Investigating functional phenotypes of PARP inhibitor treatment by advanced flow cytometry

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© Idun Dale Rein, 2022

Series of dissertations submitted to the Faculty of Medicine, University of Oslo

ISBN 978-82-8377-949-3

reproduced or transmitted, in any form or by any means, without permission.

Cover: Hanne Baadsgaard Utigard.

Print production: Reprosentralen, University of Oslo.

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Acknowledgments

The work presented in this thesis was carried out at departments of Radiation Biology and Core Facilities, Institute for Cancer Research, Oslo University Hospital, from March 2013 until August 2021. Financial support from Oslo University Hospital has been greatly appreciated. The PhD-program at the Medical Faculty, University of Oslo has provided excellent courses and facilitated my education.

I would like to thank senior scientist Trond Stokke for being my enthusiastic supervisor, for all the discussions, your open door and never-ending support. You are generous and selfless in your willingness to help others, lending out your curiosity and intelligence.

I would also like to thank all the current members of the Molecular Radiation Biology research group, Sebastian, Kari-Anne, Monica and Heidi, for all the help I have received in my project. We have had many other wonderful group members and FCCF staff throughout the years. I would specifically like to mention, my mentor in providing excellent service with a smile, Kirsti Solberg Landsverk. I still miss our daily contact, your perspectives and top- notch people-skills.

The department of Radiation Biology has provided a welcoming and encouraging scientific environment. In addition, I would like to give my love to my work family,

“Lønningspilsgjengen”, for all the good times (and beers).

I would like to thank my biological family for stimulating my inquisitiveness and

independence during my childhood, and your belief in my abilities. Thank you for your time and support! I am ever grateful for my son, Johannes, the love of my life. His caring soul and smartness brighten my days.

Oslo, August 2021

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Abbreviations

ADP Adenosine diphosphate

aEJ Alternative end-joining

APC/C Anaphase-promoting complex/cyclosome ATM Ataxia telangiectasia mutated

ATMi ATM inhibitor

BER Base excision repair

BIR Break-induced replication

BRCA Breast cancer susceptibility protein CCN Cyclin

CDK Cyclin-dependent kinase

CDT1 Chromatin licensing and DNA replication factor 1 CLL Chronic lymphocytic leukemia

cNHEJ Classical non-homologous end-joining

DDR DNA damage response

DNA Deoxyribonucleic acid

DSB Double-strand break

dsDNA Double-stranded DNA

FA Fanconi anemia

FCM Fluorescence flow cytometry GMNN Geminin DNA replication inhibitor

HR Homologous recombination

ICL Interstrand crosslink

IR Ionizing radiation

LTGC Long-tract gene conversion

MC Mass cytometry

MCL Mantle cell lymphoma

MMR Mismatch excision repair mRNA Messenger ribonucleic acid NER Nucleotide excision repair PAR Poly(ADP-ribose)

PARG Poly(ADP-ribose) glycohydrolase PARP Poly(ADP-ribose) polymerase PARPi(s) PARP inhibitor(s)

PCD Programmed cell death PLK1 Polo-like kinase 1

RB1 Retinoblastoma protein 1 RCD Regulated cell death

RNA Ribonucleic acid

RP Restriction point

SAC Spindle assembly checkpoint

SDSA Synthesis-dependent strand annealing

SSA Single-strand annealing

SSB(R) Single-strand break (repair) ssDNA Single stranded DNA

WT Wild-type

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List of papers

I. Idun Dale Rein, Kirsti Solberg Landsverk, Francesca Micci, Sebastian Patzke, and Trond Stokke.

Replication-induced DNA damage after PARP inhibition causes G2 delay, and cell line-dependent apoptosis, necrosis and multinucleation.

Cell Cycle, 2015, volume 20, issue 14, p3248-60.

II. Idun Dale Rein, Caroline Stokke, Marwa Jalal, June H. Myklebust, Sebastian Patzke, and Trond Stokke

New distinct compartments in the G2 phase of the cell cycle defined by the levels of γH2AX.

Cell Cycle, 2015, volume 20, issue 14, p3261-3269

III. Idun Dale Rein, Heidi Ødegaard Notø, Monica Bostad, Kanutte Huse, and Trond Stokke.

Cell cycle analysis and relevance for single-cell gating in mass cytometry.

Cytometry A, 2020, volume 97, issue 8, p832-844

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Norwegian summary

Utvikling av kreft er forårsaket av endringer i det genetiske materialet som er mer eller mindre unike for hver pasient. En gen-mutasjon som ga kreftcellene en fordel under kreftutvikling kan brukes som mål for terapi, ettersom kroppens normale celler ikke har samme mutasjon. Dette kalles gjerne målrettet behandling eller presisjonsmedisin, og er skreddersydd til kreftpasienten. Et slikt eksempel er behandling av tumorer med BRCA1/2- mutasjon med PARP-inhibitor (PARPi). Normale celler tåler tap av BRCA1/2 eller PARP hver for seg, mens tap av begge fører til celledød (såkalt syntetisk letalitet). Tumorcellene har ikke BRCA1/2 og dør ved behandling med PARPi. Arbeidet som presenteres i denne

avhandlingen startet med å undersøke om PARPi behandling også kan være effektiv

behandling av krefttilfeller der ATM er mutert. Tap av ATM er vanlig i flere krefttyper, bl.a.

mantelcelle-lymfom og kronisk lymfocyttisk leukemi. Både ATM og BRCA1/2 er involvert i cellens respons på DNA skade, og tap av disse øker sjansen for å utvikle kreft.

Selv om det var kjent at PARPi behandling førte til celledød for visse kreftceller var mekanismen ikke kjent. Vi undersøkte derfor effekt av PARPi behandling på proliferasjon, celledødsmekanisme, cellesyklusfordeling og DNA-skade relatert til cellesyklusfase i flere cellelinjer fra lymfom/leukemi. Vi kunne vise at PARPi behandling førte til DNA-skade i S- fase, med påfølgende forlenget G2-fase, noe som ikke tidligere var vist. Mangel på ATM aktivitet økte effekten av PARPi-behandlingen. Behandlingen hindret ikke proliferasjon, og gjentatte runder i cellesyklus økte effekten av PARPi. De ulike cellelinjene møtte ulike skjebner etter behandling, noen cellelinjer døde ved apoptose, mens andre døde etter problemer med å gjennomgå celledeling. Våre undersøkelser av DNA-skade relatert til cellesyklusfase gjorde oss oppmerksomme på en uttalt variasjon i nivåer av DNA-skade (merket med γH2AX) i G2 fase. Årsaken til variasjonen i cellenes oppholdstid i G2 innad i en homogen cellekultur er ikke kjent. Vi ønsket derfor å undersøke hvordan cellenes oppnådde nivå av DNA-skade fra S-fase endrer seg i løpet av G2. Gjennomstrømming av celler fra S- fase inn i og gjennom G2 og mitose ble studert med hensyn på nivået av DNA-skade. Vi utviklet et program for å simulere hvordan celler definert av høyt eller lavt innhold av γH2AX i G2 ble populert før de gikk inn i mitose. Vi fant en variasjon på 2-7 timer for oppholdstid i G2 fase for celler i samme kultur, selv uten behandling. Simuleringen passet godt med at cellene med høyere nivåer av γH2AX hadde lengre oppholdstid i G2. Resultatene våre kan

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derfor tyde på at mengden DNA-skade avgjør lengden på G2-fase. I tillegg tyder våre data på at oppholdstiden i G2 etter at DNA-skaden har blitt reparert er mer konstant (2-3 timer), noe som ikke tidligere har blitt rapportert.

Cellesyklus kan også studerers ved bruk av masse-cytometri (MC), der man benytter isotoper av tungmetaller til deteksjon istedetfor fluorescens som i klassisk flowcytometri (FCM).

Dessverre finnes det ikke en DNA-markør med tilstrekkelig oppløsning til bruk i MC. I fravær av en god DNA-markør er det derimot enklere å kombinere mange markører i samme prøven for MC sammenlignet med FCM. MC er dermed attraktivt fordi man kan måle over 50 markører i samme prøven, noe som tidligere ikke har vært mulig med fluorescens. Vi utviklet et markør-panel til MC slik at alle fasene i cellesyklus ble godt definert: iridium-interkalator (DNA), anti-CDT1 (G1), anti-GMNN (ikke-G1), IdU (S) og pHistoneH3 (mitose).

Cellesyklus-analyse ble likevel utfordrende da det ikke fantes en god metode for å eksludere ikke-single celler i en prolifererende kultur etter MC. For å studere cellesyklus er man helt avhengig av å kunne ekskludere målinger av ikke-single celler. Aggregater av to G1 celler kan for eksempel bli feilidentifisert som celler i G2-fase. Vi utviklet derfor en dataanalyse-sekvens som begynte med FlowSOM-clustering. FlowSOM-algoritmen identifiserte prøve-materiale som vi ønsket å ekskuldere, bl.a. debris og døde celler, men definerte i tillegg

cellesyklusfasene uten unøyaktig, manuell region-setting. Ikke-single celler fra hver cellesyklusfase kunne deretter eksluderes individuelt, siden cellene innad i hver fase har relativt lik cellestørrelse. På denne måten opnådde vi like cellesyklusfraksjoner mellom prøver som ble analysert med både FCM og MC, noe som ikke tidligere var oppnådd. Vi viste at for MC-analyse av prolifererende prøvemateriale bør cellesyklusmarkører brukes for å kunne inkludere større single celler, samt ekskludere ikke-single små celler.

Arbeidet som inngår i denne avhandlingen viser at PARPi-behandling forårsaker celledød i celler uten ATM-aktivitet. PARPi kan være et klinisk behandlingsalternativ der kreftcellene viser mangel på ATM-aktivitet. Vi har vist PARPi sin påvirkning på cellesyklus, at PARPi skaper DNA-skade i S-fase, som kan føre til celledød over tid eller forhindre normal celledeling. Vi har også vist at mengde DNA-skade kan avgjøre lengden på G2-fase, til og med i ubehandlet kultur. Avhandlingen har også ført til utvikling av en robust metode for cellesyklus-analyse og eksklusjon av ikke-single celler ved MC.

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1 Introduction

The development of cancer is a miniature evolutionary process, where multiple cell functions are altered to avoid cell death and increase proliferation. Cancer treatment should be tailored for the individual patient since this process is unique for that patient. Although the traditional treatment options, radiation and/or chemotherapy are effective in many cases, the serious side effects make it necessary to stratify the responders from the non-responders. A tailored therapy approach is important both to save lives and to improve quality of life for cancer survivors. Precision medicine is a strategy to identify and exploit the cancer-specific

alterations unique to each patient. This strategy aims to maximize the effect on the malignant cells and reduce the side effects.

DNA damage occurs thousands of times in each cell of our body, each day. Luckily, most genetic aberrations are non-functional, others are corrected by DNA repair mechanisms, and some might lead to cell death. Loss of DNA repair mechanisms at an early stage in cancer development increases the genomic instability, and allows carcinogenic errors to accumulate.

As a result, many cancer cells have deficiencies in the DNA damage response (DDR)

machinery, while the normal cells in the same patient do not. Genome instability and mutation has been defined as a cancer enabling characteristics in the revised hallmarks of cancer

(Hanahan et al., 2011). Traditional treatment options like radiation and some

chemotherapeutics induce massive DNA damage. The cancer cells are more sensitive to DNA damage both due to lack of repair and due to persistent proliferation in the presence of

damage. However, precision medicine has evolved to exploit this vulnerability by targeting the specific DNA repair deficiencies in each cancer.

Homologous recombination (HR) repair is an error-free pathway to repair the most deleterious DNA damage; DNA double-strand breaks (DSB). Loss of proteins in the HR pathway increases cancer risk, as seen for the hereditary deficiency in the BReast CAncer susceptibility gene 1 and 2 (BRCA1/2). Effective treatment of cancer cells with loss of HR function by the use of Poly(ADP-Ribose) Polymerase (PARP) inhibitors was a major discovery (Bryant et al., 2005, Farmer et al., 2005). This treatment option was very intriguing due to the relatively gentle side effect-profile of PARP inhibitors (Guo et al., 2018). PARP1 is involved in sensing DNA damage and recruiting other DDR-factors. Loss of BRCA1/2 and inhibition of PARP was the first, and is currently the only, treatment-based on the principle of

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The present work covers how PARPi-treatment with or without ATM-activity affected proliferation, cell cycle progression, induction of DNA damage and cell death. To understand cell cycle traverse with endogenous and PARPi-induced DNA damage, we further developed a tool for simulation of the residence times in the phases at the end of the cell cycle.

Additionally, we developed an improved method for cell cycle analysis by mass cytometry, which may be utilized to look at a higher number of markers of interest and to identify all single cell events reliably. The introduction of the thesis will be focused on DNA damage and repair, regulation of the cell cycle and cell death modalities. Synthetic lethality, PARP

function and inhibition will then be further detailed. At the end there is a section dedicated to the development of multiparameter cytometry methods.

1.1 The DNA damage response (DDR)

Subtle changes to the DNA sequence are the most common genetic aberrations, i.e. base substitutions and small deletions or insertions. However, the genome may be rearranged further by translocations as well as amplifications/deletions of large segments or even whole chromosomes. Many genetic aberrations are common in different cancer types, and while some are drivers of malignant transformation, others are byproducts of the genomic

instability. Protection of genomic integrity is essential to prevent cancer development. The genome can suffer damage from both endogenous (e.g. replication and reactive oxygen species) and exogenous factors (e.g. radiation and toxins). Ionizing radiation (IR) is carcinogenic, but is additionally an effective cancer treatment in many cases. High-energy radiation damages the DNA and may thus induce either malignant transformation or cell death. To cope with genomic insults, a complex machinery of DNA damage repair and cell cycle checkpoints has evolved. To prevent harm to the rest of the organism, cell suicide (regulated cell death) may ultimately be induced if the level of damage is too high in a given cell. These pathways make up the DNA damage response (DDR), and they guard and

preserve our genetic information.

1.1.1 Single-strand damage repair

The error rate of replication is as low as one base per 10⁷ incorporated nucleotides in yeast cells. The low error-rate is due to proofreading of the DNA sequence and correction of base substitutions by DNA polymerases (St Charles et al., 2015). Transferred to the human genome, this would yield hundreds of errors during each cell division. However, the DNA mismatch

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excision repair (MMR) further corrects base pair-mismatches and small insertions or deletions caused by replication.

The majority of the DNA damage a cell suffers can be mended by evolutionary-conserved high fidelity DNA excision-repair pathways. The most common point mutation

(C5mCT), is the result of unrepaired depurinated nucleotides. Depurinated nucleotides are normally repaired by base excision repair (BER). Essential factors for BER in mammalian cells are APEX1 (Xanthoudakis et al., 1996), POLB (Matsumoto et al., 1995), XRCC1

(Thompson et al., 1982), and LIG3 (Cappelli et al., 1997). Single strand break repair (SSBR) is sometimes referred to as a separate entity of DNA repair. However, SSB structure is an intermediate during BER, and the machinery implicated in SSBR is the same as in BER. If ribonucleotides are incorporated instead of deoxyribonucleotides, ribonucleotide excision repair will correct the mistake. Distortions of the α-helical structure of dsDNA due to chemically altered bases are recognized and repaired by Nucleotide base excision repair (NER). Base pair-mismatches and small insertions or deletions during replication are largely repaired by the DNA mismatch excision repair (MMR). In the presence of intact

complimentary DNA strands, the repair options are largely BER, NER and MMR. However, the more severe DNA damage, DNA double-strand breaks (DSBs), requires other DNA repair mechanisms.

1.1.2 DNA double-strand break sensing

Extensive force, IR and certain chemotherapeutics may create DNA DSBs. Our genomic information is protected by the robust structure of dsDNA, where each strand is stabilized by its sugar-phosphate backbone. Thus, DSBs are rare events, and is estimated to occur 10-50 times in each nucleated human cell per day (Vilenchik et al., 2003). Identification and stabilization of DSBs are the first steps in the repair process. Seconds after a DSB, H2AX histones around the DNA breakage site become phosphorylated at Serine139 (Rogakou et al., 1998). The H2AX-variant of histone H2A make up 2.5%-25% of the total H2A in mammalian cells (Rogakou et al., 1998). Phosphorylated H2AX (at S139 in humans) is termed γH2AX.

This modification precedes the dynamic remodeling of the chromatin-structure at the site of damage. A recent super-resolution microscopy study revealed that each DSB have a cluster of γH2AX-nanofoci around it, linked by the 3D chromatin structure (Natale et al., 2017).

Chromatin areas with γH2AX facilitate binding and recruitment of DDR-factors (Burgess et

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al., 2014, Natale et al., 2017). However, retention of repair factors at the γH2AX-marked site seems to be the most crucial (Bassing et al., 2004, Celeste et al., 2003). In addition to

promoting local repair activity, a high level of γH2AX-marked damage leads to a global DNA damage response (further described in section 1.2.2) (Bantele et al., 2020).

DNA-PKCS (PRKDC), Ataxia Telangiectasia Mutated (ATM) and Ataxia Telangiectasia and Rad3 related (ATR) are the three members of the family of phosphoinositide 3-kinase related kinases (PIKKs) responsible for phosphorylating S139 on H2AX. While ATR is responsible for formation of γH2AX in response to replication stress (Ward et al., 2001), ATM and/or DNA-PKCS phosphorylate H2AX in response to DSBs (Burma et al., 2001). MCD1 binds to γH2AX and this binding further amplifies the γH2AX signal (Stucki et al., 2005). The MRN complex (MRE11–RAD50–NBS1) is recruited by MCD1 (Uziel et al., 2003) and MRN is involved in ATM activation (Lukas et al., 2004). ATM is one of the key upstream players in the DNA damage response (Blackford et al., 2017), in addition to DNA-PKCS and ATR.

Ataxia telangiectasia (A-T) is an autosomal recessive disorder that predisposes to cancer development and affects the nervous- and immune-system of individuals with inherited mutations in both ATM alleles. ATM is dispensable for the actual repair of most DSBs, as defects in ATM activation leads to about 20% unrepaired breaks in resting cells (Riballo et al., 2004). ATM mediates repair of DSBs with blocked DNA ends (Alvarez-Quilon et al., 2014) and is required for efficient repair of IR-induced DSBs (Goodarzi et al., 2008, Beucher et al., 2009). This may explain why A-T confers one of the most profound radio-sensitivities in humans.

Full activation of ATM requires binding to NBS1 (Horejsi et al., 2003, Uziel et al., 2003, Cerosaletti et al., 2006), autophosphorylation of S1981 and subsequent ATM dimer

dissociation (Bakkenist et al., 2003). ATM phosphorylates and activates CHEK2 (Matsuoka et al., 2000) and TP53 (Chen et al., 2003) among others.

Unlike ATM, ATR is activated specifically by replication stress and not DSBs directly.

However, replication fork collapse produces DSBs too. ATR and its binding partner ATRIP are recruited by Replication protein A (RPA) coated ssDNA (Zou et al., 2003). Thus, γH2AX is not only a mark of DSBs but also stretches of ssDNA. ATM is required for end-

resectioning and subsequent RPA-coated ssDNA, thus ATR activation in S and G2 phase is partly ATM-dependent (Jazayeri et al., 2006). ATR requires activation by TOPBP1 (Kumagai et al., 2006). ATR is an essential protein for dividing cells (Brown et al., 2000), and thereby

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differs from ATM and DNA-PKCS. ATR is the major activator of CHEK1, responsible for induction of several DDR cell cycle checkpoints. The role of Poly(ADP-ribose) polymerase 1 (PARP1) in DNA damage sensing will be discussed in a separate chapter towards the end of the introduction.

Interstrand crosslinks (ICL) are covalent bonds between the two opposite DNA strands.

Repair of these lesions is crucial for faithful transcription and replication. Impaired repair of ICL results in a disease marked by genomic instability and increased risk of cancer

development, Fanconi Anemia (FA) (Hashimoto et al., 2016). ICLs are normally detected during S phase when encountered by the replication machinery, and ICL-repair is regulated by the FA-protein family (Ceccaldi et al., 2016b). ICL repair is challenging as it involves both DNA strands, and involves factors from both SSB and DSB repair pathways.

1.1.3 Repair of double-strand breaks

The different types of DSBs and availability of templates for repair guide the decision of DSB repair pathways. In the context of this thesis, it is of special interest to go further into DSBs that arise during replication of DNA. The repair of classical DSBs (e.g. formed by irradiation) is shown in Figure 2 to give a background.

Non-homologous end-joining repair (NHEJ) is the main pathway of DSB repair in G0/G1

phase (before replication of DNA), but it is additionally active throughout interphase in mammalian cells (Her et al., 2018, Murray et al., 2018). Classical NHEJ (cNHEJ) ligates a DSB with no (or minimal) sequence homology. Unless the DSB ends are perfectly compatible, loss of nucleotides and even translocations can result from cNHEJ. Blunt ends or ends with a short ssDNA overhang quickly bind the abundant Ku70/80 heterodimer. Ku70/80 bound to DNA recruits DNA-PKCS (catalytic subunit), and these three form the DNA-dependent protein kinase (DNA-PK) (Gottlieb et al., 1993, Smith et al., 1999). The DSB ends are tethered together by DNA-PK (DeFazio et al., 2002) and a complex of DNA ligase 4 (LIG4) and XRCC4, in addition XLF or PAXX is needed for scaffolding. If end-processing is needed to ensure the compatibility of the ends before ligation, Artemis, Pol λ, Pol μ, PNKP and TDP1 are recruited to the break site (Figure 2). NHEJ is a fast DSB repair mechanism, acting within minutes after break induction (Okayasu et al., 1993). The choice of DSB repair pathway is thought to be guided through a multifaceted decision tree (Scully et al., 2019). Ku70/80 may be displaced from DSB ends by extensive end-resectioning when sister-chromatids are available as a

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template for error-free repair. ATM directly activates DSB repair proteins and promotes both cNHEJ via TP53BP1(Jowsey et al., 2004), and HR via BRCA1 (Cortez et al., 1999) and CtIP

(Sartori et al., 2007). Rapid, yet transient cNHEJ factor assembly at DSBs precedes, but does not inhibit, the slower and persistent retention of HR factors (Kim et al., 2005).

Figure 2: The major pathways of DNA double‐strand break repair. Reprinted by permission from Springer Nature Customer Service GmbH: Springer Nature NATURE REVIEWS, DNA double‐strand break repair‐pathway choice in somatic mammalian cells (Scully et al., 2019),

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DSB repair by Homologous recombination (HR) is considered an error-free mechanism (Figure 2). The homologous sequence from the sister chromatid is preferred as a guide for repair rather than the homologous chromosome (Kadyk et al., 1992); HR is thus largely restricted to S and G2 phases of the cell cycle (Takata et al., 1998). The MRN complex

(MRE11-RAD50-NBS1) initiates end-resectioning of the DSB, which ultimately leads to long 3’ ssDNA tails covered by RAD51. MRN can directly activate the inactive homodimer of ATM (Lee et al., 2004, Lee et al., 2005). Active ATM recruits CtIP to DSBs (You et al., 2009). CtIP promotes efficient end-resectioning by enhancing the nuclease activity of MRE11

(Sartori et al., 2007). EXO1 (exonuclease 1), BLM (Bloom syndrome helicase) and DNA2 (DNA endonuclease 2) further unwind and digest the 5’ DNA strand. To inhibit further resectioning and to protect the vulnerable ends, RPA complexes bind to the ssDNA (Wold, 1997). The RPA coating must later be substituted by a coating of the recombinase RAD51 (creating a nucleoprotein filament) that can pair to homologous ssDNA. BRCA2 is important for the displacement of RPA in favor of RAD51 in humans (Yang et al., 2005). Partner and localizer of BRCA2 (PALB2) binds to both BRCA2 and BRCA1 and is crucial to connect their functions in HR (Simhadri et al., 2019). ATM facilitates BRCA1 retention at the break site (Khurana et al., 2014). A complex of BRCA1 and BARD1 promotes the pairing of DNA when the nucleoprotein filament searches for and invades the homologous sequence (strand invasion) (Zhao et al., 2017b). A displacement loop (D-loop) is formed when the homologous sequence displaces one of the DNA strands (Figure 2).

Several pathways of HR resolution can proceed from this point. During meiosis, formation of a double cross-shaped structure (a Holiday junction) by capture of the second DNA-end can lead to non-crossover and crossover (recombination) of genetic material from the homologous chromosome. After DNA synthesis of the 3’-invading strand, nascent strand displacement will lead to a non-crossover synthesis-dependent strand annealing (SDSA) (Figure 2). The SDSA pathway resolves the majority of DSBs repaired by HR in somatic mammalian cells. SDSA is considered a conservative pathway. However, gene conversion occurs when there are

differences in the homologous template and the donor template. BLM, RTEL1 and FANCM can disassemble the D-loops before a long nascent strand is formed, and thereby limit the extent of gene conversion (Barber et al., 2008, Xue et al., 2015). If the second end is not captured or the nascent strand does not get displaced, error-prone replicative HR responses will take over. Although the two described modalities Break-induced replication (BIR) and

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Long-tract gene conversion (LTGC) are partially overlapping mechanistically, a few factors are critical for each, at least in bacteria and yeast. Both BIR and LTGC involve extensive nascent strand synthesis through a migrating bubble mechanism leaving long ssDNA tracts vulnerable to mutation and rearrangement (Figure 2). Finally, further DNA synthesis restores both the displaced and the invading strand and several helicases and nucleases are required to resolve the intermediate repair structure.

Single-strand annealing (SSA) is a RAD51-independent DSB repair pathway, which is inherently error-prone. SSA only requires homology of a tandem repeat to anneal the 3’

ssDNA ends, at the cost of deleting the sequence between the homologous repeats. SSA requires extensive end-resectioning and RPA displacement to expose a complementary ssDNA sequence. RAD52 promotes, but is not essential for SSA in mammals (Bennardo et al., 2008). RAD52 has been implicated in alternative end-joining (aEJ) specifically during class switch recombination (Zan et al., 2017). Unlike cNHEJ, aEJ ligates the break if limited RPA displacement reveals microhomology between the 3’ ssDNA ends. The helicase function of POLQ (DNA polymerase θ) can displace RPA and the polymerase function can stabilize the two DNA ends. Although not completely essential for aEJ in mammals, mice lacking Polq suffer genomic instability (Shima et al., 2004). Thus, aEJ is part of the genomic maintenance strategy, despite being error-prone.

Replication forks cannot bypass interstrand crosslinks (ICL), the repair of which involves factors from the SSBR pathway NER and HR, in addition to genes that give rise to Fanconi anemia (FA) when mutated (Hashimoto et al., 2016). One-ended double-strand breaks at the sites of broken or collapsed replication forks comprises a distinctive challenge for the DSB repair system. As there is only a solitary dsDNA-end, there is no partner for end-joining or second-end capture for error-free SDSA. RAD51-mediated one-ended strand invasion results in BIR/LTGC (Figure 2). Additionally, there are RAD51-independent forms of LTGC reported at stalled replication forks (Willis et al., 2014). Loss of CtIP, BRCA1 or 2 increases the frequency of LTGC (Willis et al., 2014). Limited DNA synthesis (a few hundred base pairs) at the single 3’ end may lead to microhomology-binding to another ssDNA. This

microhomology-mediated template switching is a dangerous outcome of one-ended breaks.

Repeated cycles of microhomology-mediated or HR-mediated template-switching, could explain complex breakpoints involving sequences from multiple loci in cancer (Zhang et al., 2013).

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16 1.2 The mammalian cell cycle

The choice of DNA repair pathway relies on the cell cycle phase, and DNA damage may be induced during normal cell cycle traverse. Unperturbed G1 phase and mitotic cells have few γH2AX-foci, as most DSBs are repaired before mitotic entry is allowed. However, DNA replication in normal cells leads to accumulation of γH2AX in S and G2 phase cells by itself.

Treatment with scavengers of reactive oxygen species (ROS) decrease the S- and G2- associated γH2AX levels, thus damage during normal DNA replication is largely caused by ROS (Huang et al., 2006). Cell cycle progression in the presence of damage should be avoided to maintain genome integrity. DNA damage checkpoints have evolved in eukaryotic cells to delay cell cycle progression until repair is completed. These checkpoints will be described after an introduction to the unperturbed mammalian cell cycle.

The cell cycle consists of strictly ordered series of events to ensure cell survival. Interphase includes the two gap phases, G1 and G2, and the intervening S phase (DNA synthesis).

Following interphase, cells in mitosis (M phase) prepare and divide the duplicated genome for the two daughter cells. Mitosis is divided into mitotic sub-phases (Figure 3). Chromosome- condensation takes place in mitotic prophase. Nuclear envelope breakdown in pro-metaphase allow attachment of spindle pole-microtubules to the chromatids. Alignment of the

chromatids in metaphase forms the metaphase plane. The spindle assembly checkpoint (SAC) prevents anaphase onset until all sister chromatids are attached to one microtubule from each spindle pole. During anaphase, the cohesion between the sister-chromatids is degraded, allowing them to be pulled towards each pole. Two separated nuclei form and the chromosomes are decondensed in telophase. Physical cell division, cytokinesis, finally abscises the cell into two daughter cells. The daughter cells can now enter a new round of the cell cycle (G1) or stop cycling and enter a state of quiescence (G0) (Figure 3). Cells in G0 can reenter the cell cycle if stimulated, in opposition to senescent cells that are more irreversibly arrested (He et al., 2017).

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17 1.2.1 Regulation of the unperturbed cell cycle

The order of the cell cycle phases is highly organized and regulated by binding of cyclins (CCNs) to the cyclin dependent kinases (CDKs). The expression and subsequent degradation of the different CCNs control the progression through the cell cycle, while the levels of CDKs are almost constant. CDKs are activated by binding to a CCN subunit (Figure 3). However, other kinases, phosphatases and ubiquitin-ligases further modulate the activity of the CDK- CNN complexes. Mitogenic signals induce cell proliferation by activation of CDK4/6 by binding to D-type cyclins, allowing it to phosphorylate Retinoblastoma protein 1 (RB1). The modification relieves RB1-mediated inhibition of E2F transcription factors, responsible for transcribing S phase-promoting genes like E-type cyclins. CCNE-CDK2 complexes further phosphorylate RB1 and other targets at the so-called restriction point (RP).

Although it takes several hours before DNA replication starts after phosphorylation of RB1 (Figure 3) (Stokke et al., 1993), the cell is now irreversibly committed to traverse the rest of the cell cycle (Pardee, 1974). Binary switches, like the RP, ensure cell cycle progression in an ordered manner, as completion of replication and mitosis is essential. The evolved system of cell cycle control switches has been termed checkpoints. The breaks at the RP are encoded by CDKN2A/B (inhibiting CCND-CDK4/6) or by CDKN1A/B (inhibiting CCNE-CDK2). All of which are activated by anti-proliferative signals and the latter is additionally activated by TP53 in response to DNA damage.

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GMNN is expressed from the onset of S phase and degraded in metaphase. GMNN-

degradation is mediated by the Anaphase-promoting complex (APC/C^CDC20) (Clijsters et al., 2013). GMNN forms a stable complex with CDT1 during S phase. This is a separate

prevention-mechanism towards re-licensing as it keeps any remaining CDT1 molecules from interacting with MCMs. When GMNN is degraded in metaphase, CDT1 is free to license origins again.

E2F-dependent transcription of CCNA2 (in somatic cells) leads to activation of the CCNA- CDK2 complex. CCNA-CDK2 helps complete replication and prevents re-replication by phosphorylation of CDT1 and other licensing factors (Sugimoto et al., 2004). This

phosphorylation marks the licensing factors for protein degradation. CCNA-CDK2 inhibits CCNE-CDK2 and marks it for degradation, ensuring a sequential transition from CCNE to CCNA. Activation of CCNA-CDK2 triggers transcription of and prevents degradation of several mitotic regulatory proteins, like PLK1 and B-type cyclins. The CCNA-CDK2 complex re-localize from the nucleus to the cytoplasm during S/G2 transition. From the cytoplasm CCNA-CDK2 can activate the mitotic kinase PLK1 via BORA (Cascales et al., 2020). B-type cyclins are expressed from late S and peak at the end of G2 phase. CCNB- CDK1 accumulation and activity in the nucleus triggers the onset of mitosis.

CDK1 is the only essential CDK for cell cycle progression in mammalian cells (Santamaria et al., 2007). Although it can bind to all cyclins (in the absence of other CDKs) it normally binds to A-type and B-type cyclins. Premature activation of CDK1 leads to premature onset of mitosis, and is inhibited by kinases WEE1, MYT1 and activated by CDC25A/B/C phosphatases. PLK1, among other kinases, positively regulate CDC25-activation.

Chromosome decatenation (i.e. detangling) is dependent on the activity of DNA

Topoisomerase II (TOP2). A decatenation checkpoint in G2 represses mitotic entry until chromosomes are properly detangled (Bower et al., 2010). A second decatenation checkpoint in mitosis represses the onset of anaphase. TOP2-checkpoints are linked to the strand passage activity of TOP2, directly monitoring the catalytical function of the enzyme (Furniss et al., 2013). Upon mitotic entry, CCNB-CDK1 concentration is high and accumulated at the centrosome. Activated CCNB-CDK1 finally triggers an inner feedback-loop where it enhances its own activity by inhibiting its inhibitors (WEE1 and MYT1) and activating its activators (PLK1 and CDC25A/B/C) (Lindqvist et al., 2009). This signaling cascade ensures that mitotic entry is yet another point-of-no-return, like the RP. Levels of both B-type cyclins

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and GMNN remain high if the spindle assembly checkpoint (SAC) is engaged, but if not they will be marked for degradation in metaphase by APC/C^CDC20. APC/C^CDC20 is kept inactive by the activity of MAD2, until all centrosomes are connected to kinetochore from the opposing spindle poles (Figure 3). Active APC/C^CDC20 helps degrade both the cohesion between sister chromatids and CCNB, leaving the cell free to complete cell division.

Variability in total cell division time, even in monoclonal cell culture and between different cells, has mainly been contributed to the duration of G1 phase (Zetterberg et al., 1985, Stokke et al., 1993, Arata et al., 2019). An unperturbed and rapidly diving cell line may have a mean G1 duration of 11 hours, with a duration range of 7-17 hours (Lezaja et al., 2018). Variation in both S and G2 duration has been shown to be substantial even in unperturbed culture (Larsson et al., 2008a, Larsson et al., 2008b, Araujo et al., 2016, Zerjatke et al., 2017). Genome

duplication is initiated at multiple replication origins. Origins are timed to fire throughout the duration of S phase. Replication forks travel bidirectionally at different speeds from each origin. Lastly, replication forks encounter a variable number of obstacles on their way, like active transcription and DNA damage. The variation in S phase duration from cell to cell is naturally dependent on these factors (Hawkins et al., 2013). Originally, G2 duration was considered to be related to protein synthesis and cell growth. Recent results have shown that regulation of mitotic entry is not dependent on de-novo protein synthesis (Lockhead et al., 2020). In contrast to the high degree of variation in G1, S and G2 phase durations, duration of mitosis has been shown to be short, remarkably constant and uncoupled from total cell cycle time (Araujo et al., 2016). The durations of unperturbed cell cycle phases have additionally been shown to be completely uncoupled from each other (Chao et al., 2019).

1.2.2 Cell cycle checkpoints induced by DNA damage

Checkpoints in G1, intra-S, G2 and in metaphase are activated by insults to the genomic integrity of the cell, i.e. DNA damage (lightning bolts in Figure 3). DNA damage checkpoints enforce an arrest on cell cycle progression, and can further initiate DNA damage repair, senescence or regulated cell death. Unlike the other DNA damage checkpoints, the intra-S checkpoint does not cause a complete arrest. Instead, it actively slows down replication forks, and suppresses origin-firing as well as transcription of genes needed for S phase progression.

Common for all the DNA damage checkpoints is the order of initiation: DNA damage is recognized by Sensor proteins, which relay the signal to Transducers, transducers in turn

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control the Effector proteins (blue box in Figure 3). The checkpoint effectors either directly or indirectly (through transcription) induce cell cycle arrest. Arrest is followed by either DNA repair or a complete termination of cell proliferation (cell death or senescence). The three related transducer kinases, ATM, ATR and DNA-PKCS, are at the heart of this signaling network. They have both common and specific targets and pathways, and in turn regulate each other (Blackford et al., 2017). The canonical pathway for activating the intra-S DNA damage-checkpoint involves sensor TOPBP1, transducer ATR, and effector CHEK1 (Iyer et al., 2017). Replication of damaged DNA can further be avoided before S phase entry by the checkpoint in G1. Although initiation of DDR is distinct from the RP, the G1-checkpoint engages many of the same downstream effectors. DNA damage present at mitosis may lead to unfaithful division of genomic content. Thus, entry into mitosis is refused by the DNA

damage checkpoint in G2. Cells are further protected against DNA damage during the vulnerable state of mitosis at the metaphase to anaphase-transition. This DNA damage- checkpoint induces effectors of SAC. The checkpoints that monitor TOP2-status and thereby proper decatenation of chromosomes in G2 and in mitosis are distinct from both the G2 DNA damage checkpoint and SAC (Lee et al., 2019). The intrinsic ordering of the TOP2 checkpoints in relation to the DDR checkpoints does not seem clear from the current literature.

Checkpoint kinase ATM has many downstream targets, like CHEK2 and TP53 and predominantly responds to DSBs. Phosphorylation by ATM induces the kinase activity of CHEK2. In turn, CHEK2 can further positively regulate TP53. Both ATM and CHEK2 phosphorylate BRCA1 and BRCA2 promoting HR and DDR (Zannini et al., 2014). PALB2 interaction with either BRCA1 or BRCA2 is important for initiation and maintenance of the DDR checkpoint in G2, respectively (Simhadri et al., 2019). The G2 DNA damage checkpoint is enforced by activation of CDK1-inhibiting kinases WEE1 and MYT1, and by repression of the CDK1-activating CDC25-phosphatases. ATR-target CHEK1 promotes HR by

phosphorylation of RAD51 (Sorensen et al., 2005) and BRCA2 (Bahassi et al., 2008). Activated CHEK1 induces several DDR checkpoints (Liu et al., 2000) and slows down

replication (Bartek et al., 2004). Deactivation of DNA damage checkpoints after repair of DNA damage has been completed is primarily achieved by PP2A-mediated dephosphorylation

(Ramos et al., 2019). Deactivation of checkpoints without complete DNA damage repair is termed checkpoint adaptation. This phenomenon is seen in some malignant cells, and is presumed to be related to persistent damage (Waterman et al., 2020).

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ATM, ATR, CHEK1 and CHEK2 can all positively regulate TP53, the central regulator of cell cycle arrest or eventually apoptosis. Highlighting the complexity of the DDR, DNA-PKcs has been shown to activate both CHEK1 and CHEK2 under certain circumstances (Zannini et al., 2014, Buisson et al., 2015). TP53-independent actions of CHEK1 include inhibition of CDC25-phosphatases (Waterman et al., 2020). Although dispensable for the engagement of the G2 checkpoint, TP53 is essential for induction of the G1 checkpoint. Tumor suppressor TP53 is inactivated in 50% of all malignancies, and is called the guardian of the genome. TP53 can induce expression of CDKN1A (also known as p21), an inhibitor of CDK-CCN complexes.

Regulating the balance between repair and cell death in response to DNA damage, TP53 either stimulates transcription of DNA repair-associated or pro-apoptotic genes.

1.3 Cell death

Both proliferation and cell death are strictly regulated to enable development and to preserve tissue homeostasis. Cell death can be induced directly (e.g. by trauma and toxins), or

indirectly (by either extrinsic or intrinsic signaling). Indirect cell death requires enzymatic activity. Cell death is defined as irreversible damage to vital cellular functions, leading to loss of cellular integrity. Cell death modality is still to a large degree classified by morphological features. However, advances in biotechnology now allow for a multitude of cell death pathways to be further elucidated. Among the regulated cell death (RCD) pathways that are recognized today are apoptosis (intrinsic and extrinsic), necroptosis, ferroptosis, pyroptosis, parthanatos and entosis, in addition to mitotic, autophagy-related and immunogenic cell death.

Accidental cell death is the term used to describe instantaneous destruction of vital cell components caused by extreme physical, mechanical or chemical insults. On the other hand, RCD relies on molecular mechanisms; and can thus be altered pharmacologically or

genetically. Programmed cell death (PCD) is now suggested to describe cell death of otherwise healthy cells in normal physiological programs like tissue turnover and

development. Conversely, RCD may occur despite cellular homeostasis and removes cells with damage although local cell numbers are appropriate. RCD is instead a strategy developed to protect against stress. There is a considerable overlap between RCD and PCD pathways mechanistically, and individual RCD pathways show a large degree of interconnectivity

(Galluzzi et al., 2018). An important aspect of RCD is the ability of the dying cell to alert the microenvironment or organism to potential threats. The danger-signals released during RCD are collectively referred to as damage-associated molecular patterns (DAMPs). RCD

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pathways are almost invariably dysregulated during development of cancer. Functional RCD, cell cycle control and DNA repair are all involved in eliminating pre-cancerous cells. As the form of RCD is caused by different cellular stresses, and results in different outcomes for the microenvironment and the organism as a whole it is of interest when studying cancer

therapeutics. Mitotic catastrophe and RCD pathways apoptosis and necrosis/necroptosis will be described in the following sections.

1.3.1 Apoptosis

Morphologically, apoptosis is characterized by cytoplasmic shrinkage, chromatin

condensation, DNA fragmentation and plasma membrane blebbing that eventually results in formation of apoptotic bodies. Appropriate induction of apoptosis is crucial to avoid

malignancies, and can be induced by both external and internal cues. Extrinsic apoptosis is mediated by either death receptor (activation by ligand binding) or dependency receptor (loss of ligand-activation) signaling. Both detect changes to the extracellular microenvironment at the plasma membrane. Intrinsic apoptosis can be initiated by lack of growth factors, DNA damage and different types of stress (endoplasmatic reticulum, replication and ROS accumulation). Caspase (CASP) protein family members are involved in the execution of extrinsic and intrinsic apoptosis. Changes in the relative levels of pro- and anti-apoptotic proteins regulate activation of the intrinsic pathway. Outer mitochondrial membrane permeabilization is the irreversible and critical step for the intrinsic pathway. Pro-apoptotic proteins (e.g. BAX and BAK1) mediate this permeabilization, while anti-apoptotic proteins (e.g. MCL1 and BCL2) mostly bind to and inhibit the function of the pro-apoptotic proteins.

Loss of mitochondrial membrane potential and extrinsic signaling activates different initiator caspases (like CASP8, CASP9 and CASP10) through distinct signaling cascades and protein- complexes. Initiator caspases cleave the prodomain from the executioner pro-caspases and thereby activate them. Active executioner caspases (like CASP3 and CASP7) directly cleave both structural (e.g. lamins and actins) and regulatory proteins (e.g. PARP1 and TP53). DNA fragmentation by DFFB (better known as CAD) is normally inhibited by DFFA (inhibitor of CAD; ICAD). CASP3 cleaves and deactivates DFFA, unleashing the DNA fragmentation activity of DFFB (Sakahira et al., 1998). DNA fragmentation is a consequence of parthanatos and autophagy-dependent cell death (Xu et al., 2010, Nezis et al., 2010), and is thus not

exclusive for classical apoptotic RCD. Apoptotic cells may present phosphatidylserine (PS) on the cell surface to initiate phagocytosis. CASP3 is involved in activating the proteins

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involved in externalization of PS (Suzuki et al., 2016). Final-stage apoptotic cells may lose their plasma membrane integrity in a process often referred to as secondary necrosis.

Recently, it has been shown that apoptosis activate the immune system by presenting immunogens, yet reduce the pro-inflammatory processes attributed to necrotic cell death

(Yatim et al., 2017).

1.3.2 Necrotic cell death modalities

Classically, necrotic cells were defined by the lack of morphologic traits of apoptosis or autophagy (cytoplasmic vacuolization). Instead, necrosis gave rise to cell swelling, loss of membrane integrity and leakage of cytoplasma. Moreover, it was not perceived as a regulated form of cell death. Several forms of RCD (e.g. apoptosis, pyroptosis and ferroptosis) display necrotic morphological features as well. Necrotic morphology thus has limited use in

classification of cell death-mechanism. One form of necrosis can be initiated by both high levels of cytosolic Ca²⁺ and severe oxidative stress, and is called mitochondrial permeability transition (MPT)-driven necrosis. MPT-driven necrosis requires the activity of CYPD,

followed by an abrupt loss to the inner mitochondrial membrane permeability and further loss of the transmembrane potential (Galluzzi et al., 2018). In addition to MPT-driven necrosis, changes to intracellular homeostasis may lead to another form of RCD coined necroptosis.

Necroptosis can be initiated from extracellular cues by specific death receptors or pathogen recognition receptors. Molecularly, MLKL and RIPK3 activity is essential for necroptotic RCD, and are part of the necrosome signaling complex. Necroptosis is antagonized by CASP8 and FADD (Kaiser et al., 2011, Zhao et al., 2017a). Release of DAMPs triggers necroptosis and necroptotic cells again releases more DAMPs, fueling an inflammatory response.

1.3.3 Mitotic catastrophe

Mitotic cell death is not a separate entity of cell death, but a variant of RCD during or after mitotic catastrophe. Mitotic catastrophe is a common denominator for the protective measures against survival of cells after aberrant mitosis. It can be triggered by premature mitosis, extensive DNA damage, failure of the mitotic checkpoints or failure to execute the physical steps of mitosis. Morphologically, it is defined by either macro- or multi-nucleation, and micronuclei may additionally occur. The outcomes of either macronucleation or

multinucleation may point to catastrophe at an early versus late the stage of mitosis

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respectively. Mitotic cell death is normally executed by the intrinsic apoptotic machinery, a variant regulated by the BCL2-family and CASP2 (Lopez-Garcia et al., 2017). In some settings and in the absence of TP53, necrotic cell death can be an outcome of mitotic catastrophe

(Mansilla et al., 2006, Galluzzi et al., 2018).

1.4 Synthetic lethality by inhibition of PARP

Synthetic lethality is cell death resulting from the combined loss of two genes/proteins, when loss of either of the two is not lethal (Figure 1). Synthetic lethality was described in

Drosophila over 75 years ago (Dobzhansky, 1946), and was proposed as a cancer treatment strategy 24 years ago (Hartwell et al., 1997). So far only one treatment based on this principle have reached the clinic (O'Neil et al., 2017): The synthetic lethal interaction between inhibition of Poly(ADP-ribose) polymerase (PARP) and defects in the Homologous recombination (HR) repair pathway. Before going further into the results of this synthetic lethal approach, PARP proteins, and the current state of PARP inhibitors will be described.

1.4.1 Poly(ADP-ribose) polymerase

The PARP family of proteins and PARP-like proteins are present in a wide variety of species, ranging from dsDNA viruses and bacteria to most eukaryotes, yet are not found in S. pombe and S. cervisiae (Otto et al., 2005, Hassa et al., 2006). PARP proteins catalyze the transfer of ADP-ribose (PAR) to target proteins (Chambon et al., 1963). PAR is synthesized from NAD⁺ molecules and releases the by-product nicotinamid (Figure 4) (Okayama et al., 1977). PAR carries a negative charge, and PAR polymers (linear or branched and up to several kDa in size) are bound by glycosylic ribose-ribose links (D'Amours et al., 1999). Since some of the 17 human PARP superfamily members are only capable of mono-ADP-ribosylation, it has been suggested the name should be changed to the ADP-ribose transferases (ARTs) (Ame et al., 2004).

PARP1 is the founding member, and is activated by binding to sites of DNA damage

(Benjamin et al., 1980a, Benjamin et al., 1980b). PARP1 is normally responsible for about 90%

of the cellular production of PAR (Shieh et al., 1998). In addition to PARP1, the polyenzyme PARP2 and the monoenzyme PARP3 are the only other PARP family members activated by DNA damage. Although Parp1 knockout mice are viable (Wang et al., 1995), Parp1 and Parp2 double knockout is embryonically lethal (Menissier de Murcia et al., 2003). Parp1 and

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Parp3 double knockout mice are more sensitive to X-ray irradiation compared to the Parp1 knockout, while Parp3 knockout mice are like WT mice in this respect (Boehler et al., 2011). Lethality at the larval stage in Drosophila melanogaster after depletion of the only PARP-like gene further suggests that PARP is essential for development in many organisms (Miwa et al., 1999, Tulin et al., 2002). The phenotype of Parp1^-/- mice is quite mild, with anti-inflammatory traits (Oliver et al., 1999, Mabley et al., 2001) and increased sensitivity to genotoxic stress (de Murcia et al., 1997). PARP1 poly-ADP-ribosylates (PARylates) over 150 acceptor proteins

(Gibson et al., 2016) and DNA damage enhances this activity (Figure 4) (Ohgushi et al., 1980). Transcription factors such as TP53 (Mendoza-Alvarez et al., 2001, Kanai et al., 2007) and NF- κB (Kraus, 2008), histones and multiple enzymes (Gibson et al., 2016) are among the targets for PARylation by PARP1. Certain DNA-structures has been shown to be targets for PARylation by PARP1 as well (Talhaoui et al., 2016, Gibson et al., 2016). PARP1 has been found to scavenge the genome for DNA damage using dsDNA as “monkey bars”, and is thus present at undamaged DNA as well (Rudolph et al., 2018). An auto-inhibitory conformation prevents NAD⁺ from entering the catalytical site of PARP1 before it binds to damaged DNA (Figure 4) (Langelier et al., 2018).

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(Bonicalzi et al., 2005). However, PARG leaves a single ADP-ribose attached to the target, which subsequently needs to be removed by OARD1 (Sharifi et al., 2013). PAR in

mitochondria is degraded by ARH3 (Niere et al., 2012). Even though PARP1 is expressed in abundance (≈ 0,5 million copies per cell (Yamanaka et al., 1988)), the basal level of PARG activity is much higher than that of PARP. Inhibitors of PARG are currently of interest as a complement to PARP inhibitors, as they seem to exploit the replication stress phenotype of cancer cells (Slade, 2020). A high level of PAR is toxic, and its removal by PARG is essential for embryogenesis (Koh et al., 2004). Hyperactivation of PARP1 can lead to cell death by depleting the cellular energy reservoirs after massive consumption of NAD⁺ and ATP.

Furthermore, accumulation of free PAR and PARylated proteins at mitochondria leading to release of Apoptosis inducing factor (AIF) from mitochondria (Wang et al., 2003).

Incidentally, PARP1 is one of the targets for cleavage by Caspase 3 (CASP3) (Lazebnik et al., 1994). Thus, PARP1 activity has been proposed to act as a cellular rheostat, tipping the DNA damage-outcome from repair to cell death in response to the level of damage. When cell death is initiated by PARP1 hyperactivation, it is termed parthanatos.

The nick-sensor PARP1 is activated by multiple types of DNA lesions, including SSBs, DSBs, incorporated ribonucleotides, DNA crosslinks and stalled replication forks (Figure 5)

(Krishnakumar et al., 2010, Zimmermann et al., 2018). Early on, PARP activity was known to relax the chromatin structure (Poirier et al., 1982). PARP1 recruits and activates chromatin remodelers like ALC1 and MACROH2A1 (De Vos et al., 2012). PARP1 promotes BER/SSBR (Figure 5A) (de Murcia et al., 1997, Dantzer et al., 1999, Fisher et al., 2007), but is not essential for the repair process of SSBs (Vodenicharov et al., 2000, Ström et al., 2011, Ronson et al., 2018). Both PARP1 and PARP2 attract the scaffold protein XRCC1 to sites of damage (El- Khamisy et al., 2003, Fisher et al., 2007, Hanzlikova et al., 2017), by XRCC1 binding to PAR

(Masson et al., 1998). Moreover, PARP1 is known to be a factor in replication fork protection.

PARP1 activity increases the chance of repair by HR at stalled replication forks (Beck et al., 2014b). PARP1 triggers either fork reversal or DSB-repair by HR depending on whether the damage has occurred on the leading or lagging strand respectively (Figure 5C&D). Proper resolution of DNA damage to both the leading and lagging strand involves the HR-pathway in addition to PARP1 (Sugimura et al., 2008, Bryant et al., 2009, Berti et al., 2013, Slade, 2020). Loss of PARP1 results in increased speed of replication forks, and the HR-pathway and CDKN1A have both been implicated in regulating fork speed together with PARP1 (Sugimura