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.

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.

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.

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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

   "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

   "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

   "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

   "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

   "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

    "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

    "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

    "This ORCID iD identifies the author of this article:" 0000-0003-0902-2364

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