KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity

  1. Marjorie Fournier  Is a corresponding author
  2. Amélie Rodrigue
  3. Larissa Milano
  4. Jean-Yves Bleuyard
  5. Anthony M Couturier
  6. Jacob Wall
  7. Jessica Ellins
  8. Svenja Hester
  9. Stephen J Smerdon
  10. László Tora
  11. Jean-Yves Masson  Is a corresponding author
  12. Fumiko Esashi  Is a corresponding author
  1. Sir William Dunn School of Pathology, University of Oxford, United Kingdom
  2. CHU de Québec Research Center, Oncology Division; Department of Molecular Biology, Medical Biochemistry and Pathology, Laval University Cancer Research Center, Canada
  3. Department of Biochemistry, University of Oxford, United Kingdom
  4. Advanced Proteomics Facility, University of Oxford, United Kingdom
  5. The Francis Crick Institute, United Kingdom
  6. Institut de Génétique et de Biologie Moléculaire et Cellulaire, France
  7. Centre National de la Recherche Scientifique, France
  8. Institut National de la Santé et de la Recherche Médicale, France
  9. Université de Strasbourg, France

Abstract

The tumour suppressor PALB2 stimulates RAD51-mediated homologous recombination (HR) repair of DNA damage, whilst its steady-state association with active genes protects these loci from replication stress. Here, we report that the lysine acetyltransferases 2A and 2B (KAT2A/2B, also called GCN5/PCAF), two well-known transcriptional regulators, acetylate a cluster of seven lysine residues (7K-patch) within the PALB2 chromatin association motif (ChAM) and, in this way, regulate context-dependent PALB2 binding to chromatin. In unperturbed cells, the 7K-patch is targeted for KAT2A/2B-mediated acetylation, which in turn enhances the direct association of PALB2 with nucleosomes. Importantly, DNA damage triggers a rapid deacetylation of ChAM and increases the overall mobility of PALB2. Distinct missense mutations of the 7K-patch render the mode of PALB2 chromatin binding, making it either unstably chromatin-bound (7Q) or randomly bound with a reduced capacity for mobilisation (7R). Significantly, both of these mutations confer a deficiency in RAD51 foci formation and increase DNA damage in S phase, leading to the reduction of overall cell survival. Thus, our study reveals that acetylation of the ChAM 7K-patch acts as a molecular switch to enable dynamic PALB2 shuttling for HR repair while protecting active genes during DNA replication.

Editor's evaluation

The manuscript provides fundamental insights into the role of acetylation of PALB2, a protein involved in Fanconi anemia and homologous recombination though its association with BRCA1 and BRCA2. The evidence that PALB2 acetylation regulates its nuclear mobility is multi-faceted and convincing and the major strength is the definition of the role of de-acetylation of the ChAM domain of PALB2 to mobilize the protein under genotoxic stress. Individuals with an interest in genome stability will be the audience for this important study.

https://doi.org/10.7554/eLife.57736.sa0

Introduction

PALB2, the partner and localizer of the breast cancer susceptibility 2 protein (BRCA2) (Xia et al., 2006), plays essential roles in the maintenance of cellular homeostasis and disease prevention in humans. Biallelic mutations in PALB2 cause Fanconi anaemia (FA), a rare genetic disorder characterised by bone marrow failure, developmental abnormalities, and an increased incidence of childhood cancers (Reid et al., 2007; Xia et al., 2007). Hereditary monoallelic PALB2 mutations also increase the risk of breast and pancreatic cancer (Jones et al., 2009; Rahman et al., 2007), similarly to inherited BRCA1 and BRCA2 mutations (ODonovan and Livingston, 2010). The physiological importance of PALB2 is further highlighted by the recent large-scale functional analysis of PALB2 mutations in cancer patients (Boonen et al., 2019; Rodrigue et al., 2019; Wiltshire et al., 2020). Canonically, PALB2 works together with BRCA1 and BRCA2 to promote error-free repair of highly genotoxic double-strand DNA breaks (DSBs) by homologous recombination (HR) (Ducy et al., 2019). In this process, BRCA1 acts as a DNA damage sensor, which in turn recruits PALB2 and BRCA2 to sites of DNA damage. Subsequently, the essential RAD51 recombinase is recruited to form nucleoprotein filaments, which catalyse the strand invasion and homology search phases of HR repair (Sy et al., 2009b; Xia et al., 2006).

Besides the role of PALB2 in promoting HR, our recent study revealed a repair-independent role of PALB2 in protecting transcriptionally active chromatin during DNA replication (Bleuyard et al., 2017b). This role of PALB2 is mediated through its high-affinity binding partner the MORF-related gene on chromosome 15 protein (MRG15), which recognises an epigenetic marker of active genes, histone H3 trimethylated at lysine 36 (H3K36me3), via its N-terminal chromodomain (Bleuyard et al., 2017b; Hayakawa et al., 2010; Sy et al., 2009a). Moreover, PALB2 intrinsic chromatin association is reinforced by its chromatin-association motif (ChAM), an evolutionarily conserved domain uniquely found in PALB2 orthologues, which directly binds to nucleosomes (Bleuyard et al., 2012). Notably, our genome-wide chromatin immunoprecipitation coupled to high-throughput sequencing (ChIP-seq) analyses revealed that PALB2 associates with a small fraction of actively transcribed genes. Notably, locus-specific analyses through ChIP followed by quantitative PCR (ChIP-qPCR) showed a decrease in PALB2 association with these genes upon exposure to an inhibitor of DNA topoisomerase I (TOP1), camptothecin (CPT), suggesting that the mode of PALB2 chromatin association is actively regulated (Bleuyard et al., 2017b). Despite these observations, the regulatory mechanism by which PALB2 switches between different modes of chromatin association (i.e. damage-induced association to promote HR repair versus steady-state association to protect active genes during DNA replication) remains unclear.

Numerous studies in recent decades have provided evidence that reversible post-translational modifications (PTMs), such as phosphorylation, ubiquitylation, SUMOylation, poly(ADP-ribosyl)ation, methylation, and acetylation, are orchestrated to promote genome stability, including the DNA damage response (DDR) (Dantuma and van Attikum, 2016). For example, the damage-responsive ATM and ATR kinases mediate phosphorylation of PALB2 at residues S59, S177, and S376, which in turn facilitates PALB2 interaction with BRCA1, RAD51 foci formation, and hence HR repair of DSBs (Ahlskog et al., 2016; Buisson et al., 2017; Guo et al., 2015). Conversely, in G1, the E3 ligase KEAP1-CUL3-RBX1 ubiquitylates PALB2 at K25, a key residue involved in BRCA1 interaction and, in this way, suppresses PALB2-BRCA1 interaction and HR activation (Orthwein et al., 2015). Furthermore, our recent work identified PALB2 as a key substrate of the lysine acetyltransferases 2A (KAT2A/GCN5) and 2B (KAT2B/PCAF) (Fournier et al., 2016), two well-known transcriptional regulators (reviewed in Nagy et al., 2010), in undamaged cells. However, the physiological role of these acetylation events is as yet unknown. Notably, KAT2A/2B use the metabolite acetyl coenzyme A (acetyl-CoA) as a cofactor (Tanner et al., 2000), and hence are proposed to fine-tune cellular processes in accordance with the metabolic status of the cell (Wellen et al., 2009). Therefore, an understanding of the functional significance of PALB2 acetylation would have important implications in the context of tumorigenesis, as cancer cells frequently exhibit reprogrammed metabolism and elevated genome instability (Fouad and Aanei, 2017).

In this study, we investigated the role of KAT2A/2B-mediated lysine acetylation in regulating PALB2. We found that KAT2A/2B acetylate a cluster of seven lysine residues (the 7K-patch) within the PALB2 ChAM. ChAM acetylation enhanced its direct association with nucleosomes. Notably, DNA damage triggered rapid ChAM deacetylation and increased the mobility of PALB2. Importantly, lysine to glutamine (Q) or arginine (R) substitutions in the 7K-patch rendered PALB2 either constitutively unbound or non-specifically chromatin-bound, resulting in impaired RAD51 foci formation in S phase and reduced cell survival. On the basis of these observations, we propose that PALB2 chromatin association is dynamically regulated by KAT2A/2B in a context-dependent manner, which plays a significant role in the maintenance of genome stability.

Materials and methods

Cell culture and cell lines

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All cells were grown at 37 °C in an incubator with a humidified atmosphere with 5% CO2. HEK293T cells were grown in Dulbecco’s Modified Eagle’s Medium (DMEM, Sigma) supplemented with 10% fetal bovine serum (FBS), 100 U/mL penicillin, and 0.1 mg/mL streptomycin. A U2OS Flp-In T-REx P2shRNA cell line (Bleuyard et al., 2017b), carrying a doxycycline-inducible shRNA targeting the endogenous PALB2 3’-UTR, referred to as U2OS-shPALB2, was used to generate stable isogenic cell lines with constitutive or inducible expression of N3xFLAG- or FLAG-EGFP-PALB2 variants, respectively. U2OS-shPALB2 cells were co-transfected with pOG44 and pcDNA5/FRT GW/N3×FLAG-PALB2 or pcDNA5/FRT/TO/FLAG-EGFP-PALB2 (7Q or 7R), and resultant stable cell lines were selected in DMEM supplemented with 10% FBS, 100 U/mL penicillin, 0.1 mg/mL streptomycin, 10 µg/mL blasticidin, 200 μg/mL hygromycin B (100 µg/mL to maintain the cell lines), and 1 µg/mL puromycin. Stable U2OS-shPALB2 lines carrying the empty pcDNA5/FRT GW/N3×FLAG vector or the pcDNA5/FRT GW/N3×FLAG-PALB2 WT vector have been described previously (Bleuyard et al., 2017b). All cell lines tested negative for Mycoplasma contamination using the MycoAlert detection kit (Lonza).

Antibodies

The primary antibodies used for western blot (WB: with their respective working dilutions) were: anti-FLAG (Sigma F1804, mouse, WB: 1/1000), anti-pan-acetyl lysine (AcK) (Cell Signaling Technology 9441 S, rabbit, WB: 1/1000), anti-PALB2 (Bethyl A301-246A, rabbit, WB: 1/500; in-house antibody raised in rabbit (Rodrigue et al., 2019), WB: 1/5000), anti-BRCA2 (Millipore OP95, mouse, WB: 1/1000), anti-RAD51 (Yata et al., 2014) (7946, rabbit, WB: 1/5000), anti-lamin A (Sigma L1293, rabbit, WB: 1/2000), anti-γ-H2A.X (Millipore 05–636, mouse, WB: 1/1000), anti-GFP (Sigma G1544, mouse, WB: 1/1000), anti-histone H3 (Bethyl A300-823A, rabbit, WB: 1/1000), anti-MRG15 (Cell Signaling Technology D2Y4J, rabbit, WB: 1/1000), anti-BRCA1 (Sigma OP107, mouse, WB: 1/1000), anti-GST (Santa Cruz Biotechnology sc-138, mouse, WB: 1/1000), biotin-HRP conjugated (Sigma A0185, mouse, WB: 1/1000), anti-KAT2A/GCN5 (Cell Signaling Technology 3305, rabbit, WB: 1/1000), anti-α-tubulin (Cell Signaling Technology 3873, mouse, WB: 1/2000) and anti-vinculin (Sigma V9131, mouse, WB: 1/200,000). Secondary antibodies coupled with horseradish peroxidase (HRP): goat anti-mouse (Dako P0447, 1/1000; Jackson ImmunoResearch 515-035-062, 1/20,000), goat anti-rabbit (Dako P0448, 1/1000; Jackson ImmunoResearch 111-035-144, 1/20,000). Antibodies used for immunofluorescence (IF) were: anti-γ-H2A.X (Millipore 05–636, mouse, IF: 1/2000) and anti-RAD51 (BioAcademia 70–001, rabbit, IF: 1/1000). Alexa Fluor conjugated secondary antibodies: goat anti-mouse (Invitrogen A-11001, IF: 1/1000; or Invitrogen A-11017, IF: 1/400) and goat anti-rabbit (Invitrogen A-11011, IF: 1/1000). For ChIP, control IgG (Jackson Immunoresearch 015-000-003, mouse) and anti-FLAG (Sigma F1804, mouse) were used.

Plasmids

For bacterial expression, full-length PALB2 and fragments 1–4 were PCR amplified using primer pairs, numbered 1 and 8 (full length), 1 and 2 (Fr. 1), 3 and 4 (Fr. 2), 5 and 6 (Fr. 3), or 7 and 8 (Fr. 4) listed in Table 1, from pCMV-SPORT6-PALB2 (IMAGE clone 6045564, Source BioSciences) and cloned into the BamHI/NotI sites of the pGEX-6P-1 vector (GE Healthcare). For mammalian expression of ChAM fragments of varying lengths, PALB2 cDNA was first PCR amplified using primer pairs, numbered 9 and 11 (#1), 9 and 12 (#2), 10 and 12 (#3), 10 and 11 (#4), or 10 and 13 (#5) listed in Table 1, cloned into the BamHI/XhoI sites of the pENTR3C Gateway entry vector (Thermo Fisher Scientific), and subsequently transferred to pcDNA-DEST53 (Invitrogen) using Gateway cloning. PALB2 Q and R missense mutations were introduced by inverse PCR, where 5’-phosphorylated oligonucleotides containing the desired mutations were used to create blunt-ended products, which were then recircularised by intramolecular ligation. For bacteria expression of ChAM missense variants, pGEX4T3-ChAM (Bleuyard et al., 2017b) was modified using primer pairs numbered 18 and 20 (7Q), 18 and 15 (3Q4K), 14 and 20 (3K4Q), or 15 and 16 (3R4K) listed in Table 1. For PALB2 7Q and 7R full-length missense variants, pENTR3C-PALB2 was modified using primer pairs numbered 18 and 19 (7Q), or 16 and 17 (7R) listed in Table 1. To generate N3xFLAG-fusion or FLAG-EGFP-fusion mammalian expression vectors, PALB2 variants were subsequently transferred to pcDNA5/FRT-GW/N3×FLAG using Gateway cloning (Bleuyard et al., 2017b), or cloned into the NotI/XhoI sites of pcDNA5/FRT/TO/FLAG-EGFP (Bleuyard et al., 2017b).

Table 1
List of oligonucleotides used in this study.
NameSequenceNo.
PALB2-F1_fo15’-atggatccatggacgagcctccc-3’1
PALB2-F1_re15’-atgcggccgcattagaacttgtgggcag-3’2
PALB2-F2_fo15’-atggatccgcacaaggcaaaaaaatg-3’3
PALB2-F2_re15’-atgcggccgctgtgatactgagaaaagac-3’4
PALB2-F3_fo15’-atggatccttatccttggatgatgatg-3’5
PALB2-F3_re15’-atgcggccgcagctttccaaagagaaac-3’6
PALB2-F4_fo15’-atggatcctgttccgtagatgtgag-3’7
PALB2-F4_re15’-atgcggccgcttatgaatagtggtatacaaat-3’8
PALB2_395_Fo5’- actggatcctcttgcacagtgcctg-3’9
PALB2_353_Fo5’- actggatccaaatctttaaaatctcccagtg-3’10
PALB2_450_Re5’- tatctcgagttaatttttacttgcatccttattttta-3’11
PALB2_433_Re5’- tatctcgagttacaaatgactctgaatgacagc-3’12
PALB2_499_Re5’- tatctcgagttacaagtcattatcttcagtggg-3’13
Patch 1-K-Rev5’-tcagagtcatttggatgtcaagaaaaaaggttt-3’14
Patch 2-WT-Fwd5’-aaaaataaaaataaggatgcaagtaaaaat-315
Patch 1-R-Rev5’-tcagagtcatttggatgtcaggagaagagggttt-3’16
Patch 2-R-Fwd_FL5’-agaaatagaaatagggatgcaagtagaaatttaaacctttccaat-3’17
Patch 1-Q-Rev5’-tcagagtcatttggatgtccagcaacaaggttt-3’18
Patch 2-Q-Fwd_FL5’-caaaatcaaaatcaggatgcaagtcaaaatttaaacctttccaat-3’19
Patch 2-Q-Fwd_ChAM5’-caaaatcaaaatcaggatgcaagtcaaaattgagcggccgcact-3’20
Beta-Actin_in3-fo5’-taacactggctcgtgtgacaa-3’21
Beta-Actin_in3-re5’-aagtgcaaagaacacggctaa-3’22
Chr5_TCOF1_peak2_fo5’-ctacccgatccctcaggtca-3’23
Chr5_TCOF1_peak2_re5’-tcagggctctatgaggggac-3’24
Chr11_WEE1_mid_fo5’-ggccgaggcttgaggtatatt-3’25
Chr11_WEE1_mid_re5’-ataaccccaaagaacacaggtca-3’26

DNA damage and drug treatment

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For ionising radiation-induced DNA damage, cells were exposed to 4 Gy γ-rays using a 137Cs source delivering a dose rate of 1.68 Gy/min (Gravatom) or a CellRad X-ray irradiator (Precision X-Ray Inc). KDAC inhibition was performed by treating cells with a cocktail of 5 mM sodium butyrate (NaB, Sigma 303410), 5 μM trichostatin (TSA, Sigma T8552) and 0.5 mM nicotinamide (NaM) for 2 hr at 37 °C. DMSO was used as negative vehicle control.

siRNA treatment

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For KAT2A/GCN5 and KAT2B/PCAF knockdowns, U2OS cells at 30% confluence were transfected using DharmaFECT 1 (Dharmacon) according to the manufacturer’s instructions, with 50 pmol each of ON-Targetplus SMARTpools siRNAs targeting KAT2A (Dharmacon L-009722-02-0005) and KAT2B (Dharmacon L-005055-00-0005) in serum-free DMEM. As a negative control, 100 pmol of ON-TARGETplus non-targeting pool siRNAs (Dharmacon D001810-10-05) were used. The serum-free medium was replaced with DMEM supplemented with 10% FBS at 24 hr after transfection, and after further 48 hr incubation, the cells were collected by trypsinisation (total of 72 hr siRNA exposure).

Fluorescence recovery after photobleaching (FRAP)

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Cells were plated into CELLview cell culture dishes (Greiner Bio-One) and analysed in phenol red-free Leibovitz’s L15 medium (Gibco). FRAP experiments were performed on a spinning-disk confocal microscope (Ultra-View Vox, Perkin Elmer) mounted on an IX81 Olympus microscope with an Olympus 60x1.4 oil PlanApo objective, in a controlled chamber at 37 °C and 5% CO2 (TOKAI HIT stage top incubator). The fluorescence signal was detected using an EMCCD camera (ImagEM, Hamamatsu C9100-13). Cells were bleached in the GFP channel at maximum laser power with a single pulse for 20 ms, within a square region of interest of 5 µm2. After bleaching, GFP fluorescence recovery was monitored within the bleached area every second for 40 s. FRAP parameters were controlled using Volocity software 6.0 (Quorum Technologies). FRAP data were fitted and normalised for overall bleaching of the entire cell (whole-cell) using the FRAP plugins in ImageJ/Fiji (https://imagej.net/mbf/intensity_vs_time_ana.htm#FRAP) (Schindelin et al., 2012). From the FRAP curve fitting, half-time recovery time values after photobleaching (t1/2) were extracted and plotted in GraphPad Prism 7.02 (GraphPad Software), in which statistical analyses were performed.

Protein purification

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FLAG-KAT2A, FLAG-KAT2A catalytic mutant, and FLAG-KAT2B proteins were purified as described previously (Fournier et al., 2016). GST-PALB2 full-length and fragments were purified from 1 L of ArcticExpress cells (Agilent Technologies), grown at 37 °C in LB broth medium containing 50 µg/mL ampicillin and 25 µg/mL gentamycin. Protein expression was induced by 0.1 mM IPTG exposure for 24 hr at 13 °C. Cells were collected by centrifugation for 15 min at 1,400 × g at 4 °C and washed with ice-cold phosphate-buffered saline (PBS). Cells lysis was performed by 30 min incubation on ice in 15 mL of ice-cold extraction buffer (50 mM Tris-HCl pH 8.0, 150 mM KCl, 1 mM EDTA, 2 mM DTT, 10% glycerol, and protease inhibitor cocktail (PIC, Sigma P2714)) supplemented with 2 mg/mL Lysozyme (Sigma) and 0.2% Triton X-100, followed by sonication. Cell lysates were collected after 45 min centrifugation at 35,000 × g, 4 °C. GST-fusion proteins were pulled down with glutathione Sepharose 4B beads (GE Healthcare), pre-washed with ice-cold PBS. After overnight incubation at 4 °C on a rotating wheel, beads were washed three times with ice-cold extraction buffer, three times with 5 mL of ice-cold ATP-Mg buffer (50 mM Tris-HCl pH 7.5, 500 mM KCl, 2 mM DTT, 20 mM MgCl2, 5 mM ATP, and 10% glycerol) to release chaperone binding from the recombinant protein PALB2 and three times with ice-cold equilibration buffer (50 mM Tris-HCl pH 8.8, 150 mM KCl, 2 mM DTT, and 10% glycerol). Proteins were eluted from beads in ice-cold elution buffer (50 mM Tris-HCl pH 8.8, 150 mM KCl, 2 mM DTT, 0.1% Triton X-100, 25 mM L-glutathione, and 10% glycerol).

For GFP-ChAM purification for mass spectrometry analysis, HEK293T cells (3×107 cells) transiently expressing GFP-ChAM were collected by centrifugation for 5 min at 500 × g, 4 °C and washed once with ice-cold PBS. Cells were further resuspended in 5 mL ice-cold sucrose buffer (10 mM Tris-HCl pH 8.0, 20 mM KCl, 250 mM sucrose, 2.5 mM MgCl2, 10 mM benzamidine hydrochloride (Benz-HCl) and PIC). After addition of Triton X-100 (Sigma) to a final concentration of 0.3% w/v, the cell suspension was vortexed four times for 10 s at 1 min intervals. The intact nuclei were collected by centrifugation for 5 min at 500 × g, 4 °C, and the supernatant was discarded. The nuclear pellet was washed once with ice-cold sucrose buffer and resuspended in ice-cold NETN250 buffer (50 mM Tris-HCl pH 8.0, 250 mM NaCl, 2 mM EDTA, 0.5% NP-40, 10 mM Benz-HCl and PIC). After 30 min incubation on ice, the chromatin fraction was collected by centrifugation for 5 min at 500 × g, 4 °C, washed once with 5 mL ice-cold NETN250 buffer and lysed for 15 min at room temperature (RT) in ice-cold NETN250 buffer supplemented 5 mM MgCl2 and 125 U/mL benzonase (Novagen 71206–3). After addition of EDTA and EGTA to respective final concentrations of 5 mM and 2 mM to inactivate the benzonase and centrifugation for 30 min at 16,100 × g, 4 °C, the supernatant was collected as the chromatin-enriched fraction. GFP-ChAM was pulled down using 15 µL GFP-Trap Agarose (Chromotek), pre-washed three times with ice-cold NETN250 buffer and blocked for 3 hr at 4 °C on a rotating wheel with 500 µL ice-cold NETN250 buffer supplemented with 2 mg/mL bovine serum albumin (BSA, Sigma). After 3 hr protein binding at 4 °C on a rotating wheel, the GFP-Trap beads were collected by centrifugation for 5 min at 1,000 × g, 4 °C and washed four times with ice-cold NETN250 buffer.

For the analysis of ChAM acetylation upon DNA damage, a GFP-ChAM fusion was transiently expressed from pDEST53-GFP-ChAM for 24 hr in HEK293T cells. Whole-cell extracts (WCEs) were prepared from ∼1.5×107 cells resuspended in NETN150 buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 2 mM EDTA and 0.5% NP-40 alternative [NP-40 hereafter] [Millipore 492018]) supplemented with 1 mM DTT, PIC, lysine deacetylase inhibitor (5 mM NaB), 1 mM MgCl2 and 125 U/mL benzonase. After 30 min incubation on ice, cell debris was removed by 30 min centrifugation at 4 °C, and the supernatant was collected as WCE. WCE was then incubated with 15 µl of GFP-Trap Agarose for GFP-pull down. After 1 hr protein binding at 4 °C on a rotating wheel, GFP-Trap beads were collected by 5 min centrifugation at 500 × g at 4 °C and washed three times with NET150 buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl and 2 mM EDTA) supplemented with 0.1% NP-40, 1 mM DTT, PIC, 5 mM NaB and 1 mM MgCl2. Proteins were eluted from beads by heating at 85 °C for 10 min in Laemmli buffer supplemented with 10 mM DTT. The proteins were separated by SDS-PAGE and analysed by western blot.

For the nucleosome pull-down assays, GFP-ChAM variants were affinity-purified from HEK293T cells (3×107 cells) following transient expression. After collecting cells by centrifugation for 5 min at 500 × g, 4 °C, the cell pellet was washed twice with ice-cold PBS and resuspended in ice-cold NETN150 buffer supplemented with 10 mM Benz-HCl and PIC. After 30 min incubation on ice, the chromatin was pelleted by centrifugation for 5 min at 500 × g, 4 °C, and the supernatant was collected as NETN150 soluble fraction and centrifuged for 30 min at 16,100 × g, 4 °C to remove cell debris and insoluble material. For each sample, 10 µL of GFP-Trap Agarose were washed three times with 500 µL ice-cold NETN150 buffer. NETN150 soluble proteins (2.5 mg) in a total volume of 1 mL ice-cold NETN150 buffer were incubated with the GFP-Trap beads to perform a GFP pull-down. After 2 hr incubation at 4 °C on a rotating wheel, the GFP-Trap beads were collected by centrifugation for 5 min at 500 × g, 4 °C and washed four times with ice-cold NETN150 buffer. Human nucleosomes were partially purified from HEK293T cells (4×107 cells), collected by centrifugation for 5 min at 500 × g, 4 °C, washed twice with ice-cold PBS and lysed in ice-cold NETN150 buffer supplemented with 10 mM Benz-HCl and PIC. After 30 min of incubation on ice, the chromatin was pelleted by centrifugation for 5 min at 500 × g, 4 °C, washed once with ice-cold NETN150 buffer and digested for 12 min at 37 °C with 50 gel units of micrococcal nuclease (NEB) per milligram of DNA in NETN150 buffer containing 5 mM CaCl2, using 200 µL buffer per mg of DNA. The reaction was stopped with 5 mM EGTA and the nucleosome suspension cleared by centrifugation for 30 min at 16,100 × g, 4 °C.

Acetyltransferase assays

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Radioactive 14C-acetyltransferase assays on recombinant proteins were performed by incubating purified GST-PALB2 (full-length and fragments) or purified RAD51 with purified FLAG-KAT2B in the presence of 14C-labeled acetyl-CoA in the reaction buffer (50 mM Tris-HCl pH 8.0, 10% glycerol, 100 mM EDTA, 50 mM KCl, 0.1 M NaB, PIC, and 5 mM DTT) for 1 hr at 30 °C. The reactions were stopped by addition of Laemmli buffer containing 10% beta-mercaptoethanol, boiled for 5 min, resolved by SDS-PAGE, and stained using Coomassie blue to reveal overall protein distribution. The acrylamide gel was then dried and exposed to phosphorimager to reveal 14C-labeled proteins. Non-radioactive acetyltransferase assays were performed as described above using cold acetyl-CoA instead. After 1 hr incubation at 30 °C, the reactions were stopped by addition of Laemmli buffer containing 10 mM DTT, boiled for 5 min, resolved by SDS-PAGE, and after Ponceau S staining of the membrane to reveal overall protein distribution, analysed by western blot using anti-acetyl lysine antibody. Acetyltransferase assays performed for mass spectrometry analyses were performed as previously described (Fournier et al., 2016).

Nucleosome pull-down assay

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Nucleosome pull-down assays were performed by mixing 250 µg of partially purified nucleosomes and GFP-ChAM variants immobilised on GFP-Trap beads in NETN150 buffer supplemented with 2 mg/mL BSA, followed by 30 min incubation at RT, then 1.5 h incubation at 4 °C, on a rotating wheel. GFP-Trap beads were further washed four times with NETN150 buffer, and samples were analysed by SDS-PAGE and western blot.

Chemical cell fractionation and whole-cell extract

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HEK293T cells transiently expressing GFP-ChAM variants were collected using TrypLE Express reagent (Gibco), washed twice with ice-cold PBS and resuspended in sucrose buffer (10 mM Tris-HCl pH 7.5, 20 mM KCl, 250 mM sucrose, 2.5 mM MgCl2, 10 mM Benz-HCl and PIC), using 1 mL buffer per 100 mg of weighed cell pellet. After addition of Triton X-100 (Sigma) to a final concentration of 0.3% w/v, the cell suspensions were vortexed three times for 10 s at 1 min intervals. The intact nuclei were collected by centrifugation for 5 min at 500 x g, 4 °C, and the supernatant collected as the cytoplasmic fraction. The nuclei pellet was washed once with ice-cold sucrose buffer and resuspended in ice-cold NETN150 buffer (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 2 mM EDTA, 0.5% NP-40, 10 mM Benz-HCl and PIC), using 400 µL buffer per 100 mg of initially weighed cell pellet. After 30 min incubation on ice, the chromatin fraction was collected by centrifugation for 5 min at 500 x g, 4 °C and the supernatant collected as nuclear soluble fraction. The chromatin pellet was washed once with ice-cold NETN150 buffer and finally solubilised for 1 hr on ice in NETN150 buffer containing 2 mM MgCl2 and 125 U/mL Benzonase nuclease (Merck Millipore), using 250 µL buffer per 100 mg of initial weighed cell pellet. Cytoplasmic, nuclear soluble and chromatin-enriched fractions were centrifuged for 30 min at 16,100 x g, 4 °C to remove cell debris and insoluble material. For whole-cell extract, cells were directly lysed in NETN150 buffer containing 2 mM MgCl2 and 125 U/mL Benzonase for 1 hr on ice and centrifuged for 30 min at 16,100 x g, 4 °C to remove cell debris and insoluble material.

Cell survival

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U2OS-shPALB2 cells complemented with FLAG-PALB2 WT or its variants were seeded in 96-wells plates and grown in the presence or absence of 2 μg/mL doxycycline for 4 days at 37 °C. Cell survival was then measured using WST-1 reagent (Roche Applied Science) following manufacturer’s protocol.

Protein structure prediction with AlphaFold2

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The predictions of full-length PALB2 (amino acid 1–1186) and the ChAM variants (amino acid 395–450) were conducted via the ColabFold: AlphaFold2 using MMseqs2 (https://colab.research.google.com/github/sokrypton/ColabFold/blob/main/AlphaFold2.ipynb) (Mirdita et al., 2022; Steinegger and Söding, 2017). The resultant structures were visualised using UCSF Chimera (https://www.cgl.ucsf.edu/chimera/) (Pettersen et al., 2004).

Immunofluorescence microscopy

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For γ-H2A.X foci analysis, cells were grown on coverslips and washed with PBS before pre-extraction with 0.1% Triton in PBS for 30 s at RT. Cells were then fixed twice with 4% PFA in PBS, first for 10 min on ice and then for 10 min at RT. After permeabilisation in 0.5% Triton X-100 in PBS for 10 min at RT, cells were blocked with 5% BSA in PBS supplemented with 0.1% Tween 20 solution (PBS-T-0.1) and incubated with anti-γ-H2A.X antibody for 3 hr at RT. After washing with PBS-T-0.1 for 5 min at RT, cells were incubated with secondary antibodies coupled with a fluorophore, washed with PBS-T-0.1 for 5 min at RT, and mounted on slides using a DAPI-containing solution. Cells were analysed on a spinning-disk confocal microscope (Ultra-View Vox, Perkin Elmer) mounted on an IX81 Olympus microscope, with a 40x1.3 oil UPlan FL objective. The fluorescence signal was detected using an EMCCD camera (ImagEM, Hamamatsu C9100-13). Images were processed in Image J (https://imagej.nih.gov/ij/) (Schneider et al., 2012).

Click-iT fluorescent EdU labelling and immunofluorescence microscopy

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For γ-H2A.X and RAD51 foci analysis, U2OS-shPALB2 stably expressing FLAG (EV) or FLAG-PALB2 variants were grown on coverslips in the presence of 2 μg/mL doxycycline for 4 or 5 days. When applicable, cells were exposed to irradiation at 4 Gy using a CellRad X-ray irradiator (Precision X-Ray Inc) and returned at 37 °C for 2 hr 30 min. Then, cells were incubated with 10 μM EdU in media for 30 min at 37 °C before washing with PBS and fixation in 4% PFA in PBS for 10 min. After permeabilisation in 0.5% Triton X-100 in PBS for 5 min, cells were blocked with 1% BSA, 10% goat serum in PBS for 30 min. EdU staining was performed with the Click-iT Alexa Fluor 647 Imaging Kit for 30 min (Invitrogen C10340), using 1/50 the recommended volume of Alexa Fluor azide, and samples were protected from light from this point on. Primary antibody incubation was performed for 1 hr with anti- γ-H2A.X (Millipore 05–636) and anti-RAD51 (BioAcademia 70–001) diluted in PBS-1% BSA at 1:2000 and 1:1000, respectively. Secondary antibodies Alexa Fluor 488 goat anti-mouse (Invitrogen A-11001) and Alexa Fluor 568 goat anti-rabbit (Invitrogen A-11011) were diluted 1:1000 in PBS-1% BSA and applied for 1 hr. Nuclei were stained for 10 min with 4, 6-diamidino-2-phenylindole (DAPI) prior to mounting on slides with ProLong Gold antifade solution (Invitrogen). All immunofluorescence steps were performed at RT with 3 intervening PBS washes. Slide images were acquired on a CellDiscoverer 7 widefield imaging system (Carl Zeiss Microscopy) using a 50 x/1.2 water immersion objective with a ×0.5 magnification changer. Acquired images were processed and analysed using Zeiss ZEN (blue edition) software.

Chromatin Immunoprecipitation

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ChIP was performed according to Bleuyard et al., 2017a with modifications. In brief, U2OS-shPALB2 stably expressing FLAG (EV) or FLAG-PALB2 variants were treated with 2 μg/mL doxycycline for 5 days and, where indicated, with 4 Gy IR before being harvested with trypsin and washed twice with PBS. For each ChIP, 2X107 cells were pelleted and resuspended in 2 ml of PBS to be then fixed for 8 min at RT with 1% formaldehyde and quenched for 5 min with 125 mM glycine. After two washes with ice-cold PBS, cells were incubated for 10 min on ice in 1 mL of lysis buffer (10 mM PIPES pH 7.5, 85 mM KCl, 0.5% NP-40, 10 mM Benz-HCl, PIC). Isolated nuclei were pelleted, resuspended in 1 mL of ChIP buffer (20 mM Tris·HCl pH 7.5, 150 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 1% Triton X-100, 0.1% sodium deoxycholate, 0.1% SDS, 10 mM Benz-HCl, PIC) and incubated for 10 min on ice. Sonication was carried out at 4 °C in 15 ml polystyrene Falcon tubes using a water-bath Bioruptor (Diagenode) on high setting for 40 cycles of 30 s on and 30 s off. Chromatin samples were centrifuged for 10 min at 16,100 × g, 4 °C and pre-cleared by adding 20 µL of protein G Dynabeads (Thermo Fisher 10003D) and incubating for 1 hr at 4 °C on a rotating wheel. From this, 800 μL chromatin was mixed with 10 μg of control mouse IgG (Jackson Immunoresearch 015-000-003) or mouse anti-FLAG antibody (Sigma F1804) in a final volume of 1 mL ChIP buffer and incubated overnight at 4 °C on a rotating wheel. A total of 100 μL of protein G Dynabeads blocked overnight with 5 mg/mL BSA in ChIP buffer was added to each sample. After 2 hr of incubation at 4 °C on a rotating wheel, beads were washed twice with low-salt wash buffer (20 mM Tris·HCl pH 8, 150 mM NaCl, 2 mM EDTA, 1% Triton X-100, 0.1% SDS, 10 mM Benz-HCl, PIC), twice with high-salt wash buffer (20 mM Tris·HCl pH 8, 500 mM NaCl, 2 mM EDTA, 1% Triton X-100, 0.1% SDS, 10 mM Benz-HCl, PIC), once with LiCl wash buffer (20 mM Tris·HCl pH 8, 250 mM LiCl, 1 mM EDTA, 1% NP-40, 1% sodium deoxycholate, 10 mM Benz-HCl, PIC), and twice with TE buffer (10 mM Tris·HCl pH 8.0, 1 mM EDTA). Beads were resuspended in 200 μL elution buffer (TE buffer, 1% SDS) and incubated for 1 hr at 65 °C, with vortexing every 15 min. After overnight cross-link reversal and proteinase K and RNase A treatment, samples were extracted with phenol/chloroform and ethanol-precipitated. ChIP-qPCR analysis was performed on a QuantStudio 7 Pro real-time PCR system (Thermo Fisher) using Applied Biosystems PowerUp SYBR Green Master Mix (Thermo Fisher). Primer sequences are listed in Table 1.

Time-lapse microscopy analysis of PALB2 recruitment to laser-induced DNA damage

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For time-lapse microscopy, U2OS-shPALB2 cells expressing FE-PALB2 WT, 7R and 7Q, and depleted of endogenous PALB2 via doxycycline exposure, were micro-irradiated using point bleach mode for 200 ms with a 405 nm UV-laser (100% output) at the following settings: format 512X512 pixels, scan speed 100 Hz, mode bidirectional, zoom 2 X, 16-bit image depth. To monitor the recruitment of FE-PALB2 to laser-induced DNA damage sites, cells were imaged every 30 s for 15 min on a Leica TCS SP5 II confocal microscope driven by Leica LAS AF software. The fluorescence intensity of FE-PALB2 at DNA damage sites relative to an unirradiated area was quantified and plotted over time.

Sequence alignment analysis

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Sequences of PALB2 orthologues from 12 species were retrieved from Ensembl (http://www.ensembl.org) and NCBI (https://www.ncbi.nlm.nih.gov) and aligned using MUSCLE (http://www.ebi.ac.uk/Tools/msa/muscle/).

Results

PALB2 is acetylated within a 7K-patch in its ChAM domain

Our previous shotgun mass spectrometry (MS) study identified a number of lysine residues within the central region of PALB2 as KAT2A/2B-dependent endogenous acetylation acceptors in HeLa cells (Fournier et al., 2016; Figure 1A, highlighted in blue). To confirm the direct acetylation of PALB2 by KAT2A/2B, we performed in vitro acetylation assays using either recombinant full-length PALB2 or a series of PALB2 fragments (Figure 1A) with purified KAT2A or KAT2B. Lysine acetylation of full-length PALB2 and fragment 2 (residues 295–610), which encompasses the DNA/chromatin binding region of PALB2, was clearly visible by western blot against acetylated lysine (pan-AcK) or 14C-autoradiography following in vitro acetylation with KAT2A or KAT2B, but not with a catalytically inactive mutant of KAT2A (KAT2A-ED) (Figure 1B, Figure 1—figure supplement 1A and B). Our MS analyses of these products identified seven acetylated lysine residues: K436, K437, K438, K441, K443, K445 and K449, in a cluster in the PALB2 chromatin association motif (ChAM), denoted the 7K-patch (Figure 1C, highlighted in orange). Notably, while sequence alignment of PALB2 orthologues showed some divergence in the C-terminal region of ChAM, an enrichment of lysine residues was consistently detected (Bleuyard et al., 2012; Figure 1D). Clusters of lysine residues were also frequently found in other KAT2A/2B substrates (Fournier et al., 2016), corroborating our finding that the ChAM C-terminal region is favourably targeted for KAT2A/2B-mediated acetylation.

Figure 1 with 1 supplement see all
KAT2A/2B acetylate PALB2 within a 7K-patch in its ChAM domain.

(A) PALB2 lysine residues identified as acceptors of KAT2A/2B-dependent acetylation in vivo and in vitro by tandem MS analyses are shown in blue and black, respectively. An asterisk indicates residues that are only detected in endogenous PALB2, but not in PALB2 acetylated in vitro. The acetylated lysine residues within the ChAM are highlighted in red. Full-length PALB2 (FL,~131,3 kDa) and fragments 1–4 used for in vitro acetylation assays are also shown. 1: PALB2 1–320 (36.2 kDa), 2: 295–610 (35.6 kDa), 3: 580–900 (35.3 kDa) and 4: 867–1186 (35.3 kDa). (B) In vitro acetylation of PALB2 by KAT2A and catalytically inactive KAT2A-ED. Purified GST-fusions of PALB2 full-length (158.1 kDa) and fragments 1 (63 kDa), 2 (62.4 kDa), 3 (62.1 kDa), and 4 (62.1 kDa) (depicted in A) were incubated with either purified Flag-KAT2A or Flag-KAT2A-ED in the presence of acetyl-CoA, followed by SDS-PAGE. Total and acetylated proteins were visualized by Ponceau S and anti-acetyl lysine (pan-AcK) western blot, respectively. PALB2 fragment 2 acetylation by KAT2A is highlighted with a red dashed box. (C) A heat map of acetylated lysine residues, as identified by quantitative MS analysis of in vitro acetylated PALB2 FL or fragment 2 by KAT2B, KAT2A or a catalytically inactive KAT2A (KAT2A-ED). The abundance of each acetyl-lysine was evaluated as previously described (Fournier et al., 2016). The lysine residues which were detected as endogenous acetylation acceptor are shown to the left. (D) ChAM protein sequences from twelve PALB2 orthologues were aligned using MUSCLE (EMBL-EBI sequence analysis tool) and visualised using the ClustalW software with default colour-coding. Lysine residues acetylated by KAT2A and KAT2B in vitro are respectively highlighted by red and green dots. Hs (Homo sapiens, human), Pt (Pan troglodytes, chimpanzee), Pp (Pan paniscus, bonobo), Bt (Bos taurus, cow), Ml (Myotis lucifugus, little brown bat), Ss (Sus scrofa, wild boar), Ec (Equus caballus, horse), Cl (Canis lupus familiaris, dog), Am (Ailuropoda melanoleuca, giant panda), Oa (Ovis aries, sheep), Gg (Gallus gallus, red junglefowl), Tg (Taeniopygia guttata, zebra finch).

The 7K-patch promotes ChAM nucleosome association

To evaluate the impact of the 7K-patch on ChAM chromatin association, we generated a series of GFP-tagged ChAM truncations (Figure 2A). Critically, when expressed in HEK293T cells, the ChAM fragment (PALB2 residues 395–450) was found enriched in the chromatin fraction in highly acetylated forms (Figure 2A and B, Figure 2—figure supplement 1A and B). Further analysis of ChAM truncations revealed that the region containing the 7K-patch was essential for its chromatin association (Figure 2A and B). Considering the possibility that the deletion of the 7K-patch might lead to aberrant subcellular localisation and contribute to the reduced chromatin enrichment, we also affinity purified the corresponding fragments from the cytoplasmic fraction of transfected HEK293T cells and assessed their interaction with separately prepared nucleosomes (Figure 2—figure supplement 1A). The pull-down assay showed that deletion of the 7K-patch impaired ChAM interaction with nucleosomes (Figure 2A and C), corroborating our observations above.

Figure 2 with 2 supplements see all
The 7K-patch promotes ChAM nucleosome association.

(A) Diagram showing the full-length (FL) PALB2, GFP-ChAM fragments #1 to #5 and GST-ChAM variants used in this study. ChAM C-terminal lysine-rich patch (7K-patch, residues 436–449) is highlighted in red. (B) Western blot analysis assessing the chromatin enrichment of GFP-ChAM fragments transiently expressed in HEK293T cells. Empty GFP vector (EV) was used as a negative control. Whole-cell extract was also prepared to compare GFP-ChAM fragments expression levels. Lamin A and histone H3 were used as controls for the cellular fractionation. (C) In vitro nucleosome binding assay using GFP and GFP-ChAM fragments. Partially purified human nucleosomes captured by GFP and GFP-ChAM fragments immobilised on beads were detected by anti-histone H3 western blot and Ponceau S staining. (D) Coomassie blue staining of GST and GST-ChAM 7R and 7Q variants purified from E. coli and separated by SDS-PAGE. (E) In vitro binding assays using immobilised GST-ChAM 7R and 7Q variants and purified HeLa poly-nucleosomes. GST was used as a negative control. The nucleosome binding efficiency was assessed by anti-histone H3 western blot. (F) Coomassie blue staining of GST and GST-ChAM 3Q4K, 3K4Q and 7Q variants purified from E. coli and separated by SDS-PAGE. (G) In vitro binding assays using immobilised GST-ChAM 3Q4K, 3K4Q and 7Q variants and purified HeLa mononucleosomes. GST was used as a negative control. The nucleosome binding efficiency was assessed by anti-histone H3 western blot.

To further assess the direct role of 7K-patch acetylation in nucleosome binding, we produced recombinant GST fusions of ChAM and its variants harbouring glutamine (Q) or arginine (R) substitutions at all seven lysine residues (7Q or 7R, respectively), three conserved lysine residues, K436, K437, and K438 (3Q4K), or the four remaining lysine residues, K441, K443, K445, and K449 (3K4Q) within the 7K-patch (Figure 2A). K to Q substitutions nullify the positive charge of the lysine side chains and are therefore commonly considered to mimic acetyl-lysine, albeit with a reduction in the side chain size (Figure 2—figure supplement 2A). Conversely, K to R substitution maintains a positive charge but is unable to accept acetylation and, hence, is widely considered to mimic constitutively non-acetyl-lysine, albeit with an increase in the side chain size (Figure 2—figure supplement 2A). Our in silico AlphaFold2 modelling suggest that ChAM has a propensity to form an alpha-helical structures with a less defined C-terminal section, potentially allowing it to explore a spectrum of structural configurations for optimum nucleosome binding (Figure 2—figure supplement 2B). On the contrary, 7Q or 7R substitutions make this section more defined, potentially constraining its capacity to bind nucleosomes in optimal configurations (Figure 2—figure supplement 2C and D).

Significantly, in vitro nucleosome pull-down assays showed that, while the 7R variant largely maintains its interaction with nucleosomes (Figure 2D and E), all variants with Q substitutions abolished this nucleosome binding property (Figure 2F and G). This observation appeared to be at odds with our observation that ChAM acetylation was most notable on the chromatin-enriched fraction (Figure 2—figure supplement 1A and B). Considering that the size of the glutamine side chain is significantly smaller than the corresponding acetyl-lysine side chain (Figure 2—figure supplement 2A and D), we reasoned it highly unlikely that the K to Q substitutions accurately mimic the properties of acetylated ChAM. Collectively, our experimental results showed that K to Q substitutions within the 7K-patch fully perturbed ChAM ability to bind nucleosomes; the 7Q variant is hereafter considered to behave as a ChAM null mutant, rather than an acetyl-mimetic.

Acetylation within the 7K-patch enhances ChAM nucleosome association

KAT2A/2B associate with chromatin and facilitate transcription by acetylating histones (Nagy et al., 2010). Concordantly, the level of acetylation in the GFP-ChAM fragment affinity purified from the chromatin-enriched fraction was notably higher than that from the cytoplasmic or nuclear soluble fraction (Figure 2—figure supplement 1A and B). Our MS analysis of the chromatin-associated GFP-ChAM fragment detected acetylation on all seven lysine residues of the 7K-patch (Figure 3A, marked with arrows). To accurately assess the impact of the ChAM acetylation, synthetic peptides corresponding to the minimal ChAM (PALB2 residues 395–450) and containing acetyl-lysine at the evolutionarily conserved K436, K437, and/or K438 were generated, and their biochemical properties were characterised. Markedly, nucleosome pull-down assays revealed clear nucleosome binding by acetylated ChAM peptides, but not by their non-acetylated counterpart, where salmon sperm DNA was provided in excess to outcompete ChAM electrostatic interaction with DNA (Figure 3B). This effect was readily detectable with the ChAM peptide containing a mono-acetylation at K438, which was detected in our original MS study of the endogenous HeLa KAT2A/2B-acetylome (Fournier et al., 2016; Figures 1A, C, D ,, 3A, highlighted in blue). Together, these results demonstrate that acetylation within its 7K-patch enhances ChAM nucleosome binding.

Figure 3 with 5 supplements see all
Acetylation within the 7K-patch enhances ChAM nucleosome association.

(A) In vivo acetylated lysine residues in chromatin-associated ChAM, detected by tandem MS analysis, are highlighted by arrows in the upper part (MS). Synthetic biotin-ChAM peptides non-acetylated (non-ac) or acetylated at single lysine residues 436 (436-ac), 437 (437-ac), and 438 (438-ac), or all three lysine residues (All 3-ac) are shown in the lower part. The position of the 7K-patch is highlighted in red. The position of the K438 residue, the endogenous target of KAT2A/2B within the ChAM (shown in Figure 1A), is highlighted in a box. (B) In vitro nucleosome binding assays for the synthetic biotin-ChAM peptides. After incubation with purified HeLa polynucleosomes in the presence of 5 µg salmon sperm DNA, biotin-ChAM peptides were pulled-down using streptavidin beads (Strep pulldown), and associated nucleosomes were detected by anti-histone H3 western blot. (C-D) FRAP assay of FE-PALB2 expressed in U2OS-shPALB2 cells, treated either with siRNA targeting KAT2A and KAT2B (siKAT2A/siKAT2B) or with negative control siRNA (siCntl). Representative images of live cells before bleaching (pre-bleaching) and during a 37.5 s recovery period (post-bleaching) are shown in (C), where dashed circles indicate bleached areas. FRAP data are quantified and plotted in (D). Dots represent values of half-recovery time (t1/2) for individual cells and bars mean values ± SD (siCntl, n=59; siKAT2A/2B, n=51). Statistical analyses were performed using GraphPad Prism 7.02 and p-values are for the unpaired Student’s t-test (*** p<0.0006). (E-F) As in C-D, except cells were treated with a cocktail of lysine deacetylase inhibitors (KDACi, 5 mM sodium butyrate, 5 µM trichostatin A, and 5 mM nicotinamide) or DMSO as a control. Representative images of live cells before bleaching (pre-bleaching) and during a 37.5 s recovery period (post-bleaching) are shown in (E), where dashed circles indicate bleached areas. FRAP data are quantified and plotted in (F). Dots represent values of half-recovery time (t1/2) for individual cells and bars mean values ± SD (DMSO, n=61; KDAC, n=94). Statistical analyses were performed using GraphPad Prism 7.02 and p-values are for the unpaired Student’s t-test (**** p<0.0001). Representative real-time cell images are shown in Figure 3—video 1 (siCntl), Figure 3—video 1 (siKAT2A/siKAT2B), Figure 3—video 1 (DMSO) and Figure 3—video 4 (KDACi).

PALB2 mobility increases upon deacetylation

Next, we set out to investigate whether lysine acetylation might facilitate PALB2 chromatin association in a cellular context. To evaluate the impact of native PALB2 acetylation, we down-regulated KAT2A/2B activity using siRNA and assessed PALB2 mobility as a surrogate measure of its chromatin association. To this end, a tandem FLAG- and EGFP-tagged full-length wild-type (WT) PALB2 (FE-PALB2) was conditionally expressed in a U2OS cell line in which endogenous PALB2 was down-regulated by a doxycycline-inducible short hairpin RNA (U2OS-shPALB2) (Figure 3—figure supplement 1A), and the mobility of FE-PALB2 was assessed using fluorescence recovery after photobleaching (FRAP). This analysis revealed that KAT2A/2B siRNA treatment led to an increase in FE-PALB2 diffusion rate (reduced FRAP t1/2; Figure 3C and D, Figure 3—figure supplement 1B and C, Figure 3—videos 1; 2), concomitant with reduced levels of global acetylation (Fournier et al., 2016). Conversely, the inhibition of lysine deacetylases (KDAC), which increased the global lysine acetylation levels, decreased the diffusion rate of FE-PALB2 (increased FRAP t1/2; Figure 3E and F, Figure 3—figure supplement 1D and E, Figure 3—videos 3 and 4). It is noteworthy that hyper-acetylation of histones promotes chromatin relaxation, which can increase the local mobility of chromatin. Hence, the reduced PALB2 mobility in cells treated with KDAC inhibitors suggests that PALB2 chromatin association is stimulated by lysine acetylation.

DNA damage triggers ChAM deacetylation and increases PALB2 mobility

Our observations above indicate that PALB2 acetylation promotes its association with nucleosomes/chromatin, thereby restricting its mobility (Figure 4A). However, static PALB2 chromatin association could be toxic to cells when DSBs at random locations need to be recognised by the repair complex. Hence, we hypothesised that ChAM acetylation might be dynamically controlled upon stochastic DNA damage, such that PALB2 can be mobilised. Indeed, in HEK293 cells exposed to ionizing radiation (IR), rapid deacetylation of the ChAM fragment was detected at 15 min and persisted for at least 2 hr (Figure 4B and C). Furthermore, FRAP analyses of full-length FE-PALB2 in U2OS cells revealed a clear increase in the diffusion rate following IR treatment, decreasing FRAP t1/2 by 22% compared to control conditions (Figure 4D and E, Figure 4—figure supplement 1A-C, Figure 4—videos 1 and 2). In both HEK293 and U2OS cells, IR-induced DNA damage response was confirmed by γ-H2A.X staining (Figure 4B, Figure 4—figure supplement 1B). Since PALB2 mobility is suppressed when lysine acetylation is up-regulated (Figure 3E and F), it is unlikely that the damage-induced increase of PALB2 mobility is associated with chromatin relaxation caused by histone acetylation, an event which is observed upon DNA damage (Ziv et al., 2006). Together, these results support the notion that DNA damage triggers ChAM deacetylation and PALB2 mobilisation.

Figure 4 with 3 supplements see all
DNA damage triggers ChAM deacetylation and PALB2 mobilization.

(A) Depiction of PALB2 acetylation and nucleosome binding, controlled by KAT2A/2B and KDACs. (B) HEK293T cells transiently expressing GFP-ChAM were treated with 4 Gy IR, and affinity-purified GFP-ChAM acetylation was assessed using anti-acetyl-lysine (pan-AcK) western blot. γ-H2A.X signal in whole cell lysate was detected to monitor DNA damage. (C) Histogram showing the relative level of ChAM acetylation following irradiation. The levels of acetylated and total GFP-ChAM were quantified by ImageJ, and the ratio of acetyl/total GFP-ChAM was normalised against that of the pre-irradiation sample (time 0). Error bars indicate SD from two independent experiments (n=2). Statistical analyses were performed using GraphPad Prism 9.4.1 and p-value is the unpaired one-way ANOVA (**p<0.01). (D-E) FRAP assay of FE-PALB2 expressed in U2OS-shPALB2 cells, either non-treated (NT) or treated with 4 Gy IR. Representative images of live cells before bleaching (pre-bleaching) and during a 37.5 s recovery period (post-bleaching) are shown in (D), where dashed circles indicate bleached areas. FRAP data are quantified and plotted in (E). Dots represent values of half-recovery time (t1/2) for individual cells and bars mean values ± SD (NT, n=20; IR, n=21). Statistical analyses were performed using GraphPad Prism 7.02 and p-values are for the unpaired Student’s t-test (**** p<0.0001). Representative real-time cell images are shown in Figure 4—video 1 (NT) and Figure 4—video 2 (IR).

PALB2 ChAM 7K-patch acetylation defines its context-dependent association with actively transcribed genes

Exposure to IR stochastically induces various DNA lesions, including DSBs, single-strand breaks (SSBs), base damage and DNA-protein cross-links, at random genomic locations. In S phase, these lesions perturb DNA replication and, in some circumstances, could be converted to DSBs. Given that IR triggers ChAM deacetylation and PALB2 mobilisation, we considered that PALB2 ChAM deacetylation might promote translocation of PALB2 from its intrinsically associated actively transcribed loci to randomly distributed sites of DSBs to promote HR repair.

To address this idea, we made use of full-length PALB2 variants in which lysine residues within the 7K-patch were substituted with Q (7K-patch null mutant) or R (non-acetylatable positively charged 7K-patch). These PALB2 variants, named 7Q or 7R, as well as PALB2 WT or an empty vector (EV), were stably expressed as FLAG-fusions at equivalent levels in U2OS-shPALB2 cells (Figure 5A, Figure 5—figure supplement 1A). As expected, our subcellular fractionation analyses showed a reduced level of PALB2 7Q in the chromatin-enriched fraction compared to PALB2 WT or 7R (Figure 5B and C), agreeing with the direct nucleosome binding properties of the ChAM variants (Figure 2D and E). Similarly, our FRAP analyses of the PALB2 variants, conditionally expressed as FE-fusions in U2OS-shPALB2 cells, showed an increase in PALB2 7Q diffusion kinetics compared to those of PALB2 WT and 7R (Figure 5D and E, Figure 5—videos 1–3). A PALB2 variant with an internal deletion of ChAM, which abolishes its chromatin association (Bleuyard et al., 2012), equally exhibited a significant increase in PALB2 diffusion rate (Figure 5—figure supplement 1B-D, Figure 5—videos 4; 5). Collectively, we concluded that PALB2 7Q is constitutively highly mobile and is unable to engage with chromatin.

Figure 5 with 6 supplements see all
ChAM 7K-patch mediates PALB2 global chromatin association.

(A) Western blot analysis of U2OS-shPALB2 cells constitutively expressing FLAG (EV) or FLAG-PALB2 variants. Where indicated, cells were treated with 2 µg/mL doxycycline (+Dox) to deplete endogenous PALB2. Lamin A was used as a loading control. (B-C) Subcellular distribution of FLAG-PALB2 variants in cytoplasmic (Cyt), nuclear soluble (Nuc) and chromatin-enriched (Chr) fractions upon doxycycline-induced PALB2 depletion. Histone H3 was used as a control for cellular fractionation. Protein levels, detected by western blotting, were quantified using ImageJ. PALB2 levels in the chromatin fraction were normalised against PALB2 levels in whole-cell extracts shown in (A) and expressed as a percentage of the WT in (C). Data indicate mean values ± SD from three independent experiments (n=3). Statistical analyses were performed using GraphPad Prism 7.02, and p-values are from the unpaired Student’s t-test for pairwise comparisons (ns, not significant; **** p<0.0001). (D-E) FRAP analysis of FE-PALB2 WT, 7R and 7Q in U2OS-shPALB2 cells in which endogenous PALB2 was depleted via doxycycline exposure. Representative images before bleaching (pre-bleaching) and during the recovery period (post-bleaching) are shown in (D), where dashed circles indicate bleached areas. FRAP data are quantified and plotted in (E). Dots represent values of half-recovery time (t1/2) for individual cells and bars mean values ± SD (WT, n=33; 7R, n=31; 7Q, n=26). Statistical analyses were performed using GraphPad Prism 7.02 and p-values are from the unpaired Student’s t-test for pairwise comparisons (ns, not significant; **** p<0.0001). Representative real-time cell images are shown in Figure 5—video 1 (WT), Figure 5—video 2 (7R) and Figure 5—video 3 (7Q).

PALB2 7K-patch is important for damage-induced mobility change

We next turned to investigate the impact of the mutations of the 7K-patch on PALB2 recruitment to randomly distributed DNA damage sites. Here, we exposed FE-PALB2-expressing U2OS-shPALB2 cells to doxycycline (Dox) for 4 days (Figure 6A and B) for fuller depletion of endogenous PALB2, which might otherwise mask the phenotypes of the 7Q or 7R mutants by forming heterodimers (Buisson and Masson, 2012). Localised DNA damage was subsequently induced by laser micro-irradiation and the recruitment of FE-PALB2 variants was assessed by live cell imaging. This analysis revealed that the 7R substitution, but not the 7Q counterpart, impeded efficient recruitment of PALB2 to the micro-irradiated chromatin areas (Figure 6C and D). These impacts are unlikely to be associated with defective PALB2 protein complex formation, as all PALB2 variants maintained equivalent levels of interaction with BRCA1, BRCA2 and RAD51 in cells treated with IR (Figure 6—figure supplement 1). Rather, this observation is best explained if constitutively non-acetylated ChAM 7K-patch, as mimicked by the 7R substitutions, elicits sparse and non-specific chromatin association, preventing active displacement of PALB2 from these regions upon DNA damage. Hence, we consider that PALB2 7R is a variant which randomly binds chromatin with no locus specificity and is less efficiently mobilised upon DNA damage.

Figure 6 with 1 supplement see all
ChAM 7K-patch affect PALB2 localisation upon micro-irradiation.

(A) Depletion of endogenous PALB2 in the U2OS-shPALB2 cell line following the exposure to doxycycline over 4 days. Vinculin was used as a loading control. (B) Diagrams of experimental procedures, assessing the recruitment of PALB2 upon micro-irradiation following depletion of endogenous PALB2 with 4 days of doxycycline exposure. (C) Time plots of fluorescence intensity of FE-PALB2 at laser damage sites relative to an unirradiated area. Data represent the mean of relative fluorescence intensity ± SEM from four independent experiments (a total of at least n=100 cells per condition). Statistical analysis was done for the last time-point using GraphPad Prism 8 and p-values are for the Mann–Whitney test (ns, not significant; ***p<0.001). (D) Representative images of FE-PALB2 variants conditionally expressed in U2OS-shPALB2 cells upon micro-irradiation. Dashed circles indicate irradiated areas.

PALB2 7K-patch is important for efficient recruitment of RAD51 at randomly distributed DNA damage sites

Having established the constitutively highly mobile (7Q) or weakly mobile (7R) PALB2 variants, we further evaluated the impact of PALB2 mobility on the proficiency of HR by assessing the formation of RAD51 foci in S phase cells upon IR exposure. For these experiments, cells were exposed to doxycycline for 4 or 5 days (Figure 7A) to further ensure the full depletion of endogenous PALB2. Cells depleted of endogenous PALB2 for 4 or 5 days readily exhibited impaired IR-induced RAD51 foci formation in S phase, marked by the incorporation of thymidine analogue EdU. This phenotype was efficiently rescued by PALB2 WT complementation, but only partially by PALB2 7R or 7Q (Figure 7B). Additionally, while we observed no significant increase of γ−H2A.X in these cells at 4 days after doxycycline exposure (Figure 7C), more noticeable increases of γ−H2A.X were detected in PALB2 7Q or 7R cells exposed to doxycycline for 5 days compared to WT cells (Figure 7D–F). Given that RAD51 foci and γ−H2A.X foci formation closely reflect HR proficiency and DNA damage, respectively, this finding suggested that both the 7Q and 7R substitutions confer HR defects and accumulation of DNA damage in the longer term.

7K-patch-mediated PALB2 chromatin association affects IR-induced RAD51 foci formation.

(A) Diagrams of experimental procedures, assessing RAD51 in S phase cells, marked by EdU pulse labelling. (B-C) Quantification of the number of RAD51 foci (B) or γ-H2A.X foci (C) per cell, upon doxycycline exposure for 4 days and subsequently exposed to 4 Gy IR. Dots represent values for individual cells and bars mean values from three independent experiments (a total of n=600 cells per condition). Statistical analyses were performed using GraphPad Prism 8 and p-values are for the Mann–Whitney test (ns, not significant; **p<0.01; ****p<0.0001). (D-E) Quantification of the number of RAD51 foci (D) or γ-H2A.X foci (E) per cell, upon doxycycline exposure for 5 days. Dots represent values for individual cells and bars mean values from three independent experiments (a total of n=600 cells per condition). Statistical analyses were performed using GraphPad Prism 8 and p-values are for the Mann–Whitney test (****p<0.0001). (F) Representative images of RAD51 and γ-H2A.X foci in U2OS-shPALB2 cells stably expressing FLAG (EV) and FLAG-PALB2 variants, following doxycycline-induced endogenous PALB2 depletion for 5 days and subsequently exposed to 4 Gy IR. DAPI was used for nuclear staining.

The PALB2 7K-patch is important for normal cell growth

While conducting the above experiment, we noticed that, similarly to IR treated conditions, PALB2-depleted, but otherwise untreated, cells expressing 7Q or 7R exhibited impaired spontaneous RAD51 foci and an increase of γ−H2A.X in S phase (Figure 8A–F). The physiological importance of this phenomenon is highlighted by the reduced survival of PALB2-depleted cells complemented with 7Q PALB2 and, more significantly, with 7R PALB2, compared to those expressing WT PALB2 in untreated cells (Figure 9A and B).

7K-patch-mediated PALB2 chromatin association affects spontaneous RAD51 foci formation in normally growing cells.

(A) Diagrams of experimental procedures, assessing RAD51 in S phase cells, marked by EdU pulse labelling. (B-C) Quantification of the number of RAD51 foci (B) or γ-H2A.X foci (C) per cell, upon doxycycline exposure for 4 days. Dots represent values for individual cells and bars mean values from three independent experiments (a total of n=600 cells per condition). Statistical analyses were performed using GraphPad Prism 8 and p-values are for the Mann–Whitney test (ns, not significant; **p<0.01; ****p<0.0001) (D-E) Quantification of the number of RAD51 foci (E) or γ-H2A.X foci (F) per cell, upon doxycycline exposure for 5 days. Dots represent values for individual cells and bars mean values from three independent experiments (a total of n=600 cells per condition). Statistical analyses were performed using GraphPad Prism 8 and p-values are for the Mann–Whitney test (****P<0.0001). (F) Representative images of RAD51 and γ-H2A.X foci in U2OS-shPALB2 cells stably expressing FLAG (EV) and FLAG-PALB2 variants, following doxycycline-induced endogenous PALB2 depletion for 5 days. DAPI was used for nuclear staining.

Figure 9 with 1 supplement see all
7K-patch is important for normal cell survival.

(A) Diagrams of experimental procedures, assessing survival of U2OS-shPALB2 stably expressing FLAG (EV) or FLAG-PALB2 variants upon doxycycline (Dox)-induced endogenous PALB2 depletion by WST-1 assay. (B) Cellular survival normalised against those without doxycycline exposure. Data represent mean values ± SD from three independent experiments (n=3). Statistical analyses were performed using GraphPad Prism 9 and p-values are the unpaired one-way ANOVA (*p<0.05; **p<0.01; ***p<0.001). (C) Diagrams of experimental procedures, assessing the FLAG-PALB2 fusion enrichment at previously reported PALB2-bound loci by ChIP-qPCR. (D) ChIP-qPCR analysis of FLAG-PALB2 WT, 7Q and 7R enrichment at known PALB2-bound loci (Bleuyard et al., 2017a), namely the coding regions of the ACTB, TCOF1, and WEE1 genes, in cells untreated or treated with 4 Gy IR. Data are expressed as fold enrichment over IgG and represent mean values ± SD from three independent experiments (n=3) with triplicate qPCR reactions. Statistical analyses were performed using GraphPad Prism 8 and p-values are from two-way ANOVA with Tukey’s multiple comparisons test (*p<0.05; **p<0.01). (E) Model for PALB2 acetylation function in the maintenance of genome stability. MRG15 and KAT2A/2B-mediated ChAM acetylation, which occurs locally at active genes, jointly promote PALB2 enrichment at undamaged transcriptionally active chromatin. DNA damage triggers ChAM deacetylation at these loci and temporarily releases PALB2 from chromatin. This allows PALB2 to interact with damage sensors, such as BRCA1, which in turn recruits the entire HR complex to sites of DNA damage. ChAM binding to naked DNA through the deacetylated 7K-patch may promote RAD51 loading and HR repair. The constitutively highly mobile PALB2 variant (7Q) is efficiently recruited to sites of DNA damage but, unlike WT PALB2, unable to promote efficient RAD51 foci formation. The randomly chromatin-bound and less capably mobilised PALB2 variant (7R) is inefficiently recruited to sites of DNA damage and hence fails to promote HR.

Intrigued by these observations, we sought the underlying molecular mechanism. It is widely described that transcriptionally active loci are vulnerable to DNA damage during DNA replication, particularly when challenged by genotoxic reagents that stabilise RNA:DNA hybrids, such as camptothecin (CPT) (Marinello et al., 2013). In undamaged cells, WT PALB2 is enriched at a small subset of, but not all, H3K36me3 marked active genes, which in turn prevents CPT-induced DNA damage at these loci (Bleuyard et al., 2017a). We therefore asked whether the PALB2 7K-patch affects its occupancy at these previously identified PALB2-bound loci. Indeed, our ChIP-qPCR analyses revealed significant impairments of PALB2 7R and 7Q occupancies at these loci, namely the ACTB, TCOF1 and WEE1 genes (Figure 9C and D). These observations are in line with the view that the 7Q substitutions impede overall chromatin association, while 7R substitutions elicit sparse and non-specific chromatin association, limiting PALB2 enrichment at these loci. It is important to note, however, that reduced PALB2 association at these loci per se does not confer DNA damage in cis unless cells are challenged by CPT (Bleuyard et al., 2017a). Hence, this did not fully explain the increase of γ−H2A.X in cells expressing the 7Q or 7R variant in unperturbed conditions as shown in Figure 8E.

We postulated that, even in unperturbed cells, spontaneous DNA lesions may be induced at any locations of genomic DNA due to naturally occurring genotoxic metabolites, such as reactive oxygen species, and be processed to DSBs when encountered by the DNA replication machinery. In this scenario, the mobility of PALB2 would be important for the repair of these randomly distributed DSBs, as it is following exposure to IR. Indeed, our ChIP-qPCR assessment of PALB2 occupancy at the known PALB2-bound loci revealed that the occupancy of PALB2 WT at active genes was dramatically reduced upon IR exposure to levels equivalent to those of the untreated 7Q or 7R mutant (Figure 9D). Together, these observations indicate that the 7Q or 7R variants are defective for HR repair of randomly induced DNA damage outside of active genes.

Discussion

In this study, we have demonstrated that reversible lysine acetylation controls the mode of PALB2 chromatin association, fine-tuning RAD51 recruitment during S phase. Taking all our findings collectively, we propose the following model (illustrated in Figure 9E): (1) KAT2A/2B-mediated ChAM acetylation, jointly with MRG15, governs PALB2 enrichment at undamaged, transcriptionally active chromatin; (2) global DNA damage signalling triggers ChAM deacetylation, which in turn releases PALB2 from active genes; (3) in this way, PALB2, in complex with BRCA1, BRCA2 and RAD51, is effectively recruited to sites of damaged chromatin; and (4) deacetylated ChAM binding to exposed damaged chromatin allows appropriate engagement of this complex with damaged DNA and hence promotes RAD51 loading and HR repair.

These findings further our understanding of how the ChAM domain interacts with nucleosomes and how this association is dynamically regulated. It was previously demonstrated that the evolutionarily highly conserved N-terminal part of the ChAM affects its association with nucleosomes (Bleuyard et al., 2017a; Bleuyard et al., 2017b), while an interface composed of basic residues across the ChAM binds to an acidic surface on histone H2A in the nucleosome core particle (Belotserkovskaya et al., 2020). Our results presented here introduce an additional complexity, showing that the C-terminal 7K basic patch is essential for context-dependent ChAM interaction with nucleosomes. Specifically, ChAM binding to nucleosomes at active genes is promoted by 7K patch acetylation, while its dynamic recruitment to damaged chromatin is stimulated by 7K patch deacetylation (Figures 3 and 9C and D). Considering negative charge of nucleosomal DNA, it is not surprising that lysine acetylation, which neutralises its positive charge, might impact ChAM interaction with nucleosomes. However, our results have also revealed that the acetylation of the 7K patch provides a non-electrostatic and as-yet incompletely understood mechanism to enhance its association with nucleosomes. Intriguingly, the flanking regions of the ChAM also appear to influence ChAM binding to nucleosomes (Figure 2A, B and C): an N-terminal extension enhanced ChAM interaction with nucleosomes, while a C-terminal extension had a negative impact. These regions are rich in serine and threonine residues, including potential targets for cell cycle regulators (CDKs and PLK1) and DNA damage-responsive kinases (ATM and ATR) (Figure 9—figure supplement 1). These observations suggest that additional PTMs other than lysine acetylation might be involved in regulating ChAM association with nucleosomes. These would undoubtedly merit further investigation.

The increase in PALB2 mobility during the DDR shown in this study (Figure 4D and E) is reminiscent of previous reports demonstrating increased RAD51 mobility upon replication stress (Srivastava et al., 2009; Yu et al., 2003). Yu et al. showed that a fraction of RAD51, which is in complex with BRCA2, is immobile in unchallenged cells, but becomes mobile upon hydroxyurea-induced replicative stress (Yu et al., 2003). Notably, their data suggested that such increased RAD51 mobility was unlikely to be a consequence of altered interaction with BRCA2 upon replicative stress. Similarly, Jeyasekharan et al. reported increased BRCA2 mobility upon IR-induced DNA damage, which was dependent on the DNA damage signalling kinases ATM and ATR and coincided with active HR events (Jeyasekharan et al., 2010). Given that BRCA2 and PALB2 form a stable complex together with a fraction of RAD51 and that their interaction is important for HR repair, we suggest that the BRCA2-PALB2 complex and associated RAD51 are mobilised together upon genotoxic stress.

The mechanisms finely regulating the mobilisation and localisation of HR factors are not fully elucidated but undoubtedly involve PTMs. We propose that dynamic modes of PTM-mediated chromatin association synergistically govern the recruitment of repair factors to defined regions of chromatin. In undamaged cells, PALB2 is enriched at actively transcribed genes through KAT2A/2B-mediated ChAM acetylation and H3K36me3-mediated MRG15 interaction, while maintaining transient chromatin interaction. This transient mode of association may allow a severance mechanism, in which PALB2 continuously monitors the presence of DNA damage across these regions. Indeed, cells expressing PALB2 7R, which is capable of bulk chromatin association (Figure 5) but fails to bind specific active genes (Figure 9D), exhibited an increase in the basal level of γ−H2A.X compared to those expressing PALB2 WT (Figure 8F) and reduced survival (Figure 9B), underlining the importance of PALB2 acetylation in unperturbed cells. Upon DNA damage, however, ChAM is deacetylated (Figure 4B and C) and, as a consequence, PALB2 affinity for undamaged chromatin is reduced (Figure 9D). We suggest that damage-induced ChAM deacetylation allows its recruitment to sites of DNA damage through its interaction with damage-sensing factors, fulfilling its function in promoting HR repair.

Markedly, PALB2 occupancy at active genes, as detected by ChIP-qPCR (Figure 9D), was effectively reduced to null following IR exposure (i.e. comparable to the level in cells complemented with empty vector), although it may represent only the sub-fraction of PALB2 bound to the loci tested. Regardless, these observations raise a fundamental question: why does PALB2 dissociate from active genes with the potential risk of leaving these regions unprotected? Notably, the abundance of endogenous PALB2 protein is estimated to be considerably lower than that of BRCA2 or RAD51, for example ~60 times less than BRCA2 and ~600 times less than RAD51 in HeLa cells (Kulak et al., 2014). The majority of PALB2 is also found to be associated with BRCA2 on nuclear structures (Xia et al., 2006) and with MRG15 (Bleuyard et al., 2017b). Hence, we envision that, in endogenous conditions, PALB2 primarily associates with BRCA2 and RAD51 on actively transcribed loci to protect these regions (though PALB2-free pools of BRCA2 and RAD51 presumably exist), but when DNA damage occurs elsewhere, the same complex needs to be mobilised to promote HR repair. In light of these considerations, our experimental platform assessing cells expressing exogenous PALB2 variants, at higher than endogenous levels, might have somewhat compromised sensitivity to highlight the physiological impact of the PALB2 mobilisation. Either way, we envision that the under-enrichment of PALB2 at active genes is unlikely to confer severe consequences for the transcription-replication conflict, as IR would trigger the global DNA damage checkpoint, slowing replication fork progression (Lajitha et al., 1958; Ord and Stocken, 1958; Watanabe, 1974) while broken DNA is repaired.

Intriguingly, while PALB2 7K variants, with either Q or R substitutions, equally interact with BRCA1 in both undamaged and IR-treated conditions (Figure 6—figure supplement 1), their recruitment to sites of DNA damage appeared more affected with the 7R mutant (Figure 6). We postulate that the random mode of the 7R mutant chromatin association prevents its damage-induced mobilisation, which is normally triggered by ChAM deacetylation specifically occurring at transcriptionally active chromatin upon DNA damage and hence making PALB2 globally available for HR. In the meantime, PALB2 recruitment itself does not appear to guarantee RAD51 assembly. Both the 7R and 7Q substitutions elicited significant impairment of RAD51 foci formation (Figures 7 and 8), suggesting that the 7K-patch promotes optimal RAD51 engagement with broken DNA, conceivably in part through its direct interaction with the nucleosome acidic patch regions exposed at damaged chromatin (Belotserkovskaya et al., 2020). It should be noted, however, that the full impact of the 7R and 7Q mutations on HR repair remains unclear. We endeavoured to assess HR proficiency of these cell lines using our reporter systems, which monitor the HR-mediated integration of a promoterless GFP gene and the resultant expression of fluorescent protein as the readout (Rodrigue et al., 2019; Yata et al., 2012). Through these approaches, significant HR defects were not detected (data not shown). This could potentially be explained by the model that the PALB2 7K-patch is particularly important for HR repair of DSBs arising outside active genes, the detection of which is not straightforward using available techniques. Further investigation using more elaborate strategies, which allow the evaluation of HR repair outside active genes, is warranted to fully appreciate the importance of PALB2 ChAM acetylation in HR repair.

Overall, the results of this study highlight the importance of the dynamic regulation of PALB2, where both its acetylated and non-acetylated forms play critical roles in genome integrity control. Our work also suggests that caution should be exercised in the use of amino acid substitutions for functional studies of lysine acetylation. K to Q substitutions confer charge loss, as seen in lysine acetylation, but also significantly reduce the size of the side chain (compared to acetylated lysine) and, hence, do not truly mimic acetyl-lysine. Non-acetylatable K to R substitution similarly affects the size of the side chain, albeit less dramatically than K to Q. These limitations are distinctly reflected in our biochemical analyses of ChAM variants with amino acid substitutions, properties of which did not match those of synthetic ChAM peptides with and without acetylated lysine residues (compare Figures 2G and 3D). Robust in vivo assessment of acetylation events would ideally involve the development of a chemically modifiable residue that exhibits improved similarity to acetylated lysine in a reversible manner.

This study highlights a new molecular mechanism by which KAT2A/2B controls genome stability. Our KAT2A/2B-dependent acetylome study identified 398 novel potential protein targets (Fournier et al., 2016), indicating that the mechanism of KAT2 action in controlling genome stability is multifaceted and more complex than previously anticipated. It is important to note that, in the cell, KAT2A/2B do not act on their own but within large protein complexes, namely the SAGA (Spt-Ada-Gnc5-Acetyltransferase) and the ATAC (Ada-two-A-containing Acetyltransferase) complexes, at transcriptionally active chromatin (Nagy and Tora, 2007). Components of these complexes regulate their enzymatic activity and substrate specificity. Markedly, ENY2 and ATXN7L3, components of the SAGA module that catalyses deubiquitination of histone H2Bub1 have been shown to play a role in preventing unscheduled HR repair in human cells (Evangelista et al., 2018). We postulate that KAT2A/2B-containing complexes concurrently regulate the HR machinery and chromatin in undamaged cells, such that they ensure the activation of HR repair only when needed. KAT2A/2B-containing complexes are key regulators of processes controlling genome stability and will merit further investigation in the future.

More broadly, lysine acetylation is known to control biological processes, such as transcription, based on the metabolic status of the cell (Lee et al., 2014). Therefore, a better appreciation of the role of lysine acetylation in the DDR could expand the scope of studies aiming to understand how reprogrammed metabolism could increase genome instability in cancer cells (Nowell, 1976). Since PTMs are essential for physiological homeostasis and metabolites are necessary cofactors for the deposition of these PTMs, we envision that reprogrammed metabolism in cancer cells could alter the PTM landscape of DDR proteins and hence contribute to genome instability. Future studies assessing aberrant PALB2 lysine acetylation in cancer tissues and whether KDAC or bromodomain inhibitors, currently in clinical trials for cancer therapy, would modulate PALB2 acetylation may offer therapeutic potential. Furthermore, deciphering the metabolic regulation of the DDR and DNA repair could highlight additional cellular pathways that might be targeted for cancer therapy, possibly in combination with existing therapies that otherwise may give rise to resistance.

Appendix 1

Appendix 1—key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Strain, strain background (Escherichia coli)ArticExpress (DE3)Agilent Technologies
Cell line (Homo-sapiens)HEK293TATCC
Cell line (Homo-sapiens)U2OS Flp-In T-RExProf. Daniel Durocher (Lunenfeld-Tanenbaum Research Institute)
Cell line (Homo-sapiens)U2OS Flp-In T-REx FE-PALB2-WTThis studyU2OS Flp-In T-REx harbouring inducible Flag-EGFP-PALB2 WT
Cell line (Homo-sapiens)U2OS Flp-In T-REx FE-PALB2-ΔChAMThis studyU2OS Flp-In T-REx harbouring inducible Flag-EGFP-PALB2 ΔChAM
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNABleuyard et al., 2017bU2OS Flp-In T-REx harbouring inducible shRNA targeting endogenous PALB2
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA-EVBleuyard et al., 2017bU2OS Flp-In T-REx P2shRNA cells, complemented with empty vector (EV)
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA-FLAG-PALB2Bleuyard et al., 2017bU2OS Flp-In T-REx P2shRNA cells, complemented with 3xFLAG-PALB2
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA-FLAG-PALB2 7QThis paperU2OS Flp-In T-REx P2shRNA cells, complemented with 3xFLAG-PALB2-7Q
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA-FLAG-PALB2-7RThis paperU2OS Flp-In T-REx P2shRNA cells, complemented with 3xFLAG-PALB2-7R
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA- FE-PALB2-WTThis paperU2OS Flp-In T-REx P2shRNA cells complemented with inducible Flag-EGFP-PALB2 WT
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA- FE-PALB2-7QThis paperU2OS Flp-In T-REx P2shRNA cells complemented with inducible Flag-EGFP-PALB2-7Q
Cell line (Homo-sapiens)U2OS Flp-In T-REx P2shRNA- FE-PALB2-7RThis paperU2OS Flp-In T-REx P2shRNA cells complemented with inducible Flag-EGFP-PALB2-7R
Antibodyanti-FLAG (Mouse monoclonal)SigmaCat# F1804WB (1:1000)
ChIP (10 μg per sample)
AntibodyControl IgG (Mouse monoclonal)Jackson Immunoresearch015-000-003WB (1:1000)
ChIP (10 μg per sample)
Antibodyanti-pan-acetyl-lysine (Rabbit polyclonal)Cell signalling TechnologyCat# 9441 SWB (1:1000)
Antibodyanti-PALB2 (Rabbit polyclonal)BethylCat# A301-246AWB (1:500)
Antibodyanti-PALB2 (Rabbit polyclonal)Rodrigue et al., 2019WB (1:5000)
Antibodyanti-BRCA2 (Mouse monoclonal)MilliporeCat# OP95WB (1:1000)
Antibodyanti-RAD51 (Rabbit polyclonal)Yata et al., 20147946WB (1:5000)
Antibodyanti-RAD51 (Rabbit polyclonal)BioAcademia70–001IF (1/1000)
Antibodyanti-lamin A (Rabbit polyclonal)SigmaL1293WB (1:2000)
Antibodyanti-vinculin (Rabbit polyclonal)SigmaV9131WB (1:200000)
Antibodyanti-gamma H2AX (Mouse monoclonal)Millipore05–636WB (1:1000)
IF (1:2000)
Antibodyanti-MRG15 (Rabbit polyclonal)Cell signalling TechnologyCat# D2Y4WB (1:1000)
Antibodyanti-BRCA1 (Mouse monoclonal)SigmaCat# OP107WB (1:1000)
Antibodyanti-GST (Mouse monoclonal)Santa Cruz BiotechnologyCat# sc-138WB (1:1000)
Antibodyanti-biotin coupled with horseradish peroxidase (HRP) (Mouse monoclonal)SigmaCat# A0185WB (1:1000)
Antibodyanti-KAT2A/GCN5 (Mouse monoclonal)Cell signalling TechnologyCat# 3305WB (1:1000)
Antibodyanti-alpha-tubulin (Mouse monoclonal)Cell signalling TechnologyCat# 3873WB (1:2000)
AntibodySecondary antibody coupled with horseradish peroxidase (HRP) /
Goat anti-mouse (Goat polyclonal)
DakoCat# P0447WB (1:1000)
AntibodySecondary antibody coupled with horseradish peroxidase (HRP) /
Goat anti-rabbit (Goat polyclonal)
DakoCat# P0448WB (1:1000)
AntibodySecondary antibody coupled with horseradish peroxidase (HRP) /
Goat anti-mouse (Goat polyclonal)
Jackson ImmunoResearch515-035-062WB (1: 20000)
AntibodySecondary antibody coupled with horseradish peroxidase (HRP) /
Goat anti-rabbit (Goat polyclonal)
Jackson ImmunoResearch111-035-144WB (1: 20000)
AntibodySecondary antibody coupled with Alexa Fluor/
Goat anti-mouse (Goat polyclonal)
invitrogenA-11001IF (1/1000)
AntibodySecondary antibody coupled with Alexa Fluor/
Goat anti-mouse (Goat polyclonal)
invitrogenA-11017IF (1/400)
AntibodySecondary antibody coupled with Alexa Fluor/
Goat anti-rabbit (Goat polyclonal)
invitrogenA-11011IF (1/1000)
Chemical compound, drugsodium butyrate (NaB)Sigma303410
Chemical compound, drugTrichostatin (TSA)SigmaT8552
Chemical compound, drugWST-1 reagentMerck Life Sciences Uk Limited5015944001
Sequence-based reagentsiRNA: nontargeting controlDharmaconD001810-10-0550 pmole
Sequence-based reagentsiRNA: targeting KAT2ADharmaconL-009722-02-000550 pmole
sequence-based reagentsiRNA: targeting KAT2BDharmaconL-005055-00-000550 pmole
Recombinant DNA reagentpcDNA5/FRT-GW/N3×FLAGBleuyard et al., 2017b
Recombinant DNA reagentpENTR3CInvitrogen
Recombinant DNA reagentpcDNA-DEST53Invitrogen
Recombinant DNA reagentpCMV-SPORT6-PALB2Source BioSciencesIMAGE clone 6045564
Recombinant DNA reagentpGEX-6P-1GE HealthcareGST expression in bacteria cells
Recombinant DNA reagentpGEX-6P-1_PALB2This paperGST-PALB2 full length expression in bacteria cells
Recombinant DNA reagentpGEX-6P-1_PALB2_Fr1This paperGST-PALB2 fragment 1 (1-320) expression in bacteria cells
Recombinant DNA reagentpGEX-6P-1_PALB2_Fr2This paperGST-PALB2 fragment 2 (295-610) expression in bacteria cells
Recombinant DNA reagentpGEX-6P-1_PALB2_Fr3This paperGST-PALB2 fragment 3 (580-900) expression in bacteria cells
Recombinant DNA reagentpGEX-6P-1_PALB2_Fr4This paperGST-PALB2 fragment 4 (867–1186) expression in bacteria cells
Recombinant DNA reagentpOG44Invitrogen
Recombinant DNA reagentpENTR3C-ChAM #1This paperChAM #1 (395-450) in the gateway entry vector
Recombinant DNA reagentpENTR3C- ChAM #2This paperChAM #2 (395-433) in the gateway entry vector
Recombinant DNA reagentpENTR3C- ChAM #3This paperChAM #3 (353-433) in the gateway entry vector
Recombinant DNA reagentpENTR3C- ChAM #4This paperChAM #4 (353-450) in the gateway entry vector
Recombinant DNA reagentpENTR3C- ChAM #5This paperChAM #5 (353-499) in the gateway entry vector
Recombinant DNA reagentpENTR3C-PALB2Bleuyard et al., 2017bFull length PALB2 in the gateway entry vector
Recombinant DNA reagentpENTR3C-PALB2-7QThis paperFull length PALB2-7Q in the gateway entry vector
Recombinant DNA reagentpENTR3C-PALB2-7RThis paperFull length PALB2-7R in the gateway entry vector
Recombinant DNA reagentpcDNA-DEST53- ChAM #1This paperGFP-ChAM #1 (395-450) expression in mammalian cells
Recombinant DNA reagentpcDNA-DEST53- ChAM #2This paperGFP-ChAM #2 (395-433) expression in mammalian cells
Recombinant DNA reagentpcDNA-DEST53- ChAM #3This paperGFP-ChAM #3 (353-433) expression in mammalian cells
Recombinant DNA reagentpcDNA-DEST53- ChAM #4This paperGFP-ChAM #4 (353-450) expression in mammalian cells
Recombinant DNA reagentpcDNA-DEST53- ChAM #5This paperGFP-ChAM #5 (353-499) expression in mammalian cells
Recombinant DNA reagentpGEX4T3GE HealthcareGST expression in bacteria cells
Recombinant DNA reagentpGEX4T3-ChAMBleuyard et al., 2012GST-ChAM WT expression in bacteria cells
Recombinant DNA reagentpGEX4T3-ChAM-3Q4KThis paperGST-ChAM-3Q4K expression in bacteria cells
Recombinant DNA reagentpGEX4T3-ChAM-3K4QThis paperGST-ChAM-3K4Q expression in bacteria cells
Recombinant DNA reagentpGEX4T3-ChAM-7QThis paperGST-ChAM-7Q expression in bacteria cells
Recombinant DNA reagentpGEX4T3-ChAM-3R4KThis paperGST-ChAM-3R4K expression in bacteria cells
Recombinant DNA reagentpcDNA5/FRT-GW/N3×FLAG-PALB2Bleuyard et al., 2017bConstitutive 3xFlag- PALB2 expression in mammalian cells
Recombinant DNA reagentpcDNA5/FRT-GW/N3×FLAG-PALB2-7QThis studyConstitutive 3xFlag- PALB2 7Q expression in mammalian cells
Recombinant DNA reagentpcDNA5/FRT-GW/N3×FLAG-PALB2-7RThis studyConstitutive 3xFlag- PALB2 7R expression in mammalian cells
Recombinant DNA reagentpcDNA5/FRT/TO/FE-PALB2Bleuyard et al., 2012Inducible Flag-EGFP-PALB2 fusion expression in mammalian cells
Recombinant DNA reagentpcDNA5/FRT/TO/FE-PALB2 7QThis paperInducible Flag-EGFP-PALB2 7Q expression in mammalian cells
Recombinant DNA reagentpcDNA5/FRT/TO/FE-PALB2 7RThis paperInducible Flag-EGFP-PALB2 7R expression in mammalian cells
Recombinant DNA reagentpcDNA5/FRT/TO/FE-PALB2_ΔChAMBleuyard et al., 2012Inducible Flag-EGFP-PALB2 ΔChAM expression in mammalian cells
Sequence-based reagentPALB2-F1_fo1This paperPCR primers5’-atggatccatggacgagcctccc-3’
Sequence-based reagentPALB2-F1_re1This paperPCR primers5’-atgcggccgcattagaacttgtgggcag-3’
Sequence-based reagentPALB2-F2_fo1This paperPCR primers5’-atggatccgcacaaggcaaaaaaatg-3’
Sequence-based reagentPALB2-F2_re1This paperPCR primers5’-atgcggccgctgtgatactgagaaaagac-3’
Sequence-based reagentPALB2-F3_fo1This paperPCR primers5’-atggatccttatccttggatgatgatg-3’
Sequence-based reagentPALB2-F3_re1This paperPCR primers5’-atgcggccgcagctttccaaagagaaac-3’
Sequence-based reagentPALB2-F4_fo1This paperPCR primers5’-atggatcctgttccgtagatgtgag-3’
Sequence-based reagentPALB2-F4_re1This paperPCR primers5’-atgcggccgcttatgaatagtggtatacaaat-3’
Sequence-based reagentPALB2_395_FoThis paperPCR primers5’- actggatcctcttgcacagtgcctg-3’
Sequence-based reagentPALB2_353_FoThis paperPCR primers5’- actggatccaaatctttaaaatctcccagtg-3’
Sequence-based reagentPALB2_450_ReThis paperPCR primers5’- tatctcgagttaatttttacttgcatccttattttta-3’
Sequence-based reagentPALB2_433_ReThis paperPCR primers5’- tatctcgagttacaaatgactctgaatgacagc-3’
Sequence-based reagentPALB2_499_ReThis paperPCR primers5’- tatctcgagttacaagtcattatcttcagtggg-3’
Sequence-based reagentPatch 1-K-RevThis paperPCR primers5’-tcagagtcatttggatgtcaagaaaaaaggttt-3’
Sequence-based reagentPatch 2-WT-FwdThis paperPCR primers5’-aaaaataaaaataaggatgcaagtaaaaat-3’
Sequence-based reagentPatch 1-R-RevThis paperPCR primers5’-tcagagtcatttggatgtcaggagaagagggttt-3’
Sequence-based reagentPatch 2-R-Fwd_FLThis paperPCR primers5’-agaaatagaaatagggatgcaagtagaaatttaaacctttccaat-3’
Sequence-based reagentPatch 1-Q-RevThis paperPCR primers5’-tcagagtcatttggatgtccagcaacaaggttt-3’
Sequence-based reagentPatch 2-Q-Fwd_FLThis paperPCR primers5’-caaaatcaaaatcaggatgcaagtcaaaatttaaacctttccaat-3’
Sequence-based reagentPatch 2-Q-Fwd_ChAMThis paperPCR primers5’-caaaatcaaaatcaggatgcaagtcaaaattgagcggccgcact-3’
Sequence-based reagentBeta-Actin_in3-foThis paperPCR primers5’-taacactggctcgtgtgacaa-3’
Sequence-based reagentBeta-Actin_in3-reThis paperPCR primers5’-aagtgcaaagaacacggctaa-3’
Sequence-based reagentChr5_TCOF1_peak2_foThis paperPCR primers5’-ctacccgatccctcaggtca-3’
Sequence-based reagentChr5_TCOF1_peak2_reThis paperPCR primers5’-tcagggctctatgaggggac-3’
Sequence-based reagentChr11_WEE1_mid_foThis paperPCR primers5’-ggccgaggcttgaggtatatt-3’
Sequence-based reagentChr11_WEE1_mid_reThis paperPCR primers5’-ataaccccaaagaacacaggtca-3’
Peptide, recombinant proteinChAM-WTThis paperFrancis Crick Institute Peptide Chemistry Technology PlatformAEKHSCTVPEGLLFPAEYYVRTTRSMSNCQRKVAVEAVIQSHLDVKKKGFKNKNKDASKN
Peptide, recombinant proteinChAM-K436(Ac)-K437(Ac)-K438(Ac)This paperFrancis Crick Institute Peptide Chemistry Technology PlatformAEKHSCTVPEGLLFPAEYYVRTTRSMSNCQRKVAVEAVIQSHLDVK(Ac)K(Ac)K(Ac)GFKNKNKDASKN
Peptide, recombinant proteinChAM-K436(Ac)This paperFrancis Crick Institute Peptide Chemistry Technology PlatformAEKHSCTVPEGLLFPAEYYVRTTRSMSNCQRKVAVEAVIQSHLDVK(Ac)KKGFKNKNKDASKN
Peptide, recombinant proteinChAM-K437(Ac)This paperFrancis Crick Institute Peptide Chemistry Technology PlatformAEKHSCTVPEGLLFPAEYYVRTTRSMSNCQRKVAVEAVIQSHLDVKK(Ac)KGFKNKNKDASKN
Peptide, recombinant proteinChAM-K438(Ac)This paperFrancis Crick Institute Peptide Chemistry Technology PlatformAEKHSCTVPEGLLFPAEYYVRTTRSMSNCQRKVAVEAVIQSHLDVKKK(Ac)GFKNKNKDASKN
Software, algorithmImage Jhttps://imagej.nih.gov/ij/Schneider et al., 2012
Software, algorithmProteome Discoverer v1.4ThermoFischer Scientific
Software, algorithmFlowJo software V10FlowJo
Commercial assay, kitSensiFAST SYBR No-Rox kitBiolineBIO-98005

Data availability

The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE (Perez-Riverol et al., 2019) partner repository with the dataset identifier PXD014678 and PXD014681. All other data generated or analysed during this study are included in the manuscript and supporting file. Raw images of western blots and DNA/protein gels are avilable through the Open Science Framework.

The following data sets were generated
    1. Fournier M
    2. Esashi F
    (2022) Open Science Framework
    KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity.
    https://doi.org/10.17605/OSF.IO/8E9MS

References

    1. Fouad YA
    2. Aanei C
    (2017)
    Revisiting the hallmarks of cancer
    American Journal of Cancer Research 7:1016–1036.

Decision letter

  1. Wolf-Dietrich Heyer
    Reviewing Editor; University of California, Davis, United States
  2. Jessica K Tyler
    Senior Editor; Weill Cornell Medicine, United States

In the interests of transparency, eLife publishes the most substantive revision requests and the accompanying author responses.

[Editors' note: this paper was reviewed by Review Commons.]

Thank you for submitting your article "KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by a Reviewing Editor and Jessica Tyler as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

The manuscript by Fournier et al. highlights the importance of acetylation in the ChAM domain of PALB2 in regulating nucleosome binding and DNA repair. The text is well-written text and the experiments are well-designed. we read the manuscript, the reviews as well as the rebuttal and plans for a revision. We believe the revision and the proposed added experiments will be needed to cement the conclusions. Of the 5 experiments, #1 and #2 are critical, and we believe #4 and #5 are also important as BRCA1 is a key factor for PALB2 (#5) and the effect on HR (#4) should be documented experimentally. We do not consider experiment #3 (PALB2 foci) as critical. We encourage the authors to plan and execute this revision as they outlined with the above exception of experiment #3. You may want to consider combining KAT2A/B depletion and/or using KDAC inhibitors in the experiments with the 7R and 7K mutants, but we leave that suggestion to them. The authors may take as long as necessary to deliver the revision, under the present COVID19 guidelines of eLife. We look forward to receiving the revised manuscript.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity" for further consideration by eLife. Your revised article has been evaluated by Jessica Tyler (Senior Editor) and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

The efforts in this revision were appreciated by all reviewers. However, as detailed in reviews #1 and 3, there are significant concerns about the overall model and specific experiments, which will require significant additional experimentation.

The key issues are:

1) A fuller characterization of the 7R mutant is required in the HR-reporter assay, the sensitivity assays, in the nucleosome binding assay, and in ChIP assays at transcriptionally-active loci along with some controls. The importance of the analysis of the 7R mutant was highlighted in the previous review cycle.

2) The authors should acknowledge inconsistencies and revise their model to be compatible with their data.

The reviews contain specific critiques and concerns that will require significant changes in the text. The authors may want to consider removing the DNA binding assays.

Reviewer #1 (Recommendations for the authors):

In this revised manuscript by Fournier et al. (eLife-57736R1), the authors have added experiments and analysis to address concerns raised previously. This reviewer appreciates the elegant systems employed in much of this work, as well as the effort that went into this revision. The revised manuscript continues to offer potential insights into the regulation and function of PALB2 in DNA repair, and into the role of acetylation in regulating DNA damage responses. However, there are still major concerns, which in the opinion of this reviewer, should be addressed prior to potential publication. These include issues with certain newly added figures, and apparent inconsistencies or contradictions between different results. As a consequence, the model presented in Figure 7D is not yet fully supported.

1. The 2nd paragraph on p. 21 of the unmarked revision discusses levels of γ-H2AX shown in Figures 5D-E in the context of DNA replication (this paragraph is part of a subsection of the Results beginning on p. 20). Earlier in this same paragraph the authors hypothesize that "PALB2 association with active genes … prevents the induction of DNA damage during DNA replication". In this context, Figures 5D-E are misleading and are not necessarily related to DNA replication. Specifically, the cells were not synchronized and DNA damage arises over several days, leading to questions about the relationship to DNA replication. This would be better supported by performing ChIP assays using γ-H2AX antibody to determine whether DNA damage specifically accumulates at transcriptionally active sites, where conflicts with DNA replication may arise, as compared to other sites. Further, no measures of statistical significance are given for Figure 5E.

2. While HR data was added at the request of a reviewer, the intent is presumably to strengthen the hypothesis that ChAM acetylation regulates PALB2 function in DNA repair. Four points about this:

A) No measure of significance is given for the HR results shown in Figure 7—figure supplement 1. Further, the error bars are often very large. As such, no convincing conclusions can be made about the effects of the 7Q and 7R mutants on HR.

B) In any case, if the finding was sufficiently established that both the 7Q and 7R mutants disrupt DSB-HR, as suggested by the authors, how do they rectify this with the observation that only 7Q but not 7R sensitizes U2OS cells to Olaparib in Figure 7C? Does the 7R mutant perturb HR with no consequences for cellular survival in the presence of DNA damage? And, if so, what additional function is perturbed by the 7Q mutant that results in sensitivity to Olaparib?

C) In the last paragraph on p. 26, the authors write, "We suggest that damage-induced ChAM deacetylation allows its recruitment to sites of DNA damage through its interaction with damage sensing factors, fulfilling its function in promoting HR repair". If all of this is true, since PALB2-7R was never acetylated and therefore did not need to be deacetylated or released from sites of active transcription, then, according to their model, why is this mutant defective for HR (Figure 7—figure supplement 1)? Is this finding consistent with the model presented in Figure 7D?

D) On p. 27 (Discussion), the authors state, "Interestingly, the PALB2 7R variant, unlike its 7Q counterpart, had little impact on RAD51 foci formation … This could be explained by our findings that DNA damage triggers ChAM deacetylation and promotes PALB2 mobilisation, which we propose is the critical event in initiating HR repair". This statement seems to overlook data shown in Figure 7—figure supplement 1 that claims PALB2-7R appears is deficient for HR. Further, proficiency for RAD51 foci in response to IR typically is associated with functional HR, but 7R has nearly normal RAD51 foci while suggested to be deficient for HR. Can the authors' explain this based upon their model?

3. In the model presented in Figure 7D, the authors suggest (1st paragraph of the Discussion, on p. 24) that DNA damage triggers ChAM deacetylation (presumably on PALB2 localized to transcriptionally active chromatin) which "releases PALB2 from active genes and increases its mobility". I have the following concerns about the model (in addition to those detailed in point #2):

A) When discussing their model, it is currently unclear whether the authors are proposing that ChAM deactylation is occurring specifically at "transcriptionally active chromatin" in response to DNA damage and whether this makes PALB2 "globally" available for HR.

B) The data shown in Figures 4B-C seems to contradict the authors' model (Figure 7D). If the role of ChAM deacetylation is simply to release "PALB2 from active genes and increase(s) its mobility", then the 7Q mutant (mimic of acetylated lysine), by being more mobile (Figure 7D-E), should be available for "HR repair complexes" that form in chromatin. Because the 7Q mutant shows less association with chromatin than WT PALB2 or the 7R mutant (Figures 7B-C), one wonders if the 7Q mutant impairs some other PALB2 function besides release from transcriptionally active chromatin upon DNA damage. Additionally, the 7R (non-acetylable) mutant appears functional as indicated by normal cellular resistance to Olaparib (Figure 7C). This does not support any claim there is a specific need that PALB2 be recruited to transcriptionally active chromatin and subsequently released.

C) Unless I overlooked it, it seems that the authors did not specifically test the effect of the 7R mutant on the association of PALB2 (or the ChAM domain) with nucleosomes; this may be helpful in better interpreting various results in the context of their model.

D) Since the 7R mutant shows even less association with transcriptionally active chromatin (Figure 5F) than PALB2-7Q, and PALB2-7R is resistant to Olaparib (Figure 7C), then it would seem that protection of "transcriptionally active chromatin … during DNA replication (2nd paragraph of p. 24) is not essential for cell survival in response to DNA damage.

E) The model that acetylation of the 7K patch releases PALB2 from transcriptionally active chromatin would be better supported by testing this in the presence of IR (or Olaparib), since DSB-initiated HR and cellular resistance to Olaparib are among DNA damage response-related assays featured in the manuscript. Does IR or Olaparib indeed release WT PALB2 from transcriptionally-active loci?

F) According to the model presented, are transcriptionally-active loci left deprotected after acetylation of the ChAM motif that results in increased mobilization of PALB2 for HR?

G) In the absence of assays of PALB2 with transcriptionally active loci after exposure to IR (or PARP inhibitor), the authors have not convincingly demonstrated that PALB2 is mobilized from these loci after DNA damage. While the authors note that Bleuyard et al. 2017b demonstrate that CPT induces decreased association of PALB2 with transcriptionally active chromatin, CPT could have a different effect than IR by inducing more replication stress.

4. The authors note that both K to R and K to Q substitutions alter the size of the side chain, albeit to different degrees. How confident are the authors that K to R and K to Q substitutions are accurate non-acetyl and acetyl mimics, respectively? Is it possible that both substitutions are simply disrupting the structure of the ChAM, and perhaps PALB2 more globally, to some degree?

5. In contrast to association of PALB2 with transcriptionally active chromatin regulating interaction with BRCA1, as proposed, structural changes across the ChAM, or in PALB2 more broadly, could potentially cause the observed decreased interactions of the 7R and 7Q PALB2 mutants with BRCA1.

Reviewer #2 (Recommendations for the authors):

Most of the concerns raised previously have been taken into account and the manuscript has now been much improved. I can therefore recommend this manuscript for publication.

Reviewer #3 (Recommendations for the authors):

Fournier et al. – KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity

The authors have responded to the comments made by each reviewer, and have made several amendments to the manuscript itself, generally improving its legibility and providing additional supporting data / experiments. Saying this, there are still some outstanding questions that should be addressed:

A) It is still not clear as to the (intended) purpose of the GST-ChAM DNA-binding experiments and what these actually contribute to the manuscript as a whole. They could be readily removed from the manuscript without impacting the set of conclusions.

Note: there is no appreciable DNA binding by any of the mutant forms of the GST-ChAM construct as compared to the two controls, WT or indeed PALB2-FL (how was this protein produced, is this the bacterially expressed protein?).

It is therefore not clear how the authors can support the following statement.

Page 18, Lines 444-446: "… however, the ChAM 3R4K variant also showed impaired binding to DNA in an EMSA assay, albeit less pronounced than that of its K to Q (3Q4K) counterpart (Figure 2H)".

B) Whilst it is appreciated that the authors have now created two additional GST-ChAM construct (3Q4K and 3R4K) they haven't performed the requested control experiment, i.e., with the 7R construct. This is necessary to support the set of experiments described at a later point in the manuscript. What is the effect of the 7R construct in the GST-pulldown experiments – does this still bind to nucleosomes / histone H3?

C) Figure 3F, why is there a such a significant increase in PALB2 mobility in the FRAP experiment for the negative control siCntrl? See also Figure 3—figure supplement 1

D) The data for MMC/Olaparib treatment presented in the response to authors (Figure R3) should be included as supplementary data within the revised manuscript.

Page 23, Lines 572-576

"… though similar changes in ChAM acetylation were not detectable upon MMC or olaparib DNA damaging treatment. We noted that IR triggered strong γ- H2A.X induction (Figure 6B), which was not observed upon exposure of HEK293T cells to MMC or olaparib, suggesting that the reduction of ChAM acetylation reflects the cellular response to DNA damage.

E) Figure 7 —figure supplement 1, panel B (HR assay)

The means from individual experiments should be displayed on the graph, as it is not clear if the wide error bar (which presumably should also extend downwards, as + and – 1 SD) and thus interpretation of the result is due to a single outlier measurement.

F) Discussion

The authors allude to unpublished data within the discussion / conclusion – such statements should ideally be removed.

Also, our preliminary results indicate that a PALB2 variant defective in BRCA1 binding exhibits higher accumulation at genic regions than its wild-type counterpart.

Indeed, our preliminary results demonstrated that extracellular glucose concentration affect PALB2 dynamics, as reducing glucose level in the growth medium increases PALB2 dynamics.

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity" for further consideration by eLife. Your revised article has been evaluated by Jessica Tyler (Senior Editor) and a Reviewing Editor.

The manuscript has been significantly improved but there are some remaining issues that need to be addressed, as outlined below:

In the current revision of their manuscript, Fournier et al. have, in general, addressed the concerns that were raised previously. This includes removal of confounding results and addition of new data, such as survival assays, recruitment to sites of DNA damage, and ChIP assays for occupancy at transcriptionally active sites with and without IR, for WT PALB2, and the 7Q and 7R mutants. Overall, the data and presentation are now much more cohesive and are integrated into an interesting model in Figure 9E. As in previous versions, an impressive array of complementary techniques have been employed in this manuscript. This manuscript will make an important contribution to the literature on DNA damage responses.

Having said all of this, it should be noted that the manuscript still requires certain clarifications in the text and/or figures. This will not require additional experimentation only text changes and clarifications that would strengthen this manuscript further and improve overall clarity.

1) There needs to more clarity, about roles in DNA repair/cell survival away from transcriptionally active genes, of the full cellular pool of PALB2 vs the sub-fraction that is specifically associated with transcriptionally active sites. a) Do the authors propose that any PALB2 remains at transcriptionally active sites after the induction of DNA damage elsewhere and if so whether this may provide ongoing protection of active genes and b) whether the authors are proposing that a PALB2-BRCA2-RAD51 complex is mobilized from transcriptionally active genes and whether the PALB2-BRCA2-RAD51 complex exists away from transcriptionally active genes at steady state.

2) The statements about the role of the 7K-patch in HR needs to be softened and clarified how results obtained with the 7R and 7Q variants provide complementary information about PALB2's regulation and function in response to DNA damage. There were reasons for removing data from HR reporter assays, but a reader may consider conclusions about the role of the 7K-patch in HR based only on RAD51 foci to not be fully supported. As such, it is suggested that the authors briefly discuss their HR reporter assay results (in either the Results or Discussion) as data not shown and indicate why these results may not be congruent with those obtained for RAD51 foci. Finally, in the absence of HR reporter assay data, I would suggest toning down the headings in various places to less definitively link the 7K patch to HR repair (line 610; lines 1026-1027; lines 1044-1045. In general, RAD51 focus formation cannot be equalized with HR.

3) A measure of the statistical significance of differences in Figure 4C is needed.

4) It looks like there is a non-specific band in the blot for PALB2 in Figure 6A. It would be helpful to indicate, with a marker in Figure 6A and/or by the description in the figure legend, which, if any, of the two bands is non-specific there.

5) The foci are often difficult to see without significant enlargement in Figures 7D and 8D. Perhaps making this figure part larger relative to the overall figure would help with this. Alternatively, the intensities could be enhanced in the images if the current relative size of the figure part is retained.

6) There is a single statement that appears to have been carried forward from the previous version which should be removed (page 22, line 524-525) 'while suppressing non-specific DNA binding'.

https://doi.org/10.7554/eLife.57736.sa1

Author response

The manuscript by Fournier et al. highlights the importance of acetylation in the ChAM domain of PALB2 in regulating nucleosome binding and DNA repair. The text is well-written text and the experiments are well-designed. we read the manuscript, the reviews as well as the rebuttal and plans for a revision. We believe the revision and the proposed added experiments will be needed to cement the conclusions. Of the 5 experiments, #1 and #2 are critical, and we believe #4 and #5 are also important as BRCA1 is a key factor for PALB2 (#5) and the effect on HR (#4) should be documented experimentally. We do not consider experiment #3 (PALB2 foci) as critical. We encourage the authors to plan and execute this revision as they outlined with the above exception of experiment #3. You may want to consider combining KAT2A/B depletion and/or using KDAC inhibitors in the experiments with the 7R and 7K mutants, but we leave that suggestion to them. The authors may take as long as necessary to deliver the revision, under the present COVID19 guidelines of eLife. We look forward to receiving the revised manuscript.

We thank all the reviewers for taking the time to evaluate our manuscript and for giving us an opportunity to revise our manuscript. As recommended by the eLife Editor, we conducted four key experiments, the results of which are now incorporated in the revised manuscript:

  • Experiment 1: Biochemical analyses of ChAM 7R mutant (Figure 2G and H).

  • Experiment 2: ChIP-qPCR of PALB2 variant at previously identified PALB2 bound genes (Figure 5F)

  • Experiment 4: PALB2 variant homology-directed gene targeting assay (Figure 7—figure supplement 1B)

  • Experiment 5: PALB2 variant interaction with BRCA1 (Figure 7—figure supplement 1D)

In the revised manuscript, we have further extended our discussion to clarify concerns raised by the reviewers. We believe that these newly incorporated results and rewriting significantly improve our manuscript, shedding light on the importance of PALB2 acetylation at the ChAM in both normally growing cells and upon DNA damage.

Below, we provide an updated version of our point-to-point response to the reviewers. Please note that the Figure numbers referenced by the reviewers in the original manuscript do not match those of the revised manuscript. To facilitate straightforward review, our responses include the relevant Figure, page and line numbers for the revised manuscript.

Point-by-point

Reviewer #1 (Evidence, reproducibility and clarity (Required)):

Fournier et al. detect acetylation within the chromatin association motif (ChAM) of PALB2 and demonstrate that KAT2 can acetylate these 7 lysine residues within this region. They then generate K to R mutations (7R) or K to Q mutations (7Q) at these sites and perform assays of fluorescence recovery after photobleaching (FRAP) to measure mobility as a measure of chromatin association, RAD51 foci, PALB2 recruitment at sites of laser-induced DNA damage, and sensitivity to olaparib. They find increased mobility of the 7Q mutant of PALB2 but not 7R in the absence of exogenous DNA damage, as well as defects in DNA damage-induced RAD51 foci and resistance to olaparib. On this basis, the authors conclude that acetylation is required for the association of PALB2 with undamaged chromatin and that deacetylation permits mobilization and association with BRCA1 to enable proper DNA repair. While the manuscript is generally well-written, many of the systems are rather elegant, and this study may yield novel insights into the regulation and function of PALB2 in DNA repair, there are some missing experiments to be added and important contradictions that should be resolved in order to fully establish the new model the authors propose.

Major comments:

1. There are some concerns about the interpretation of experiments with the 7R and 7Q mutants of PALB2. For example, in the description of results in Figure S2C, the authors state "K to R substitutions maintain the charge yet are unable to accept acetylation and hence mimic constitutively non-acetyl lysine". However, in Figure 4B the association of the 7R mutant with chromatin is similar to WT and in Figure 7D,E the relative immobility of the 7R mutant is very similar to WT PALB2. Thus, the conclusion that acetylation is required for PALB2 association with damaged undamaged chromatin and for release of PALB2 upon DNA damage does not appear justified. Perhaps the authors need to better consider whether the 7R mutant mimics acetylation because of its charge. Even so, the mutant then maintains the charge normally associated with acetylated PALB2, calling into question whether deacetylation indeed "releases PALB2 from undamaged chromatin".

We agree with the reviewer’s point that there is no or little difference between WT and the 7R mutant in regard to their enrichment on non-damaged chromatin, as detected by fractionation (Figure 4B and C), or their mobility, as detected by FRAP (Figure 4D and E). To better address the question raised by the reviewer, we have conducted biochemical analyses of the ChAM K to R substitutions, testing their direct interaction with DNA (Experiment 1; Figure 2G and H).

Our analysis of the ChAM 3R4K variant, substituting three conserved lysine residues K436, K437 and K438 with R, revealed that K to R substitutions decreased its binding to DNA, although to a lesser extent than that of a ChAM Q mutant (3Q4K), compared to WT. This observation indicates that (1) ChAM interaction with DNA is mediated not solely by its electrostatic charge, (2) K to R substitutions alters ChAM biochemical properties, and (3) the 7R mutation likely impairs PALB2 association with chromatin as a result of its reduced affinity for DNA. This is described on page 18, lines 437-446.

The discrepancy between this new finding and the fractionation/FRAP results can be explained by the fact that, in a cellular context, full-length PALB2 is enriched at a fraction of H3K36me3-marked exons (which comprise only 1-1.5 % of the genome), as shown in our previous genome-wide ChIP-seq analysis (Bleuyard et al., 2017, PNAS). We reasoned that bulk fractionation or FRAP analyses might not be sensitive enough to highlight the impact of the 7R mutation, but the ChIP-qPCR method, detecting PALB2 association at defined genic regions would be more appropriate. Thus, we have expanded our ChIP-qPCR analyses to validate our proposed model (Experiment 2: Figure 5F).

Indeed, our ChIP-qPCR analysis of PALB2 variants at previously identified PALB2-bound loci (i.e. ACTB, TCOF1 and WEE1) revealed a statistically significant decrease of PALB2 7R at these loci compared to WT PALB2 in undamaged conditions. This observation supports that PALB2 7R elicits decreased PALB2 association at these loci and that bulk fractionation or FRAP analyses were unsuited to detect the impact of the 7R mutation. This point is now discussed on page 22-23, lines 541-560.

Overall, the new results led us to revise our interpretation of the 7R mutant. While K to R substitutions alter the biochemical properties of ChAM, the R-associated electrostatic charge, analogously to non-acetylated K, allows dynamic PALB2 association with chromatin except at those actively transcribed loci which are normally bound by PALB2. We propose that, at KAT2A/B-enriched actively transcribed regions, PALB2 is targeted for acetylation, which in turn promotes PALB2 association with chromatin, likely together with MRG15-mediated tethering. This mechanism might potentially contribute to the suppression of HR repair at actively transcribed regions in undamaged cells. This point is now discussed on pages 29-30, lines 726-740.

2. Related to questions of interpreting results utilizing the 7R and 7Q mutants of PALB2, in Figure 7B,C the 7R mutant but not 7Q supports RAD51 foci and resistance to olaparib similar to WT PALB2. The authors then state in the Discussion that "our work also suggests that caution should be exercised in the use of K to Q substitutions for functional studies of lysine acetylation". Thus, which mutant is giving the correct and reliable results?

We apologise for the miscommunication if this point was unclear. Using biochemical approaches, we established that ChAM acetylation, but not K to Q substitutions, facilitates its association with nucleosomes (please compare Figure 2E and Figure 3B). This observation clearly demonstrates that K to Q substitutions do not mimic acetylation at these residues, but instead render PALB2 ChAM functionally null. The PALB2 7Q phenotypes therefore demonstrate the importance of the 7K-patch for ChAM function in HR repair, rather than its acetylation status. In the revised manuscript, we have emphasised the differences between lysine acetylation and the K to Q substitutions throughout (e.g. page 18-19, lines 446-450; page 19, lines 461-465; page 29, lines 716719).

Perhaps even more importantly, if results with the 7Q mutant are suspect, the conclusion that deacetylation is required for HR (or DNA repair) is suspect because that is the only case where the authors see a defect in RAD51 foci and resistance to olaparib. Similarly, if the 7R mutant "mimics nonacetyl-lysine" then the fact that it has normal RAD51 foci and resistance to olaparib contradicts the conclusion that deacetylation is required for DNA repair.

As responded in the previous points, we now articulated our view in the revised manuscript. Briefly, K to Q substitutions confer loss of charge but also significantly reduce the size of the lysine side chain compared to that of an acetylated lysine, as depicted in Figure 2—figure supplement 1B. We thus believe that changes in the side chain size might negatively impact on PALB2/ChAM binding to chromatin. This thus makes it impossible to assess the effect of charge loss in the context of the 7Q mutation. Importantly, however, the 7Q mutant, which behaves like a ChAMnull mutant as defined in page 18-19, lines 446-450, was used to demonstrate the importance of ChAM 7K-patch in normally growing conditions (Figure 5) and HR-mediated DNA repair (Figure. 7A-C and Figure 7—figure supplement 1B).

K to R substitutions also alter the side chain size, i.e. a modest size increase compared to nonacetylated lysine (Figure 2B). We believe that this affects impaired binding of the ChAM K to R variant to naked DNA in vitro (Figure 2H). Cells expressing the PALB2 7R variant showed delayed S phase progression, DNA damage accumulation in undamaged condition (Figure 5A-E), loss of PALB2 association at defined PALB2 target loci (Figure 5F) and somewhat impaired gene-targeting efficiency upon ZFN-induced DSB induction at the AAVS1 locus (Figure 7—figure supplement 1B). Conversely, we observed little defects of the 7R variant upon exposure to genotoxic treatments in terms of its recruitment to DNA damage sites (Figure 7—figure supplement 1E and F), RAD51 foci formation (Figure 7A and B) and olaparib resistance (Figure 7C). The seemingly normal behaviour of the 7R variant can be explained by our observation that the induction of DNA damage triggers ChAM deacetylation (Figure 6B and C) and an increase of PALB2 mobility (Figure 6D-G). In other words, under conditions where the DNA damage response (DDR) is robustly activated, K to R substitutions might more closely mimic a de-acetylated form of PALB2 and be beneficial for HR.

In summary, it seems clear that amino acid substitutions, K to either Q or R, are not fully optimal in assessing the real impact of acetylation and deacetylation in cellular contexts. Nonetheless, we believe these mutants are useful tools to understand the importance of target residues and to infer the potential impact of acetylation. This point is discussed extensively on pages 28-29, lines 690-723.

3. There are multiple concerns about Figures 5 and S5. In Figure 5A-C, difference in cell cycle progression after synchronization are relatively small and no rationale/interpretation is given for how this may be related to PALB2 function is given. In Figure 5D,E differences in the levels of γ-H2AX as a marker of DNA damage between different forms of PALB2 do not become readily apparent until about 6 or more days after addition of doxycycline. As such, it seems that these could be indirect effects and it is unclear how strongly this supports the importance of PALB2 acetylation in the DNA damage response.

We apologise if these points were unclear. We have previously established that steadystate PALB2 chromatin association, jointly mediated by the ChAM and MRG15 interaction, protects a subset of active genes from DNA damage that may otherwise arise from replication-transcription conflicts (Bleuyard et al., 2017, PNAS). The results presented in Figure 5 led us to propose that PALB2 chromatin association is, at least in part, mediated by the ChAM 7K patch, and its impairment (either by 7Q or 7R substitutions) leaves actively transcribed genes normally bound by PALB2 unprotected and exposed to DNA damage during replication. This model nicely supports our observations that both 7Q/7R mutants exhibit slow S-phase progression and accumulation of γ-H2AX over time. These points are now articulated in the revised manuscript on pages 21-23, lines 523-560:

In Figure S5, it is interesting that there are differences in the association of different forms of PALB2 with 3 distinct active loci, but no error bars or measures of statistical significance are given. Further at 2 of the 3 loci, the association of the 7Q mutant is closer to WT than the 7R mutant. Taken together, neither Figure 5 nor Figure S5 strongly support the key conclusion that acetylation regulates the association of PALB2 with actively transcribed genes to protect them.

We appreciate this criticism. To better evaluate how the ChAM 7K-patch impacts on the association of PALB2 with actively transcribed genes, we have performed biological replicates of the ChIP-qPCR analyses of Flag-PALB2 WT, 7R and 7Q at ACTB, TCOF1 and WEE1 loci (Experiment 2; Figure 5F). The results showed consistently that the 7K substitutions to either 7R or 7Q confer statistically significant decreases in their binding to these loci compared to PALB2 WT. This observation supports our model that PALB2 7Q/7R variants leave actively transcribed genes unprotected and exposed DNA damage, which can cause cellular defects such as slower cell cycle progression.

4. Figures 6D-G and S6A-D conclude that "DNA damage triggers ChAM deacetylation and induces PALB2 mobilization" based upon FRAP experiments utilizing WT PALB2. But there is no control to demonstrate that this is a specific effect driven by the state of PALB2 acetylation. For example, DNA damage might cause global acetylation changes resulting in relaxed chromatin in which proteins that are not subject to acetylation-deacetylation also show increased mobility.

We thank the reviewer for this valuable comment. It is true that we cannot formally exclude the possibility that changes in PALB2 mobility are indirect consequence of damage-induced chromatin reorganisation/increased chromatin mobility. However, our analyses clearly demonstrate that ChAM acetylation increases its association with nucleosomes (Figure 3B), while non-nucleosome binding ChAM-null (7Q or deletion) increases PALB2 mobility (Figure 2E, Figure 4E and Figure 4—figure supplement 1C). Further, WT PALB2 mobility increases after KAT2 depletion (i.e. reduction of chromatin acetylation of KAT2 targets, hence leading to chromatin compaction) (Figure 3F), but reduces upon KDAC inhibition (i.e. global increase in acetylation, hence leading to chromatin relaxation) (Figure 3G). Considering all these observations collectively, the increase in PALB2 mobility detectable upon DNA damage is unlikely to reflect global chromatin relaxation, and that PALB2 acetylation influences its mobility in both challenged and unchallenged cells. We have clarified this point on page 24, lines 583-589.

5. Figure 7B shows that the 7Q mutant has diminished RAD51 foci while Figure S7C,D suggests based upon a different methodology (laser-induced damage) that the 7Q mutant does not affect PALB2 recruitment. Since the issue of recruitment is key to the mechanism proposed, the authors should examine PALB2 foci instead as this may be a more sensitive assay of PALB2 recruitment.

We appreciate the reviewer’s point. Nonetheless, we believe that the laser-induced experiments provide high sensitivity and resolution for PALB2 recruitment kinetics, as the data were obtained with real-time live-cell imaging. In line with this observation, our preliminary foci analysis indicated that PALB2 7Q was efficient in forming foci upon DNA damage (data not shown). We therefore did not extend this analysis, in accordance with the eLife editor recommendation, where PALB2 foci analyses were considered not essential (Experiment 3).

6. The authors state in the last sentence of the Results section that "lysine residues within the ChAM 7K-patch are indispensable for PALB2 function in HR" but never test the mutants for HR using reporter assays. The manuscript would be strengthened by performing such assays.

We have now conducted homology directed recombination (HDR) reporter assays using cells expressing the PALB2 7K variants (Experiment 4; Figure 7—figure supplement 1B). Here, we assessed the gene-targeting efficiency of a GFP-reporter construct that can be integrated at the AAVS1 locus as a readout for HDR. This approach revealed that, compared to cells expressing WT PALB2, GFP integration at the AAVS1 locus upon ZFN-induced DSB was barely detectable in cells expressing PALB2 7Q or 7R (Figure 7—figure supplement 1B and outlined on pages 24-25, lines 603609). This observation supports the notion that the PALB2 7K-patch is indeed important for HDR at this locus under the conditions employed.

Of note, unlike this ZFN-induced HDR reporter assay, the PALB2 7R mutant was competent in supporting IR-induced RAD51 foci formation and olaparib resistance. We propose that different levels of DNA damage might underlie the difference between these phenotypes. 4Gy IR and 1-2 µM olaparib are expected to induce far more DNA damage compared to ZFN-induced DSB (three AAVS1 loci in triploid U2OS cells), triggering robust DDR, where the 7K-patch is naturally deacetylated on WT PALB2, hence little difference between WT and the 7R mutant. Under the condition in which a limited number of DSB is induced, i.e. modest DDR induction, 7K-patch-mediated chromatin association might remain important at least at an actively transcribed AAVS1 locus.

7. The model for the role of ChAM acetylation in regulating PALB2 function presented in Figure 7D is not fully supported by the data presented. Critically, while association with RAD51 and BRCA2 is tested in Figure S7B, the authors hypothesize that deacetylation is required to release PALB2 to enable association with BRCA1 but this is not tested utilizing the mutants.

As suggested, we have assessed BRCA1 association with PALB2 WT and 7R/7Q mutants by pull-down (Experiment 5; Figure 7—figure supplement 1D). Unexpectedly, this analysis revealed decreased binding of BRCA1 to either 7Q or 7R mutant, indicating that PALB2’s competence in associating with active genes (Figure 5F) affects its binding to BRCA1.

On the basis of this observation, we have revised our discussion on the mechanism by which PALB2 7Q and 7R variants could be recruited to sites of DNA damage (pages 28; lines 690-698). Briefly, PALB2 recruitment to sites of DNA damage might be enabled even with a reduced interaction with BRCA1, or without direct interaction with BRCA1, for example via a mechanism that is promoted by DNA-damage-associated small RNA (sdRNA) (Hatchi et al., 2021). Understanding the mechanism underlying this observation warrant future investigation and is now an active area of our investigation.

Also, there are some specific points that should be considered in the context of the model. This includes how DNA damage may trigger deacetylation, and whether it is the deacetylated state or the process of deacetylation of ChAM that is critical. Also, if acetylation is important for protecting active genes in the absence of DNA damage, is deacetylation necessary to release PALB2 local or global. This is important, because if it is local there needs to be a specific mechanism for local deacetylation, while if deacetylation is global that could result in transcriptionally active genes becoming unprotected.

We thank this reviewer for this valuable comment. We agree that, while this study establishes that ChAM is deacetylated upon DNA damage, it remains unclear whether the dynamic ‘de-acetylation’ of PALB2 is important for HR repair, and whether or not this is a local event. Regardless, we would like to highlight that PALB2-bound genes are mostly periodic, e.g. those required for cell cycle progression (Bleuyard et al., 2017, PNAS). It would therefore be reasonable to speculate that DNA damage triggers the suppression of periodic gene expression as a part of DNA damage checkpoint signalling, possibly in a KDAC-dependent manner, which then allows release of PALB2 without risking DNA damage that could otherwise be caused by replication-transcription conflict. Mobilised PALB2 might then be recruited to sites of DNA damage for HR repair. In the revised manuscript, we made our best effort to better describe our model, accompanied by the revised model shown in Figure 7D.

Further study will be required to fully evaluate this model, for example by identifying the specific KDAC involved in ChAM deacetylation and tracking individual PALB2 molecules, which we consider to be beyond the scope of the present study.

Minor Comments:

a. Some parts of the Materials and methods are overly long (such as the subsection on "Protein purification" and "immunofluorescence microscopy") and could be shortened by consolidating experimental details that are largely the same for related processes.

According to the eLife manuscript guideline, we provide all Materials and methods in the main text.

b. In the description of Figure 1D, the statement "7K-patch, which is common to PALB2 orthologs" is misleading since there is not complete conservation of each lysine residue across each ortholog.

We apologise for this error. We have now amended the description on page 17, lines 404407: ‘Notably, while sequence alignment of PALB2 orthologues showed some divergence in the Cterminal region of ChAM, an enrichment of lysine residues was consistently detected (Bleuyard et al., 2012) (Figure 1D).

c. Figures 3E,F and S3B,C perform FRAP in cells with knockdown of KAT2A/B as a surrogate for chromatin association. The authors note that this global reduction in acetylation increases PALB2 diffusion, but there is concern that this experiment is not very informative because the increased mobility may have nothing to do acetylation of PALB2.

Please refer to our answer to this reviewer’s point 4.

Reviewer #2 (Evidence, reproducibility and clarity (Required)):

This manuscript reports the control of PALB2 – chromatin interaction by the acetylation of a particular lysine-rich domain of the protein called ChAM. This acetylation is shown to be mediated by the acetyltransferases KAT2A/B. Following these investigations, the authors made an effort to place their findings in the context of DNA replication and DNA repair.

The proposed model is that the acetylation-dependent interaction of PALB2 with chromatin could ensure the protection of the genome during DNA replication and control DNA repair.

Specific remarks

1) Based on different experiments, essentially the one shown in Figure 3B, the authors conclude that the acetylation of the ChAM domain enhances its association with nucleosomes.

However, taking into account the experimental setting, this conclusion should be largely tuned down. Indeed, this enhanced acetylation-dependent nucleosome binding was observed when the experiment was carried out in the presence of excess of free naked DNA.

Under these conditions, the non-acetylated ChAM fragments became mostly trapped by DNA (clearly shown in Figure 3C/D), and hence would not be available for nucleosome binding, while the acetylated ChAM fragments would remain available for nucleosome association because of their reduced DNAbinding ability.

Consequently, the acetylation of the ChAM domain would only play a role on the availability of PALB2 for chromatin/nucleosome binding and not directly stimulate nucleosome binding. Therefore, the nucleosome-binding capacity of ChAM by itself should not be dependent on ChAM domain acetylation.

If true, this hypothesis could also be relevant in vivo since the poly-K in the ChAM domain could also non-specifically interact with nuclear RNAs and hence its acetylation, by releasing it from nuclear RNAs, would make it available for chromatin-binding.

The importance of RNAs in the regulation of PALB2 nucleosome-binding could be tested in the experiments shown in Figure 2C and 2E by adding RNase to the pull-down medium (WT +/-RNase or addition of increasing exogenous RNAs).

We are grateful for the reviewer’s detailed comments and find the potential involvement of RNA very intriguing. Indeed, transcriptionally active loci, which are bound by PALB2, are enriched in nascent RNA, and such local RNA may play an important role in promoting the association of acetylated PALB2 with nucleosomes. However, we believe that investigating the role of RNAs in PALB2 nucleosome binding is beyond the scope of this study. As discussed extensively in response to this Reviewer’s point 2 below, we believe the mode of interaction of ChAM with nucleosomes to be highly complex, being jointly mediated by the N-terminal conserved region and the C-terminal lysine cluster. In the revised manuscript, we have discussed in greater details the model of ChAM interaction with nucleosomes (pages 26-27, lines 638-660), based on published and current results presented in this study.

2) The real question is as follows. While acetylation makes the protein available for nucleosome binding, which part of the ChAM domain is actually mediating nucleosome binding and whether lysine acetylation could be directly involved in this binding. Another question would be to identify the elements in the nucleosome mediating this interaction, histones (core domain, tails, posttranslational modifications, specific histone types), histone-DNA, etc…

We entirely agree with the reviewer’s question. Despite the increasing recognition of the physiological importance of the PALB2 ChAM and our efforts in understanding the mode of association of ChAM with nucleosomes (including the potential involvement of histone tail modifications), this specific question remains enigmatic.

Explicitly, our previous work demonstrated that substitutions of residues within the evolutionarily highly conserved N-terminal part of the ChAM perturb its association with nucleosomes (Bleuyard et al., 2017, PNAS; Bleuyard et al., 2017, Wellcome Open Research). A recent study by the laboratory of Prof Jackson proposed that basic residues across the ChAM are part of a binding interface with an acidic patch of histone H2A in its nucleosomal context (Belotserkovskaya et al., Nat Comm. 2020). Our results presented in this study introduced an additional complexity, showing that the C-terminal 7K basic patch is essential for ChAM-nucleosome interaction. Intriguingly, our study also suggests that the regions flanking ChAM, which are phosphorylated at multiple residues, play roles in regulating ChAM binding to nucleosomes (Figure 2B and C; please refer also to our answer to the reviewer’s minor point 6). The summary of these observations is shown in Figure 9—figure supplement 1.

We are currently working towards solving the structure of ChAM in complex with a nucleosome, which may help to clarify this very important question. At this point, we think a complete description of all the elements required for ChAM-nucleosome interaction is beyond the scope of this manuscript, and should be addressed in future work. In the revised manuscript, we have provided an updated overview of the ChAM elements affecting nucleosome interaction and post-translational modifications (acetylation, phosphorylation) flanking ChAM regions, as summarised in Figure R1, are now presented in Figure 7—figure supplement 2.

3) Taking into account the authors conclusions on the role of ChAM domain acetylation and its impact on PALB2 mobility, in Figure 4D/E, one should expect a difference of t1/2 when wild-type and 7R mutant are assayed by FRAP. At least the measures of t1/2 in the wild-type should have been more heterogeneous compared to the 7R mutant due to the acetylation of the wild-type PALB2 by the endogenous HATs (the impact of endogenous HATs on the wild-type sequence is shown in Figure 3F). Could the authors comment on this?

We appreciate this reviewer’s point, which is related to reviewer 1’s points 1 and 3, questioning why no difference between WT and the 7R mutant was detected by FRAP assay and cellular fractionation. To address this issue carefully, we have conducted further characterisation of PALB2 K to R variants using DNA binding assays and ChIP-qPCR (Experiments 1 and 2).

As outlined in our response to reviewer 1’s points 1 and 3, we found that K to R substitutions reduced ChAM binding to naked DNA in vitro (Experiment 1; Figure 2 H) and PALB2 association with three target genes ACTB, TCOF1 and WEE1 in vivo (Experiment 2; Figure 5 F). Overall, these analyses revealed that K to R substitutions cause reduced DNA and chromatin association at defined loci in a manner independent of electrostatic charges. The discrepancy between these new results (DNA binding and ChIP-qPCR) and previously presented results (FRAP assay and cellular fractionation analysis) can be explained by the fact that PALB2 associates with only a small subset of genes (Bleuyard et al., 2007, PNAS); FRAP assays and cellular fractionation analyses were most likely not sensitive enough to detect minute but critical differences. This point is now extensively discussed in the revised manuscript p19-20, lines 487-506.

4) It would be better to remove the data presented in Figure 5 since, as currently presented, these investigations remain shallow and do not bring much information on what is happening. The presented data are rather confusing since, in the absence of further investigations, it is not clear which one(s) of the mechanisms involved in the control of DNA replication is controlled by PALB2 and many explanations, including artefacts, remain possible.

The manuscript would gain in interest if the authors would devote the functional studies only to the repair part (Figure 6 and 7).

Our ChIP-qPCR analyses revealed significantly decreased binding of PALB2 7R and 7Q compared to WT at previously defined PALB2-bound loci (Figure 5 F). This observation now nicely explains why both PALB2 7R- and 7Q-expressing cells displayed delays in cell cycle progression and accumulated DNA damage over time, likely due to a lack of protection of PALB2-bound genes during replication in both these cell lines. We therefore feel it is important to keep these results in the manuscript to allow readers to comprehend the role of the 7K-patch in both undamaged and damaged conditions.

Minor points

5) High background of non-enzymatic acetylation of PALB2 fragments makes the identification of KAT2A/B specific acetylation not very convincing.

The immunoblot detection of acetylation fragments shown in Figure S1 is much more convincing. Therefore, the authors may consider to present Figure S1 as a main Figure and Figure 1B as a supplementary one.

As suggested by the reviewer, panels Figure 1—figure supplement 1B and Figure 1B have now been swapped in the revised manuscript.

6) It would be interesting if the authors would comment on why the presence of regions flanking the ChAM domain (Figure 1A, construct #5) significantly reduces chromatin (Figure 1B) and nucleosome binding (Figure 1C).

We are grateful for this reviewer’s comment. Indeed, we noticed that the inclusion of the ChAM C-terminal flanking region perturbs its chromatin association. This region is highly enriched in serine and threonine residues which could be targeted for phosphorylation by cell cycle regulators (CDKs and PLK1) and DNA damage-responsive kinases (ATM and ATR). It is tempting to speculate that, when phosphorylated, this flanking region could mask the basic patch of the ChAM, hence facilitating the release of PALB2 from undamaged chromatin region and its recruitment to sites of DNA damage. In the revised manuscript, we provide the complete list of PTMs in Figure 7—figure supplement 2 and discuss this point on page 26-27, lines 638-660.

Reviewer #3 (Evidence, reproducibility and clarity (Required)):

KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity

The authors describe a series of experiments examining the consequence of acetylation, within a defined motif (Chromatin Association Motif; ChAM), on the cellular roles of the protein PALB2 (Partner and Localizer of BRCA2).

The key conclusions drawn by the authors are generally convincing and are supported by the presented experimental results, which indicate that acetylation of PALB2 by KAT2A/KAT2B modulates its cellular behaviour and response to DNA damage. However please see specific comments below:

Major Comments

Expression of full-length PALB2 in the heterologous host E. coli is highly problematic, as the WD40 domain is generally not correctly folded. The authors use the ArticExpress strain to try and solve/alleviate this problem – but it is clear from the Materials and methods section that an ATP-wash step has had to be introduced in order to release the recombinant protein from the chaperone system encoded by the ArticExpress system; i.e. indicating poor / mis-folding. Whilst this does not strictly have an effect on the results presented in Figure 1 (detection of in vitro acetylation sites), they have implications for the wider scientific community, as this may lead to the erroneous assumption that is possible to produce functional / folded full-length PALB2 in this way.

We apologise if the manuscript conveyed the message that we are able to produce functionally active, full-length PALB2 in bacteria, which was clearly not our intention. Our aim was to test whether KAT2A was able to acetylate PALB2 in vitro. We agree that the folding and the biochemical properties (e.g. WD40-mediated BRCA2 binding) of the bacterially produced full length PALB2 were not fully assessed. We believe that this does not affect the overall conclusions of this study. In the revised manuscript, the error has now been clarified on page 9, line 219 in Material and Methods.

In vitro modification assays are prone to producing post-translational modifications that are not fully reflective of those observed in vivo, and therefore need to be treated with some caution. This is highlighted by the relatively low modification of K438 in vitro by KAT2A; esp. as this is an acetylation site that has been previously mapped in vivo (by the authors). It would have been useful to include / see the effects on PALB2 function in vivo by modification / alteration of this single site.

We appreciate the reviewer’s comment. Redundancy of acetylation acceptor residues within a lysine cluster is common, as is also the case for many ubiquitination events, hence we analysed the 7K-patch mutant for phenotypic studies. For the same reason, we trust that the outcome of the characterisation of a K438 mutant would not significantly change our conclusions.

Figure 3C and Figure 3D do not fully support or reflect the conclusions drawn by the authors – any peptide containing a cluster of positive charged residues are likely to interact with DNA through charge neutralisation of the phosphodiester backbone, concomitantly any alteration to this region of charge (i.e. via acetylation) will perturb this interaction.

We totally agree with the reviewer’s view. In the revised main text referring to the results shown in Figure 3C and D, we state “As anticipated, lysine acetylation, which neutralises the positive charge on the lysine side chain, conferred reduced affinity for negatively charged DNA” (page 20, lines 475-477).

Furthermore, experiments performed with the synthetic acetylated peptides do not agree with those carried out with the GST-ChAM constructs – GST-ChAM interacts with the nicked and linear forms of the pBS plasmid (Figure 2F) but does not interact with the supercoiled form. The WT synthetic ChAM peptide, in contrast, interacts with all three plasmid states at high concentrations. It is suggested that these two figures are removed.

It is true that we cannot exclude potential differences between GST-ChAM and synthetic ChAM peptides: for example, the 26 kDa GST, which can form a dimer, might partly affect the biochemical properties of ChAM in DNA binding. However, we believe that the differences are more likely caused by the concentration of ChAM used. While we used the synthetic ChAM peptides at concentrations of 2.97, 5.94, 29.3 µM for Figure 3C, we used 5.94 µM of GST-ChAM for Figure 2F; we apologise for the omission of the exact experimental conditions used. This notion is supported by the side-by-side experiment shown id Author response image 1. In the revised manuscript, we have included the concentration of GST-ChAM used in Figure 2F (page 34, line 853) to be clear.

Author response image 1
ChAM binding to DNA is dose Dependent.

Indicated concentration of recombinant GST-ChAM and synthetic ChAM were incubated with 300 ng of pBluescript, and DNA-binding was assessed by EMSA. Binding to DNA was not detectable with up to 300 nM ChAM in our previous study (Bleuyard et al., 2012, EMBO Rep).

p. 18: the authors used a PALB2 variant, where the lysines in the 7K patch are mutated to arginine – but don't fully characterise the effects of introducing these particular mutations on the ability of the ChAM fragment to bind to DNA, or indeed to nucleosomes; this is an important control.

We appreciate the reviewer’s comment and, indeed, the importance of the 7R variant biochemical characterisation for accurate interpretation of in vivo phenotypes. We have now conducted the biochemical characterisation of a ChAM K to R mutant (Experiment 1; Figure 2G and H). We have generated a ChAM mutantwith K to R substitutions at positions K436, K437 and K438 (the ChAM 3R4K variant) and purified it from E. coli (Figure 2G). Our analyses of the ChAM variants revealed reduced DNA binding by ChAM 3R4K compared to wild-type (Figure 2H), albeit better binding than 3Q4K. This observation was unexpected as K to R substitutions are expected to maintain electrostatic charges.

These observations led us to revise our interpretation of the 7R variant; K to R substitution impairs the function of ChAM in associating defined transcriptionally active loci (Figure 5F), although maintains better overall chromatin association compared to the K to Q variant (Figure 4). This explains why both K to Q/R substitutions confer defects in some cellular phenotypes, such as cell growth in undamaged conditions (Figure 5A-E) and homology mediated gene-targeting efficiency (Figure 7—figure supplement 1B). This point is extensively discussed in the revised manuscript pages 21-23, lines 503-560, under the section ‘PALB2 ChAM 7K-patch acetylation is required for the protection of actively transcribed genes during DNA replication’

Please also refer to our response to reviewer 1’s points 1 and 3 and reviewer 2’s point 3.

Figure 6: it would be good to show a second supporting example for deacetylation of PALB2 in response to DNA damage – perhaps treatment with MMC?

We appreciate the reviewer’s comment. Indeed, we have conducted the analysis upon MMC and olaparib exposure. Curiously, however, no clear change in ChAM acetylation was detectable (Author response image 2). Note that, for this experiment, we assessed the acetylation level of a GFP-fusion of ChAM, exogenously expressed in HEK293T cells, along with endogenous γ-H2AX as a readout for DNA damage signalling. Unlike ionising radiation, which triggered strong induction of gammaH2AX (Figure 6), no clear increase of γ-H2AX was detectable upon the MMC/olaparib exposure conditions used. Hence, we propose that the reduction of ChAM acetylation reflects the cellular response to DNA damage. These points have now been clarified in the revised manuscript (pages 23, lines 568-576).

Author response image 2
ChAM acetylation upon DNA damage treatment.

HEK293T cells transiently expressing GFP-tagged ChAM were treated with (A) 2.5μM olaparib and collected after 0, 15, 30, 60 or 120min of treatment and (B) 1mM mitomycin C (MMC) and collected after 0, 15, 30, 60, 120 or 240min of treatment. In both conditions, affinity-purified GFP-ChAM acetylation was assessed using anti-GFP and anti-acetyl-lysine (pan-AcK) antibodies. γ-H2AX signal in whole cell lysate (input) was detected to monitor DNA damage.

Minor Comments

p. 16: 'Our MS analysis of the chromatin-associated GFP-ChAM fragment identified actelyation of all seven lysines within the 7K-patch (Figure 3A, marked with arrows).

This part of the manuscript is potentially a little confusing, as Figure 3A references a series of synthetic peptides rather than the GFP-ChAM fragments themselves.

We apologise for the confusion that we have now corrected. Indeed, Figure 3A shows (1) MS of the chromatin-associated fraction of GFP-ChAM (the top part with arrows) and (2) a schematic diagram of synthetic peptides that we used for biochemical analyses (the bottom part). Figure 3A has been modified accordingly.

p. 20: Furthermore, using the FRAP approach, we observed clear differences in diffusion rates of FEPALB2 following damage by IR, MMC, or olaparib treatment…

FE-PALB2 = FL-PALB2?

We apologise for the confusion. In our study, FE-PALB2 refers to Flag-EGFP tagged PALB2 (full-length). This is defined in the text “To this end, a tandem FLAG- and EGFP-tagged full-length wildtype (WT) PALB2 (FE-PALB2)” (page 20, lines 486-487).

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined below:

The efforts in this revision were appreciated by all reviewers. However, as detailed in reviews #1 and 3, there are significant concerns about the overall model and specific experiments, which will require significant additional experimentation.

The key issues are:

1) A fuller characterization of the 7R mutant is required in the HR-reporter assay, the sensitivity assays, in the nucleosome binding assay, and in ChIP assays at transcriptionally-active loci along with some controls. The importance of the analysis of the 7R mutant was highlighted in the previous review cycle.

We have thoroughly reconducted the requested experiments to the best of our ability. These re-evaluations have revealed more detailed properties of the PALB2 7Q and 7R mutants, leading to some key changes in our interpretation and the model.

In brief, despite our greatest efforts, we were unable to obtain concrete evidence supporting the involvement of the PALB2 7K patch in HR repair using HR reporter assays (i.e. DSB-induced gene targeting at AAVS1 or LMNA loci). Also, the olaparib sensitivity of the 7Q variant proved to be insignificant. However, it was clear that S phase RAD51 foci were less efficiently formed in both the 7Q and 7R mutants upon irradiation (Figure 7B and E). Significantly, we also found impaired RAD51 foci formation in the 7Q and 7R mutants even in unperturbed conditions (Figure 8B and E). This is followed by a surge of S phase γ-H2A.X foci (Figure 8F) and impaired cell survival (Figure 9B). These observations suggest that these mutants are defective in RAD51 foci formation at randomly distributed DNA damage, stochastically occurring as part of normal physiology (e.g. metabolically generated ROS), resulting in the accumulation of unrepaired DNA and consequential cell death.

Interestingly, PALB2-7R, but not PALB2-7Q, was less efficiently recruited to DNA damage induced by micro-irradiation (Figure 6), revealing distinct mechanisms underlying the impaired RAD51 foci formation with these mutants. Specifically, the PALB2-7R mutant fails to be recruited to these sites effectively, but PALB2-7Q, while being successfully recruited, is defective in promoting RAD51 foci formation.

Biochemically, recombinant ChAM 7R, but not ChAM 7Q, binds purified nucleosomes, albeit to a lesser extent than WT (7K) (Figure 2D and E). This finding corroborates our observations that the PALB2 7R largely remains in the chromatin-enriched fraction (Figure 5). AlphaFold2 modelling further predicts greater structural plasticity of ChAM WT (7K) compared to ChAM 7Q or ChAM 7R (Figure 2—figure supplement 2), indicating that ChAM WT has better capacity to explore various structural configurations for its optimum association with nucleosomes compared to the 7R and 7Q counterparts. Indeed, both 7R and 7Q binding to active genes are severely compromised compared to WT (Figure 9D). IR triggered robust release of WT PALB2 from these loci, concomitantly with the ChAM deacetylation.

2) The authors should acknowledge inconsistencies and revise their model to be compatible with their data.

Taking all experimental results together, we propose that the ChAM 7K-patch regulates dual modes of PALB2 chromatin association, namely (1) the steady-state PALB2 association with actively transcribed chromatin through the acetylatable 7K, and (2) the damage-induced PALB2 engagement with damaged chromatin through the non-acetylated 7K. Specifically, in unperturbed conditions, WT PALB2 resides at actively transcribed chromatin through MRG15 (Bleuyard et al., PNAS, 2017) and KAT2A-mediated acetylation of the 7K-patch (this study), which is important for protecting active genes from replicative stress (Bleuyard et al., PNAS, 2017). Spontaneous DNA damage, IR and potentially other genotoxic stresses trigger the deacetylation of the 7K-patch, enabling PALB2 mobilisation and its re-localisation to DNA damage sites, including those distant from PALB2-bound actively transcribed regions. At sites of DNA damage, the non-acetylated 7K-patch further supports efficient RAD51 loading onto ssDNA, likely involving its non-specific electrostatic interaction with exposed nucleosomes (Belotserkovskaya et al., Nat Comms, 2020). In this way, WT PALB2, but not the 7Q or 7R mutant, prevents the accumulation of DNA damage and facilitates survival.

The absence of phenotypes with the HR reporter system and upon olaparib exposure was unexpected. It is noteworthy, however, that the HR reporter systems monitor DNA damage repair within active genes (i.e., AAVS and LMNA) and that PARP1 non-randomly occupies regulatory regions of active genes (Krishnakumar, R., Science, 2008; Krishnakumar and Kraus, Mol Cell, 2010). Hence, it is likely these systems highlight HR events only at actively transcribed genes and nearby loci. We suggest that the regulation of the 7K-patch has more pronounced impact on DNA damage in regions that are distant from transcriptionally active genes. Regardless, given the uncertainty of these observations, we have removed these results from the revised manuscript.

The reviews contain specific critiques and concerns that will require significant changes in the text. The authors may want to consider removing the DNA binding assays.

As suggested, we have removed the DNA binding assays, incorporated a significant amount of new experimental data and also considerably changed the main text according to the results. We thank the reviewers for constructive criticisms, which have improved the study and made it more robust.

Reviewer #1 (Recommendations for the authors):

In this revised manuscript by Fournier et al. (eLife-57736R1), the authors have added experiments and analysis to address concerns raised previously. This reviewer appreciates the elegant systems employed in much of this work, as well as the effort that went into this revision. The revised manuscript continues to offer potential insights into the regulation and function of PALB2 in DNA repair, and into the role of acetylation in regulating DNA damage responses. However, there are still major concerns, which in the opinion of this reviewer, should be addressed prior to potential publication. These include issues with certain newly added figures, and apparent inconsistencies or contradictions between different results. As a consequence, the model presented in Figure 7D is not yet fully supported.

1. The 2nd paragraph on p. 21 of the unmarked revision discusses levels of γ-H2AX shown in Figures 5D-E in the context of DNA replication (this paragraph is part of a subsection of the Results beginning on p. 20). Earlier in this same paragraph the authors hypothesize that "PALB2 association with active genes … prevents the induction of DNA damage during DNA replication". In this context, Figures 5D-E are misleading and are not necessarily related to DNA replication. Specifically, the cells were not synchronized and DNA damage arises over several days, leading to questions about the relationship to DNA replication. This would be better supported by performing ChIP assays using γ-H2AX antibody to determine whether DNA damage specifically accumulates at transcriptionally active sites, where conflicts with DNA replication may arise, as compared to other sites. Further, no measures of statistical significance are given for Figure 5E.

We appreciate this reviewer’s comment and assessed the level of γ-H2A.X in S-phase cells, marked by EdU incorporation (Figures8 and 9). This approach is widely used to robustly assess replication-associated DNA damage. This assessment confirmed that the 7Q and 7R mutants indeed accumulate more γ-H2A.X in S phase cells upon shRNA-mediated depletion of endogenous PALB2; this difference reached statistical significance at day 5 after the expression of shRNA. This finding is reflected by the reduced survival of cells expressing the 7Q or 7R mutant even in the absence of genotoxic treatments.

Unfortunately, γ-H2A.X ChIP-qPCR proved challenging with the currently available antibody and we were unable to validate whether these S-phase γ-H2A.X foci indeed arise at transcriptionally active sites in 7R/Q cells. Nonetheless, we consider this very unlikely as, in our previous study, we observed no increase of γ-H2A.X at these loci in cells expressing another PALB2 variant which is similarly unable to bind active genes (via MRG15 interaction), unless challenged by CPT, which induces covalent TOP1-DNA binding (Bleuyard et al., PNAS, 2017).

Significantly, while characterising the cellular phenotypes, we noticed impaired S-phase RAD51 foci formation even in unperturbed 7R/Q cells on day 4, prior to the rise of γ-H2A.X in these cell lines at day 5 (Figure 8). A similar trend was found in 7R/Q cells exposed to IR on day 4 (Figure 7). Given that IR stochastically induces various types of DNA damage across the genome, which could, in turn, impede DNA replication, these observations suggest that the 7Q/R mutants are both defective in the efficient recruitment of RAD51 to DNA damage at random locations of the genome, conferring the global accumulation of DNA damage.

We further revealed that the PALB2 7R, but not the 7Q counterpart, is less efficiently recruited to laserinduced DNA damage (Figure 6), suggesting that the mechanism underlying RAD51 recruitment to sites of DNA damage is different between these mutants. We propose that defective engagement of the 7Q mutant with damaged chromatin undermines stable RAD51 foci formation, while the 7R mutant, which is ‘glued’ to chromatin non-specifically through electrostatic interaction, failed to be mobilised upon DNA damage, conferring impaired recruitment of RAD51 to sites of DNA damage.

2. While HR data was added at the request of a reviewer, the intent is presumably to strengthen the hypothesis that ChAM acetylation regulates PALB2 function in DNA repair. Four points about this:

A) No measure of significance is given for the HR results shown in Figure 7—figure supplement 1. Further, the error bars are often very large. As such, no convincing conclusions can be made about the effects of the 7Q and 7R mutants on HR.

Despite our greatest effort, we were unable to detect significant HR defects using our HR reporter systems. Specifically, we exploited two reporter assays to evaluate HR competency, by inducing DSB at AAVS1 by ZFN or at LMNA by Cas9, and providing a matching donor plasmid carrying GFP with homologous arms. Successful integration was monitored through the expression of GFP, quantified by FACS or microscopy, and was used as the readout of HR proficiency. However, neither of these assays has provided consistent results in relation to the function of the 7K-patch in HR repair. We highlight, however, several limitations of these HR reporter assays. Firstly, these assays depend on high transfection efficiency and only detect integration of a reporter plasmid at defined, actively transcribed loci (i.e., AAVS and LMNA), which might not be representative of all types of HR across the genome. Also, unlike physiological HR repair which uses an unbroken sister chromatid as a repair template, these assays use exogenously introduced plasmid DNA as an HR template. Finally, these assays introduce ‘clean’ two-ended DSB, which is likely processed differently from those naturally occurring ‘dirty’ DNA breaks. Accordingly, we have decided to remove these gene-targeting results.

In the revised manuscript, we have conducted thorough analyses of RAD51 foci (Figures7 and 8), which usually reflects the HR competency. Indeed, RAD51 foci formation is widely used as a reliable readout of HR proficiency in the field, (as the reviewer also comments in D) in this section below (‘proficiency for RAD51 foci in response to IR typically is associated with functional HR’). Accordingly, we infer HR proficiency on the basis of RAD51 foci formation.

B) In any case, if the finding was sufficiently established that both the 7Q and 7R mutants disrupt DSB-HR, as suggested by the authors, how do they rectify this with the observation that only 7Q but not 7R sensitizes U2OS cells to Olaparib in Figure 7C? Does the 7R mutant perturb HR with no consequences for cellular survival in the presence of DNA damage? And, if so, what additional function is perturbed by the 7Q mutant that results in sensitivity to Olaparib?

Through our careful re-evaluation, it has become evident that all our cell lines show comparable resistance to olaparib with similar levels of γ-H2A.X and RAD51 foci. While the source of PARP-trapping lesions remains contentious, PARP1 is shown to be intimately linked with transactivation (Krishnakumar, R. Science, 2008; Krishnakumar and Kraus, Mol Cell, 2010). Hence, it is tempting to speculate that olaparib induces more lesions at transcriptionally active chromatin. The lack of phenotypes in the PALB2 7K patch mutant upon olaparib exposure could be explained by a model in which PARP-trapped lesions at actively transcribed chromatin can be effectively repaired without significant involvement of the PALB2 7K patch or, more broadly, without the steady-state presence of PALB2 at these regions. Indeed, transcription itself has been shown to enhance HR efficiency (Ouyang et al., Nature, 2021).

Accordingly, in the revised manuscript, we shift our focus to the cellular phenotypes in unperturbed conditions and IR response, where statistically significant impairment of S-phase RAD51 foci was observed in both the 7Q and 7R mutant cells (see our response to point 1 above, also outlining our revised model that explains the molecular mechanism underlying these phenotypes).

C) In the last paragraph on p. 26, the authors write, "We suggest that damage-induced ChAM deacetylation allows its recruitment to sites of DNA damage through its interaction with damage sensing factors, fulfilling its function in promoting HR repair". If all of this is true, since PALB2-7R was never acetylated and therefore did not need to be deacetylated or released from sites of active transcription, then, according to their model, why is this mutant defective for HR (Figure 7—figure supplement 1)? Is this finding consistent with the model presented in Figure 7D?

We apologise for the confusion. In the revised manuscript, we provide additional results showing that ChAM-7R binds to nucleosomes even without acetylation (Figure 2D and E), although its association with transcriptionally active loci in vivo is drastically compromised (Figure 9D). Further, the recruitment of PALB2-7R to laser-induced DNA damage was significantly compromised (Figure 6). Collectively, we propose that PALB2 7R is randomly ‘glued’ to chromatin via electrostatic interaction, making this variant less efficiently mobilised and hence less efficiently recruited to sites of DNA damage.

D) On p. 27 (Discussion), the authors state, "Interestingly, the PALB2 7R variant, unlike its 7Q counterpart, had little impact on RAD51 foci formation … This could be explained by our findings that DNA damage triggers ChAM deacetylation and promotes PALB2 mobilisation, which we propose is the critical event in initiating HR repair". This statement seems to overlook data shown in Figure 7—figure supplement 1 that claims PALB2-7R appears is deficient for HR. Further, proficiency for RAD51 foci in response to IR typically is associated with functional HR, but 7R has nearly normal RAD51 foci while suggested to be deficient for HR. Can the authors' explain this based upon their model?

As stated above, we reassessed IR-induced RAD51 foci formation in S phase, when HR is considered most active (Figure 7B and E). This assessment revealed that both PALB2-7Q and -7R are defective in efficient RAD51 foci formation, indicating impaired HR proficiency. We revised the manuscript accordingly.

3. In the model presented in Figure 7D, the authors suggest (1st paragraph of the Discussion, on p. 24) that DNA damage triggers ChAM deacetylation (presumably on PALB2 localized to transcriptionally active chromatin) which "releases PALB2 from active genes and increases its mobility". I have the following concerns about the model (in addition to those detailed in point #2):

A) When discussing their model, it is currently unclear whether the authors are proposing that ChAM deactylation is occurring specifically at "transcriptionally active chromatin" in response to DNA damage and whether this makes PALB2 "globally" available for HR.

These are correct – we thank the reviewer for rewording. In support of this idea, our new ChIP-qPCR assessing PALB2 occupancy at transcriptionally active loci revealed that it is indeed reduced upon IR (Figure 9D). We made this clear in Descussion in the revised manuscript (page 29).

B) The data shown in Figures 4B-C seems to contradict the authors' model (Figure 7D). If the role of ChAM deacetylation is simply to release "PALB2 from active genes and increase(s) its mobility", then the 7Q mutant (mimic of acetylated lysine), by being more mobile (Figure 7D-E), should be available for "HR repair complexes" that form in chromatin. Because the 7Q mutant shows less association with chromatin than WT PALB2 or the 7R mutant (Figures 7B-C), one wonders if the 7Q mutant impairs some other PALB2 function besides release from transcriptionally active chromatin upon DNA damage. Additionally, the 7R (non-acetylable) mutant appears functional as indicated by normal cellular resistance to Olaparib (Figure 7C). This does not support any claim there is a specific need that PALB2 be recruited to transcriptionally active chromatin and subsequently released.

The reviewer is absolutely correct, except that we consider that the 7Q is functionally null in direct chromatin association rather than mimicking acetylated lysine. Indeed, our new experimental results indicate that, while PALB2-7Q can be recruited to sites of laser-induced DNA damage (Figure 6), it fails to promote RAD51 foci formation (Figure 7B and E). It was recently shown that PALB2, through its basic residues within the ChAM, can bind to the acidic surface of nucleosomes at sites of DNA damage (Belotserkovskaya et al., Nat Comms, 2020). We speculate that this engagement allows optimal loading of RAD51 onto ssDNA for HR repair. Our results also support the notion that PALB2 7R is defective in HR repair, albeit unlikely at actively transcribed genes (see our discussion on the HR reporter and sensitivity to olaparib above).

The need for steady-state PALB2 association at transcribed genes was fully evaluated in our previous publication (Bleuyard et al., PNAS, 2017). When PALB2 association at active genes is compromised, these regions accumulate DNA damage when exposed to the DNA topoisomerase I poison camptothecin (CPT). It remains unclear why PALB2 needs to be released upon DNA damage, but the copy number of PALB2 is estimated at 10 times lower than that of BRCA2 (Kulak et al., Nat Methods, 2014), and we speculate that random and static PALB2 association with chromatin, as seen with the PALB2 7R mutant, is toxic to the cells.

C) Unless I overlooked it, it seems that the authors did not specifically test the effect of the 7R mutant on the association of PALB2 (or the ChAM domain) with nucleosomes; this may be helpful in better interpreting various results in the context of their model.

We included the biochemical assessment of the 7R ChAM binding to nucleosomes in the revised manuscript (Figure 2D and E). The purified recombinant ChAM 7R, along with WT and the 7Q, was assessed for its binding to nucleosomes in vitro. Our results show that ChAM 7R indeed maintains nucleosome binding, albeit at a slightly reduced level compared to the non-acetylated WT. Conversely, the 7Q counterpart largely lost its binding to nucleosomes.

D) Since the 7R mutant shows even less association with transcriptionally active chromatin (Figure 5F) than PALB2-7Q, and PALB2-7R is resistant to Olaparib (Figure 7C), then it would seem that protection of "transcriptionally active chromatin … during DNA replication (2nd paragraph of p. 24) is not essential for cell survival in response to DNA damage.

We appreciate this critique. As discussed above, we have previously shown that PALB2 association at active genes is important to protect these regions from transcription-replication conflict, which is enhanced by CPT (Bleuyard et al., PNAS, 2017). CPT covalently links TOPI to DNA, which is significantly different from noncovalent PARP-trapping by olaparib (Murai et al., Cancer Res, 2012; Langelier et al., Science, 2012). Unlike TOPI-trapped lesions, the absence of PALB2 at active genes appears to cope well with PARP-trapped lesions, likely due to its more dynamic nature of trapping. Nonetheless, we removed data associated with olaparib treatment to make this manuscript better focused on thier response to randomly induced DNA damage.

E) The model that acetylation of the 7K patch releases PALB2 from transcriptionally active chromatin would be better supported by testing this in the presence of IR (or Olaparib), since DSB-initiated HR and cellular resistance to Olaparib are among DNA damage response-related assays featured in the manuscript. Does IR or Olaparib indeed release WT PALB2 from transcriptionally-active loci?

We have reconducted PALB2 ChIP in unperturbed conditions and also in IR-treated cells (Figure 9D). We observe clear enrichment of WT PALB2 at ACTB, TCOF and WEE1 loci in unperturbed cells, but this association is greatly reduced in IR treated cells. This observation is consistent with our FRAP results showing increased PALB2 mobility in IR-treated cells (Figure 4D and E). Both the 7Q and 7Q mutants showed reduced association with these loci in unperturbed conditions, and little change of their association upon IR-treatment.

F) According to the model presented, are transcriptionally-active loci left deprotected after acetylation of the ChAM motif that results in increased mobilization of PALB2 for HR?

It is widely known that, upon DNA damage, cells suppress overall progression of DNA replication via activation of checkpoints. Hence, we consider that these regions have a reduced risk of generating DNA damage arising from the transcription-replication conflict (Bleuyard et al., PNAS, 2017) while PALB2 acts at damaged DNA for repair.

G) In the absence of assays of PALB2 with transcriptionally active loci after exposure to IR (or PARP inhibitor), the authors have not convincingly demonstrated that PALB2 is mobilized from these loci after DNA damage. While the authors note that Bleuyard et al. 2017b demonstrate that CPT induces decreased association of PALB2 with transcriptionally active chromatin, CPT could have a different effect than IR by inducing more replication stress.

As indicated under (E) above, we now include PALB2 ChIP at transcriptionally active loci in IR-treated cells (Figure 9D). It is indeed efficiently released from these loci upon IR.

4. The authors note that both K to R and K to Q substitutions alter the size of the side chain, albeit to different degrees. How confident are the authors that K to R and K to Q substitutions are accurate non-acetyl and acetyl mimics, respectively? Is it possible that both substitutions are simply disrupting the structure of the ChAM, and perhaps PALB2 more globally, to some degree?

This is a matter of our active investigation, characterising experimentally the structure of ChAM in the complex with a nucleosome. Nonetheless, in this manuscript, we included AlphaFold2 modelling of ChAM WT (7K), 7Q and 7R mutants (Figure 2—figure supplement 2). It predicts that WT (7K) forms various structural configurations, while 7Q and 7R mutants form consistent structural variations. From this modelling, we infer that WT ChAM has a great flexibility, but that the 7Q and 7Q mutations confer rigidity. This might potentially restrict the way the 7Q and 7R mutants bind nucleosomes, preventing optimum interaction.

5. In contrast to association of PALB2 with transcriptionally active chromatin regulating interaction with BRCA1, as proposed, structural changes across the ChAM, or in PALB2 more broadly, could potentially cause the observed decreased interactions of the 7R and 7Q PALB2 mutants with BRCA1.

As commented above (point 4), we now include AlphaFold2 modelling of ChAM WT (7K), 7Q and 7R mutants (Figure 2—figure supplement 2). The region linking the ChAM and the N-terminal BRCA1 binding domain is largely disordered, and the ChAM mutation is unlikely to affect BRCA1 binding. Indeed, our reassessment of PALB2BRCA1 interaction revealed no impact of the mutations in BRCA1 binding.

Reviewer #3 (Recommendations for the authors):

Fournier et al. – KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrityThe authors have responded to the comments made by each reviewer, and have made several amendments to the manuscript itself, generally improving its legibility and providing additional supporting data / experiments. Saying this, there are still some outstanding questions that should be addressed:

A) It is still not clear as to the (intended) purpose of the GST-ChAM DNA-binding experiments and what these actually contribute to the manuscript as a whole. They could be readily removed from the manuscript without impacting the set of conclusions.

As recommended, we removed DNA binding assays from the current manuscript. As such, we conducted no further investigation to address the reviewer’s comments below.

Note: there is no appreciable DNA binding by any of the mutant forms of the GST-ChAM construct as compared to the two controls, WT or indeed PALB2-FL (how was this protein produced, is this the bacterially expressed protein?).

It is therefore not clear how the authors can support the following statement.

Page 18, Lines 444-446: "… however, the ChAM 3R4K variant also showed impaired binding to DNA in an EMSA assay, albeit less pronounced than that of its K to Q (3Q4K) counterpart (Figure 2H)".

B) Whilst it is appreciated that the authors have now created two additional GST-ChAM construct (3Q4K and 3R4K) they haven't performed the requested control experiment, i.e., with the 7R construct. This is necessary to support the set of experiments described at a later point in the manuscript. What is the effect of the 7R construct in the GST-pulldown experiments – does this still bind to nucleosomes / histone H3?

We now include 7R ChAM in the nucleosome binding assay (see also our response to reviewer 1, point 3, C), presented in Figure 2D and E. Our results show that ChAM 7R indeed maintained nucleosome binding, albeit at a slightly reduced level compared to the non-acetylated WT, while the 7Q mutant largely lost its binding to nucleosomes.

C) Figure 3F, why is there a such a significant increase in PALB2 mobility in the FRAP experiment for the negative control siCntrl? See also Figure 3—figure supplement 1

We speculate that siRNA treatment itself confers stress to the cells. Indeed, we consider siCntrl is a better control for siKAT2A/B and, as such, removed the non-treated data from the plot (Figure 3D)

D) The data for MMC/Olaparib treatment presented in the response to authors (Figure R3) should be included as supplementary data within the revised manuscript.

Page 23, Lines 572-576

"… though similar changes in ChAM acetylation were not detectable upon MMC or olaparib DNA damaging treatment. We noted that IR triggered strong γ- H2A.X induction (Figure 6B), which was not observed upon exposure of HEK293T cells to MMC or olaparib, suggesting that the reduction of ChAM acetylation reflects the cellular response to DNA damage.

In the revised manuscript, we focused on the cellular response to IR. As such, we removed the data associated with MMC and olaparib.

E) Figure 7 —figure supplement 1, panel B (HR assay)

The means from individual experiments should be displayed on the graph, as it is not clear if the wide error bar (which presumably should also extend downwards, as + and – 1 SD) and thus interpretation of the result is due to a single outlier measurement.

We repeated the HR reporter assay, targeting AAVS and LMNA loci. As extensively explained in our response to reviewer 1, point 2, (A), we were unable to gain consistent results using these systems, but would also like to highlight several limitations of the HR reporter. As such, we removed the result from the revised manuscript.

In its place, as reviewer 1 also comments in point 2 (D) (i.e. ‘proficiency for RAD51 foci in response to IR typically is associated with functional HR’), RAD51 foci formation was used as a reliable readout of HR proficiency in this study.

(F) Discussion

The authors allude to unpublished data within the discussion / conclusion – such statements should ideally be removed.

As suggested, we removed the statements below.

“Also, our preliminary results indicate that a PALB2 variant defective in BRCA1 binding exhibits higher accumulation at genic regions than its wild-type counterpart.”

“Indeed, our preliminary results demonstrated that extracellular glucose concentration affect PALB2 dynamics, as reducing glucose level in the growth medium increases PALB2 dynamics.”

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

The manuscript has been significantly improved but there are some remaining issues that need to be addressed, as outlined below:

In the current revision of their manuscript, Fournier et al. have, in general, addressed the concerns that were raised previously. This includes removal of confounding results and addition of new data, such as survival assays, recruitment to sites of DNA damage, and ChIP assays for occupancy at transcriptionally active sites with and without IR, for WT PALB2, and the 7Q and 7R mutants. Overall, the data and presentation are now much more cohesive and are integrated into an interesting model in Figure 9E. As in previous versions, an impressive array of complementary techniques have been employed in this manuscript. This manuscript will make an important contribution to the literature on DNA damage responses.

We thank the editors for taking the time to evaluate our manuscript once again and are glad to hear they found the manuscript significantly improved.

Having said all of this, it should be noted that the manuscript still requires certain clarifications in the text and/or figures. This will not require additional experimentation only text changes and clarifications that would strengthen this manuscript further and improve overall clarity.

We appreciate the remaining points and have revised the manuscript accordingly. Please find our point-by-point response below:

1) There needs to more clarity, about roles in DNA repair/cell survival away from transcriptionally active genes, of the full cellular pool of PALB2 vs the sub-fraction that is specifically associated with transcriptionally active sites. a) Do the authors propose that any PALB2 remains at transcriptionally active sites after the induction of DNA damage elsewhere and if so whether this may provide ongoing protection of active genes and b) whether the authors are proposing that a PALB2-BRCA2-RAD51 complex is mobilized from transcriptionally active genes and whether the PALB2-BRCA2-RAD51 complex exists away from transcriptionally active genes at steady state.

To address the reviewer’s point, we would like to highlight the following three observations. Firstly, the abundance of endogenous PALB2 protein is estimated to be markedly lower than that of other HR factors. For example, Kulak et al., (PMID: 24487582) estimated that there are ~150 molecules of PALB2 per cell, compared to ~10,000 for BRCA2 and ~100,000 for RAD51, in HeLa. Secondly, Xia et al., (PMID: 16793542) reported, on the basis of BRCA2 or PALB2 immunoprecipitation from fractionated cells, that ‘the majority of nuclear structure bound BRCA2 is associated with PALB2’ and that ‘much of the PALB2 is associated with BRCA2.’ Finally, we have found that PALB2 associates with MRG15, a chromodomain protein that recognises an epigenetic marker of active genes (i.e. H3K36me3) with high affinity (Bleuyard et al., PMID: 28673974).

Considering all these observations together, we envision that the majority of endogenous PALB2 is enriched at actively transcribed chromatin in complex with BRCA2, RAD51 and MRG15, although PALB2-free pools of BRCA2 and RAD51 must exist. Upon the induction of DNA damage at random loci, PALB2 at active chromatin is mobilised, and instead becomes enriched at damaged loci, likely through BRCA1-mediated DNA damage recognition. In this context, our experiment assessing cells expressing exogenous PALB2 variants at elevated levels might have limited sensitivity to highlight the physiological impact of the PALB2 mobility as in endogenous conditions, explaining the somewhat subtle cellular phenotypes in our study.

To specifically address point (a), our ChIP-qPCR data (Figure 9D) indicate that PALB2 occupancy at active genes is effectively reduced to null (i.e. comparable to the level in cells complemented with empty vector) following IR exposure, although we cannot totally exclude the possibility that PALB2 bound at other (untested) actively transcribed genes responds differently. Regardless, we consider it unlikely that damage-induced under enrichment of PALB2 at active genes severely impacts the transcription-replication conflict, as DNA damage would trigger the canonical DNA damage checkpoint (i.e. supressing cell cycle progression), hence slowing the progression of DNA replication (i.e. no active movement of the replication machinery). Regarding point (b), we indeed consider that the PALB2-BRCA1-RAD51 complex is mobilised from actively transcribed chromatin at which it is enriched at steady-state.

These points are clarified in the revised manuscript with the following paragraph (page 29):

“Markedly, PALB2 occupancy at active genes, as detected by ChIP-qPCR (Figure 9D), was effectively reduced to null following IR exposure (i.e. comparable to the level in cells complemented with empty vector), though it may represent only the sub-fraction of PALB2 bound to the loci tested. Regardless, these observations raise a fundamental question: why does PALB2 dissociate from active genes with the potential risk of leaving these regions unprotected? Notably, the abundance of endogenous PALB2 protein is estimated to be considerably lower than that of BRCA2 or RAD51, e.g. ~60 times less than BRCA2 and ~600 times less than RAD51 in HeLa cells (Kulak et al., 2014). The majority of PALB2 is also found to be associated with BRCA2 on nuclear structures (Xia et al., 2006) and with MRG15 (Bleuyard et al., 2017b). Hence, we envision that, in endogenous conditions,

PALB2 primarily associates with BRCA2 and RAD51 on actively transcribed loci to protect these regions (though PALB2-free pools of BRCA2 and RAD51 presumably exist), but when DNA damage occurs elsewhere, the same complex needs to be mobilised to promote HR repair. In light of these considerations, our experimental platform assessing cells expressing exogenous PALB2 variants, at higher than endogenous levels, might have somewhat compromised sensitivity to highlight the physiological impact of the PALB2 mobilisation. Either way, we envision that the under-enrichment of PALB2 at active genes is unlikely to confer severe consequences for the transcription-replication conflict, as IR would trigger the global DNA damage checkpoint, slowing replication fork progression (Lajtha et al., 1958; Ord and Stocken, 1958; Watanabe, 1974) while broken DNA is repaired.”

2) The statements about the role of the 7K-patch in HR needs to be softened and clarified how results obtained with the 7R and 7Q variants provide complementary information about PALB2's regulation and function in response to DNA damage. There were reasons for removing data from HR reporter assays, but a reader may consider conclusions about the role of the 7K-patch in HR based only on RAD51 foci to not be fully supported. As such, it is suggested that the authors briefly discuss their HR reporter assay results (in either the Results or Discussion) as data not shown and indicate why these results may not be congruent with those obtained for RAD51 foci. Finally, in the absence of HR reporter assay data, I would suggest toning down the headings in various places to less definitively link the 7K patch to HR repair (line 610; lines 1026-1027; lines 1044-1045. In general, RAD51 focus formation cannot be equalized with HR.

We appreciate this comment and accordingly included the following paragraph in the Discussion (page 30):

“In the meantime, PALB2 recruitment itself does not appear to guarantee RAD51 assembly. Both the 7R and 7Q substitutions elicited significant impairment of RAD51 foci formation (Figures 7 and 8), suggesting that the 7Kpatch promotes optimal RAD51 engagement with broken DNA, conceivably in part through its direct interaction with the nucleosome acidic patch regions exposed at damaged chromatin (Belotserkovskaya et al., 2020). It should be noted, however, that the full impact of the 7R and 7Q mutations on HR repair remains unclear. We endeavoured to assess HR proficiency of these cell lines using our reporter systems, which monitor the HR-mediated integration of a promoterless GFP gene and the resultant expression of fluorescent protein as the readout (Rodrigue et al., 2019; Yata et al., 2012). Through these approaches, significant HR defects were not detected (data not shown). This could potentially be explained by the model that the PALB2 7K-patch is particularly important for HR repair of DSBs arising outside active genes, the detection of which is not straightforward using available techniques. Further investigation using more elaborate strategies, which allow the evaluation of HR repair outside active genes, is warranted to fully appreciate the importance of PALB2 ChAM acetylation in HR repair.”

We have also toned down the headings, which had referred to ‘HR repair’, to a more accurate description (e.g. RAD51 foci formation)

3) A measure of the statistical significance of differences in Figure 4C is needed.

The statistical significance of changes between time 0 and all combined time points (15, 30, 60 and 120 mins), as assessed by the one-way ANOVA, is now reported in Figure 4C.

4) It looks like there is a non-specific band in the blot for PALB2 in Figure 6A. It would be helpful to indicate, with a marker in Figure 6A and/or by the description in the figure legend, which, if any, of the two bands is non-specific there.

We have now included the positions of molecular weight marker in Figure 6A and indicated the correct band for PALB2. The raw scan of the blot is also included in the ‘source data’.

5) The foci are often difficult to see without significant enlargement in Figures 7D and 8D. Perhaps making this figure part larger relative to the overall figure would help with this. Alternatively, the intensities could be enhanced in the images if the current relative size of the figure part is retained.

As suggested, we have enlarged these figure parts and increased the intensity to the best of our capacity.

6) There is a single statement that appears to have been carried forward from the previous version which should be removed (page 22, line 524-525) 'while suppressing non-specific DNA binding'.

We apologise for this error. This sentence has been removed.

https://doi.org/10.7554/eLife.57736.sa2

Article and author information

Author details

  1. Marjorie Fournier

    Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom
    Present address
    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Conceptualization, Resources, Data curation, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    For correspondence
    marjorie.fournier@bioch.ox.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4075-8719
  2. Amélie Rodrigue

    CHU de Québec Research Center, Oncology Division; Department of Molecular Biology, Medical Biochemistry and Pathology, Laval University Cancer Research Center, Québec, Canada
    Contribution
    Formal analysis, Methodology, Validation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9883-7084
  3. Larissa Milano

    CHU de Québec Research Center, Oncology Division; Department of Molecular Biology, Medical Biochemistry and Pathology, Laval University Cancer Research Center, Québec, Canada
    Contribution
    Formal analysis, Validation, Visualization, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  4. Jean-Yves Bleuyard

    Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom
    Contribution
    Resources, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-6727-5362
  5. Anthony M Couturier

    Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom
    Contribution
    Data curation, Formal analysis, Investigation, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1512-9558
  6. Jacob Wall

    Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom
    Contribution
    Data curation, Formal analysis, Visualization, Writing – review and editing
    Competing interests
    No competing interests declared
  7. Jessica Ellins

    Department of Biochemistry, University of Oxford, Oxford, United Kingdom
    Contribution
    Data curation, Formal analysis, Investigation
    Competing interests
    No competing interests declared
  8. Svenja Hester

    1. Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom
    2. Advanced Proteomics Facility, University of Oxford, Oxford, United Kingdom
    Contribution
    Data curation, Investigation, Methodology
    Competing interests
    No competing interests declared
  9. Stephen J Smerdon

    The Francis Crick Institute, London, United Kingdom
    Present address
    Institute of Cancer and Genomic Sciences, University of Birmingham, Edgbaston, United Kingdom
    Contribution
    Conceptualization, Resources, Methodology, Writing – original draft, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-5688-8465
  10. László Tora

    1. Institut de Génétique et de Biologie Moléculaire et Cellulaire, Illkirch, France
    2. Centre National de la Recherche Scientifique, Illkirch, France
    3. Institut National de la Santé et de la Recherche Médicale, Illkirch, France
    4. Université de Strasbourg, Illkirch, France
    Contribution
    Resources, Supervision, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7398-2250
  11. Jean-Yves Masson

    CHU de Québec Research Center, Oncology Division; Department of Molecular Biology, Medical Biochemistry and Pathology, Laval University Cancer Research Center, Québec, Canada
    Contribution
    Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Writing – review and editing
    For correspondence
    Jean-Yves.Masson@crchudequebec.ulaval.ca
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4403-7169
  12. Fumiko Esashi

    Sir William Dunn School of Pathology, University of Oxford, Oxford, United Kingdom
    Contribution
    Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing
    For correspondence
    fumiko.esashi@path.ox.ac.uk
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0902-2364

Funding

Wellcome Trust (101009/Z/13/Z)

  • Fumiko Esashi

H2020 European Research Council (ERC-2013-340551)

  • László Tora

Edward P Abraham Research Fund (RF 260)

  • Fumiko Esashi

Canadian Institutes of Health Research (FDN-388879)

  • Jean-Yves Masson

Medical Research Council (MR/W017601)

  • Fumiko Esashi

Edward P Abraham Research Fund (RF 282)

  • Fumiko Esashi

Canada Research Chairs (Tier I Canada Research Chair in DNA repair and cancer therapeutics)

  • Jean-Yves Masson

Francis Crick Institute

  • Stephen J Smerdon

European Research Council (Advanced Grant ERC-2013–340551 Birtoaction)

  • László Tora

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication. For the purpose of Open Access, the authors have applied a CC BY public copyright license to any Author Accepted Manuscript version arising from this submission.

Acknowledgements

We are grateful to Nicola O’Reilly (Francis Crick Institute Peptide Chemistry Technology Platform) for synthesis of ChAM peptides, Andrew Jefferson, Carina Mónico, Nadia Halidi, Niloufer Irani, Deirdre Kavanagh (Micron Oxford Advanced Bioimaging) as well as Yan Coulombe (CHU de Québec) for assistance with microscopy, Christine Ralf (Sir William Dunn School) for technical support in generating constructs, Michal Maj for assistance with FACS analysis and Chris Norbury (Sir William Dunn School) for critical reading of the manuscript. FE receives an MRC project grant (MR/W017601), was supported by a Wellcome Trust Senior Research Fellowship in Basic Biomedical Science (101009/Z/13/Z) and is thankful for the support from the Edward P Abraham Research Fund (RF 282). JYM is a Tier I Canada Research Chair in DNA repair and cancer therapeutics and is supported by a CIHR foundation grant (FDN-388879). SJS was supported by the Francis Crick Institute which receives its core funding from Cancer Research UK (FC001156), the UK Medical Research Council (FC001156), and the Wellcome Trust (FC001156). LT received funding from the European Research Council (ERC) Advanced Grant (ERC-2013–340551, Birtoaction).

Senior Editor

  1. Jessica K Tyler, Weill Cornell Medicine, United States

Reviewing Editor

  1. Wolf-Dietrich Heyer, University of California, Davis, United States

Publication history

  1. Preprint posted: August 15, 2019 (view preprint)
  2. Received: April 10, 2020
  3. Accepted: October 20, 2022
  4. Accepted Manuscript published: October 21, 2022 (version 1)
  5. Version of Record published: November 17, 2022 (version 2)

Copyright

© 2022, Fournier et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Marjorie Fournier
  2. Amélie Rodrigue
  3. Larissa Milano
  4. Jean-Yves Bleuyard
  5. Anthony M Couturier
  6. Jacob Wall
  7. Jessica Ellins
  8. Svenja Hester
  9. Stephen J Smerdon
  10. László Tora
  11. Jean-Yves Masson
  12. Fumiko Esashi
(2022)
KAT2-mediated acetylation switches the mode of PALB2 chromatin association to safeguard genome integrity
eLife 11:e57736.
https://doi.org/10.7554/eLife.57736
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