The Role of SMARCAD1 During Replication Stress Sarah Joseph

Home , Adermatoglyphia, ASCC3, BTAF1, HLTF, SLFN11, SMARCA1

The role of SMARCAD1 during replication stress

Sarah Joseph

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy under the Executive Committee of the Graduate School of Arts and Sciences

COLUMBIA UNIVERSITY

2020

Sarah Joseph

Abstract

The role of SMARCAD1 during replication stress

Sarah Joseph

Heterozygous mutations in BRCA1 or BRCA2 predispose carriers to an increased risk for breast or ovarian cancer. Both BRCA1 and BRCA2 (BRCA1/2) play an integral role in promoting genomic stability through their respective actions during homologous recombination (HR) mediated repair and stalled replication fork protection from nucleolytic degradation. SMARCAD1 (SD1) is a SWI/SNF chromatin remodeler that has been implicated in promoting long-range end resection and contributes to HR.

Using human cell lines, we show that SMARCAD1 promotes nucleolytic degradation in

BRCA1/2-deficient cells dependent on its chromatin remodeling activity. Moreover,

SMARCAD1 prevents DNA break formation and promotes fork restart at stalled replication forks. These studies identify a new role for SMARCAD1 at the replication fork. In addition to the work presented here, I discuss a method for introducing stop codons (nonsense mutations) into genes using CRISPR-mediated base editing, called iSTOP, and provide an online resource for accessing the sequence of iSTOP sgRNASs

(sgSTOPs) for five base editor variants (VQR-BE3, EQR-BE3, VRER-BE3, SaBE3, and

SaKKH-BE3) in humans and over 3 million targetable gene coordinates for eight eukaryotic species. Ultimately, with improvements to CRISPR base editors this method can help model and study nonsense mutations in human disease.

Table of Contents

List of Figures ...... v

Acknowledgments ...... vii

Dedication ...... x

Chapter 1: Introduction ...... 1

1.1 DNA replication and the replication stress response ...... 1

1.1.1 Cell division and the replication machinery ...... 1

1.1.2 Extrinsic and intrinsic obstacles to fork progression ...... 2

1.1.3 The replication stress response ...... 3

1.2 Maintenance of replication fork integrity ...... 4

1.2.1 Fork stalling ...... 4

1.2.2 Fork collapse ...... 15

1.2.3 The role of chromatin in replication fork maintenance ...... 17

1.2.4 The role of SNF2-family remodelers at the fork ...... 19

1.2.5 Methodologies to study replication fork transactions ...... 23

1.2.6 Genomic consequences ...... 26

1.2.7 Associated diseases ...... 28

1.4 The roles of BRCA1 and BRCA2 during double-strand break repair and fork

protection ...... 31

1.4.1 Sources of double-strand breaks ...... 31

1.4.2 The tumor suppressors BRCA1 and BRCA2...... 33

1.4.3 BRCA1/2 and the repair of double-strand breaks ...... 34

1.4.4 BRCA1/2 and the protection of stalled forks from nucleolytic degradation .... 37

1.5 The role of stalled fork protection and homologous recombination in cancer

predisposition ...... 38

1.6 The role of stalled fork protection and homologous recombination in cancer

therapeutics ...... 40

1.6.1 Chemosensitivity ...... 40

1.6.2 Chemoresistance ...... 43

1.6.3 Immunotherapy ...... 46

1.7 The chromatin remodeler SMARCAD1 ...... 49

1.8 Objectives ...... 52

1.9 Figures ...... 54

Chapter 2: The role of SMARCAD1 in promoting fork degradation in BRCA1 and

BRCA2-deficient cells ...... 66

2.1 Introduction ...... 66

2.2 Results ...... 69

2.4 Methods and Materials ...... 73

2.5 Figures ...... 79

Chapter 3: Discussion ...... 84

3.1 Models for the role of SMARCAD1 at stressed replication forks ...... 84

3.2 Inhibition of SMARCAD1 as a chemotherapeutic ...... 87

3.3 Conclusions and future perspectives ...... 88

3.4 Figures ...... 90

Appendix I: Programmable gene editing by CRISPR-Cas9 ...... 92

Chapter A1: Introduction ...... 92

A1.1 RNA-guided genomic editing by CRISPR-Cas9 ...... 92

A1.1.1 Genome engineering technologies using protein-DNA binding...... 92

A1.1.2 Re-purposing the RNA-guided adaptive response, CRISPR/Cas, for

genome editing...... 94

A1.1.3 The Cas9 effector as a promising molecule for genomic engineering ...... 96

A1.2 CRISPR applications ...... 99

A1.2.1 Modulating homologous recombination for increased targeting efficiency 102

A1.2.2 Programmable base editing ...... 103

A1.3 Figures ...... 107

Chapter A2: Inducing stop codons by CRISPR-mediated base editing ...... 109

A2.1 Overview ...... 109

A2.1.1 CRISPR-mediated base editing efficiently disrupts genes through iSTOP 111

A2.1.4 The iSTOP database ...... 114

A2.2 Discussion and perspectives ...... 114

A2.2.1 Advantages over DSB-mediated disruption strategies ...... 115

A2.2.2 Potential iSTOP applications ...... 116

A2.2.3 Limitations to iSTOP ...... 117

A2.2.4 Prime editing: A new precision gene editing tool ...... 118

iii

A2.2.5 Perspectives ...... 119

A2.3 Figures ...... 123

References ...... 124

List of Figures

Chapter 1

Figure 1. 1 ...... 54

Figure 1. 2 ...... 55

Figure 1. 3 ...... 56

Figure 1. 4 ...... 57

Figure 1. 5 ...... 58

Figure 1. 6 ...... 59

Figure 1. 7 ...... 61

Figure 1. 8 ...... 63

Figure 1. 9 ...... 64

Figure 1. 10 ...... 65

Chapter 2

Figure 2. 1 ...... 79

Figure 2. 2 ...... 80

Figure 2. 3 ...... 81

Figure 2. 4 ...... 83

Chapter 3

Figure 3. 1 ...... 90

Figure 3. 2 ...... 91

Appendix Chapter 1

Figure A1. 1 ...... 107

Figure A1. 2 ...... 108

Appendix Chapter 1

Figure A2. 1 ...... 123

Acknowledgments

A professor once told me that getting a PhD is not just physically demanding but emotionally draining as well. As such, this would not have been possible without the support from several amazing people. First, I would like to thank my mentor, Dr. Alberto

Ciccia, for his advice, support, and scientific discourse. Without his help none of this would have been possible. Alberto provided me with so many learning opportunities and

I am profoundly grateful for his attention to detail and fountain of knowledge. I would also like to thank the current and past members of the lab: Dr. Jen-Wei Huang, Dr.

Angelo Taglialatela, Dr. Pierre Billon, Dr. Silvia Alvarez Nanez, Dr. Giuseppe Leuzzi, Dr.

Raquel Cuella-Martin, Tarun Nambiar, and Samuel Hayward. Specifically, I would like to thank Jen-Wei, Angelo, Silvia, and Giuseppe for teaching me several of the techniques I have used throughout my career. And I want to give a special thank you to Tarun, Sam, and Jen-Wei for helping me keep my sanity all these years by providing fun conversation and career advice. Those times really meant a lot to me. I would also like to thank Pierre and Dr. Eric Bryant for taking a chance on me when I said I could make them a web resource for their database.

I would like to thank the members of my qualifying committee and thesis committee members for their amazing advice, support, and humor: Dr. Rodney

Rothstein, Dr. Lorraine Symington, Dr. Jean Gautier, Dr. Zhiguo Zhang, and Dr. Richard

Baer. In particular, I would like to thank Dr. Richard Baer for attending our lab meetings, providing his expertise, and always having great experimental suggestions. I would, also, like to thank Dr. Chao Lu for his participations in our lab meetings and advice. I

vii would also like to thank the National Science Foundation. Portions of this thesis were funded by the NSF Graduate Research Fellowship DGE-1644869.

My scientific journey would not have been possible without the several professors throughout my life that supported my growth as a scientist and a person.

Therefore, I would like to thank all of those who have mentored me in the past and fostered my curiosity and passion for science. Early on in my life, Dr. Eleanor Sterling taught me to not give up and to find new opportunities. It has been wonderful to be able to reconnect with her at Columbia some 10 years later. I would, also, like to thank Dr.

Kim McKim at Rutgers University for believing in me in high school and taking me on as a student. Additionally, I would like to thank my amazing mentor Dr. Elaine Sia and her lab at the University of Rochester. Words cannot express how helpful, kind, and supportive Dr. Sia has been as a mentor.

I would like to thank my amazing friends for their patience and encouragement. I wish I could thank each of you individually but then this section would honestly be 500- pages long. Please know, that your board game nights, musicals, video games, and gatherings made all of this possible. “Because I knew you, I have been changed for good” (Stephen Schwartz, Wicked).

I would like to thank my family. They have helped me through the injuries, tears, and struggles. I would like to thank my mom for always being there and providing me with new books or documentaries to support each job I had wanted to pursue. From paleontology to conservationist you never said I could not do it (except the subtle hint that I probably should not go to space- 100 ways to die in space…very subtle). You helped me through some of the toughest times of my life and I would not have gotten

viii this far without you. I would also like to thank my mother-in-law Sandra Meltzer for leading me through the toughest parts of my PhD. Of course, I would like to thank my sister Rachel for dealing with me all these years and for listening to my rants. Thank you for joining my science research team, showing me backstage crew, and teaching me about geology (you rock). I would like to thank my sister and brother in-laws Erica and

David for their advice, support, and endless source of fun conversations.

Unquestionably, I would like to thank my grandma Gerry and my bubbe Doris. Thank you bubbe for being such a firecracker and source of joy. And thank you grandma for letting me live in your house for months to do research and helping me become the person I am today. Without your encouragement and help, I would not be where I am today.

Finally, I would like to thank my husband, Aaron. Thank you for encouraging me to become a better scientist and web developer. Without your knowledge and immense patience, it would have taken much longer to understand backend and frontend development (even still I can barely grasp it). Thank you for running up to Columbia to eat food with me, sit next to me while I am at the microscope or in the hood, looking over my figures, and listening to me workshop this acknowledgements page (seriously, this is very hard). From the time we met to now you have always had my back. From surgery to quarantine you know how to cheer me up.

A special thank you to the front-line doctors, nurses, and essential workers risking their lives and keeping me safe during this pandemic.

Dedication

For Monroe Stavenhagen, Harry Rossen, and Kathy Storfer.

“Every individual matters. Every individual has a role to play. Every individual makes a difference. The least I can do is speak out for those who cannot speak for themselves.”

- Dr. Jane Goodall

Chapter 1: Introduction

Portions of this chapter are adapted from:

Joseph SA, et al. Replication fork remodeling, genome stability and human disease:

the role of SNF2-family DNA translocases. In submission.

This chapter was written with help from Angelo Taglialatela, Giuseppe Leuzzi, Raquel

Cuella-Martin, Jen-Wei Huang, and Alberto Ciccia.

1.1 DNA replication and the replication stress response

1.1.1 Cell division and the replication machinery

DNA replication ensures the accurate duplication of the genome during cellular proliferation. Failure to accurately duplicate the genome during DNA replication causes genomic instability, an enabling characteristic for cancer development [1-3]. In order to replicate the genome, a subset of replication initiation sites is licensed in early G1 upon assembly of the pre-replication complex (pre-RC), which is composed of the origin recognition complex (ORC1-6), CDT1, CDC6 and the replicative helicase, MCM2-7, in a head-to-head configuration [4-6]. A subset of MCM2-7 complexes loaded at replication initiation sites become activated in S-phase following the binding of CDC45 and the

GINS complex mediated by the DDK and CDK kinases, resulting in the formation of the

CDC45-MCM-GINS (CMG) complex, which catalyzes the 3’-5’ unwinding of parental

DNA strands [7-11]. Unwinding of the parental strands leads to the formation of single- stranded DNA (ssDNA), which is rapidly coated with the ssDNA-binding RPA trimeric complex (RPA1-3) to prevent promiscuous DNA reannealing[12-15]. DNA synthesis is 1

then initiated by the DNA polymerase α-primase complex and is then continued through the coordinated actions of DNA polymerase ε on the leading strand and DNA polymerase δ on the lagging strand [16, 17]. Due to the antiparallel nature of the DNA strands, the leading strand is synthesized continuously, while the lagging strand is generated discontinuously through the synthesis of Okazaki fragments [18, 19].

Replicative DNA polymerases associate with the proliferating cell nuclear antigen

(PCNA), which facilitates DNA synthesis by increasing the processivity of DNA polymerases [20, 21]. Sites of DNA synthesis, also referred to as replication forks, progress bidirectionally from an initiation site and terminate as the result of the convergence of replication forks moving in opposite directions.

1.1.2 Extrinsic and intrinsic obstacles to fork progression

Replication fork progression is frequently challenged by obstacles leading to replication stress and genomic instability [22]. Replication obstacles can occur naturally or be induced by exogenous agents. Endogenous obstacles include DNA abasic sites, modified DNA bases (e.g., oxidized or methylated bases, DNA crosslinks), DNA breaks, protein-DNA adducts, unusual DNA structures (e.g., hairpins, triplexes, G-quadruplexes, and trinucleotide repeats), replication slow zones (regions AT-rich or poor of replication initiation sites), unbalanced nucleotide pools, and collisions with transcription or non- histone DNA-bound proteins (e.g., cohesin, centromeric proteins, and pre-RCs) [23-25].

Replication impediments can also be caused by DNA lesions induced by exogenous sources. The most common exogenous sources of replication stress are ultraviolet radiation (UV), ionizing radiation (IR) and chemical compounds, such as DNA crosslinking and methylating agents (e.g., cisplatin, mitomycin C, methyl 2

methanesulfonate), topoisomerase inhibitors (e.g., camptothecin) and inhibitors of nucleotide synthesis and DNA polymerase activity (e.g., aphidicolin, hydroxyurea) [26].

These lesions can stall replication fork progression on two levels: (1) by preventing DNA polymerase or helicase movement or (2) causing the replication fork machinery to “run- off” the strand generating a one-ended DNA double-strand break and replication fork collapse [27].

1.1.3 The replication stress response

While replication obstacles on the lagging strand are bypassed by the synthesis of a new Okazaki fragment, impediments on the leading strand may result in the arrest of replication fork progression (Figure 1.1, step I) [28-30]. Small DNA lesions, such as abasic sites, UV lesions, small DNA hairpins, that block the leading strand polymerase are thought to lead to the formation of RPA-coated ssDNA regions possibly resulting from the uncoupling of the CMG helicase and the replicative polymerases [30-35]. More recent studies have now shown that bulky barriers that prevent the unwinding of the parental DNA strands by CMG also lead to the formation of ssDNA, suggesting fork processing by DNA helicases or translocases [36-38]. Formation of RPA-coated ssDNA results in the recruitment of the ATR (ataxia-telangiectasia and Rad3 related) kinase through its interacting partner ATRIP (ATR-interacting protein) [39-41]. ATR is part of the phosphoinositide-3-like kinase kinase (PIKK) family including ATM (ataxia- telangiectasia mutated), DNA-PKcs (DNA-dependent protein kinase), mTOR

(mammalian target of rapamycin), and SMG1 which, excluding mTOR, phosphorylate serine or threonine residues followed by a glutamine residue (S/TQ) on their target proteins [42]. Upon recruitment to ssDNA regions, ATR is activated by TOPBP1 3

(Topoisomerase 2-binding protein 1) in complex with RAD9-RAD1-HUS1 (9-1-1) [43-

47]. ATR phosphorylates the CHK1 kinase and a wide number of DNA replication and repair factors to suppress global origin firing, initiate local dormant origin firing, slow replication fork movement, arrest cell cycle progression, and promote replication fork stability[48-50]. ATR signaling is also required to initiate the restart of stalled replication forks by replication-coupled repair (RCR), which induces the processing of DNA lesions in coordination with the replisome to enable the restart of DNA synthesis once the obstacles have been bypassed or removed [51].

1.2 Maintenance of replication fork integrity

In order to ensure RCR can occur, several pathways have evolved to maintain the integrity of the replication fork upon replication stress. These pathways act at three levels: (1) allowing the immediate bypass of the damage, (2) pausing and eventually resuming progression once the obstacle is cleared, or (3) cleaving and repairing the fork

(Figure 1.1)[52-54]. How cells determine which pathway to take is in part dependent upon PCNA post-translational modification (PTM), the type of lesion stalling the replication fork, and cell cycle regulation [54, 55].

1.2.1 Fork stalling

Fork traverse and repriming

Large DNA-protein crosslinks (DPCs) block DNA synthesis by dramatically slowing helicase movement past the lesion [56, 57]. Exogenous agents such as UV and chemotherapeutics (cisplatin) or endogenous sources, like formaldehyde, generate

DPCs [58-60]. Spartan (SPRTN), a metalloprotease, has been identified to degrade

DPCs and allow bypass. Unlike proteasomal degradation of DPCs, SPRTN does not require poly-ubiquitination to degrade its targets [61]. Therefore, SPRTN activity is highly regulated to avoid nonspecific proteolysis of the replisome components. As such,

SPRTN is initially mono-ubiquitinated to prevent recruitment to chromatin due to the interaction between the attached ubiquitin and SPRTN’s ubiquitin binding domain (UBZ)

[62]. It is still unclear which de-ubiquitnating enzyme is responsible for removing the

PTM, but SPRTN is able to access chromatin only when non-ubiquitinated [62].

Furthermore, SPRTN binding to ssDNA activates SPRTN’s protease activity enabling

DPC degradation [62, 63]. On the other hand, SPRTN’s binding to dsDNA activates its autocatalytic activity to negatively regulate itself [62]. Notably, SPRTN’s activity also depends on nucleotide synthesis, suggesting that SPRTN can be activated on either the leading or lagging strand of the replication fork [61].Finally, SPRTN is cell cycle regulated with its expression peaking in S-phase [62, 64, 65].

Another factor involved with the removal of DPCs is the helicase RTEL1

(regulator of telomere length 1). RTEL1 maintains telomere integrity by unwinding T- loops and G-quadruplex DNA at the telomeres [66]. Recently, RTEL1 has been identified as an accessory protein that allows the CMG helicase to bypass undegraded

DPCs on the leading strand [57]. RTEL1 facilitates CMG bypass of DPCs by unwinding the DNA beyond the lesion, generating ssDNA beyond the adduct [57]. Furthermore,

RTEL1 is required for efficient DPC proteolysis by SPRTN since the absence of RTEL1 reduced the chromatin binding of SPRTN. On the other hand, for lagging strand DPCs,

MCM10 binds transiently to the N-terminal side of the CMG helicase and isomerizes the

helicase into a steric exclusion mode, in which the lagging strand is excluded from the interior of the helicase, enabling the bypass of the legion [67-69].

Similarly to DPCs, interstrand crosslinks (ICLs), the covalent linkage between nucleotides on opposing strands, generated exogenously by chemotherapeutic agents like cisplatin or mitomycin C or endogenously by aldehydes and lipid peroxidation prevents DNA helix unwinding and replication progression [70-72]. In this case, the

CMG helicases can also bypass ICLs by relying upon the coordinated actions of the

BLM, RMI1, FANCD2, FANCM, and FANCM’s interacting partners MHF1, MHF2, and

PCNA [73-76]. FANCM is a translocase that can move along the DNA duplex without

DNA unwinding [77]. The MHF1 and MHF2 (MHF1/2) heterodimer enhances FANCM

DNA binding activity crucial for FANCM traverse through an ICL while the interaction of

FNACM with PCNA through its PCNA-interacting protein (PIP) box also promotes ICL traverse [74, 77]. However, how the interaction between FANCM and PCNA promotes traversal is still unknown [74]. Additionally, FANCM requires the helicase activity of BLM together with the RMI2 for proper ATR-dependent recruitment to stalled replication forks and efficient ICL traverse [75]. Notably, the association of FANCM with the CMG helicase requires FANCD2 and loss of FANCD2 reduces ICL bypass [76]. Subsequently when two converging forks are stalled by an ICL, TRAIP-dependent ubiquitination of the

CMG helicase controls whether the ICL is repaired by the Fanconi Anemia pathway [78] or by direct unhooking by NEIL3 [79, 80]. Ultimately, after a lesion is skipped, primase activity to generate a free 3’-hydroxyl is required for DNA synthesis to resume by the replicative polymerases [81].

PRIMPOL is the mammalian DNA polymerase-primase that has been identified to re-prime the leading strand after a lesion at a stalled replication fork (Figure 1.1, step

III) [81-83]. PRIMPOL is an error-prone member of the Archeo-Eukaryotic family of primases containing a DNA polymerase domain and a DNA primase domain [84].

Because unchecked primase activity can be mutagenic [84, 85], PRIMPOL activity is inherently regulated by its weak affinity for DNA [86]. Consequently, PRIMPOL requires

RPA to be recruited to stalled forks [84, 87]. Additionally, RPA stimulates the primase function of PRIMPOL while PolDIP2 (polymerase δ-interacting protein 2) positively regulates PRIMPOL’s primer extension activity [84, 86, 87]. Notably, PRIMPOL reprimes DNA substrates containing G-quadruplexes and chain terminating nucleosides

[88, 89]. While the primary role of PRIMPOL is repriming after a lesion, PRIMPOL also plays a minor role in incorporating DNA nucleotides opposite the lesion, known as translesion synthesis (TLS) [81].

Translesion Synthesis

PRIMPOL plays a minor role in TLS by specifically only bypassing through 8-oxo- guanine, deoxyuracil, and pyrimidine dimer lesions [81-83, 90]. Unlike, PRIMPOL, the major TLS polymerases are recruited to the lesion by PCNA mono-ubiquitination at lysine 164 (K164) by the ubiquitin-conjugating enzyme (E2) RAD6 and the ubiquitin ligase (E3) RAD18 [91-94]. RAD18 is recruited in the presence of ssDNA and has a preference for forked DNA by its SAP (SAF-A/B, Acinus and PIAS) domain [93-96]. This mono-ubiquitination allows the TLS polymerases to out compete polymerase δ for access to PCNA and promote subsequent DNA synthesis in response to DNA damage

[91]. The main TLS polymerases Pol κ, Pol ι, Pol η and Rev1 belong the Y-family of 7

DNA polymerases along with Pol ζ from the B-family of polymerases (Figure 1.1, step

II) [97-103]. The other TLS polymerases include the X-family polymerases, Pol μ and

Pol λ, and the A-family polymerases, Pol θ and Pol ν that primarily play a role in gap filling for other DNA repair pathways such as base excision repair (BER) and nonhomologous end-joining (NHEJ) [104-113]. As opposed to the replicative polymerases,

TLS polymerases are characterized by low processivity, low efficiency, and low accuracy. TLS polymerases have larger active sites that permits Hoogsteen-base pairing of nucleotides and accommodates damaged or distorted backbones allowing them to incorporate nucleotides opposite a lesion but lack the 3’-5’ exonuclease proof- reading domain, thus making them error prone [114-117]. Most of the Y-family TLS polymerases interact directly with mono-ubiquitinated PCNA through their respective ubiquitin binding domains, ubiquitin binding zinc-finger (UBZ) for Pol κ and Pol η and the helical ubiquitin binding motif (UBM) for Pol ι, with additional interactions to PCNA with non-canonical PCNA interacting motifs [118-120]. REV1, on the other hand, interacts with ubiquitin through its UBM with additional contacts with PCNA through its BRCA1 C- terminus (BRCT) domain and the polymerase assisted domain (PAD) [121, 122].

Generally, TLS requires the sequential action of two groups of polymerases to (1) insert the nucleotide across from the lesion (Pol κ, Pol η, Pol ι) followed by (2) the extension the nascent strand (Pol κ or Pol ζ) (Figure 1.1, step I) [99, 123-128]. REV1 primarily acts as a molecular bridge between PCNA and different TLS polymerases to facilitate the switching from the high-fidelity replicative helicases but can also insert bases across from the lesion (specifically cytosines opposite guanines, bulky N2-guanine adducts,

and abasic sites) [129-132]. Finally, once synthesis has passed the lesion the replicative polymerases can take over to finish DNA replication [132].

Recent studies have challenged the idea of mono-ubiquitinated PCNA as the mediator for TLS recruitment [91, 133-138]. In yeast, Pol ζ and REV1 were activated independently of PCNA ubiquitination when the integrity of the interaction between

PCNA and Pol δ was impaired [91]. Furthermore, mutations in the yeast and human Pol

η UBZ domains did not compromise TLS activity through UV-induced damage [134-

137]. In avian DT40 cells, disrupting REV1 induced more fork stalling than the disruption of PCNA ubiquitination while PCNA ubiquitination was essential for post-replicative gap filling [138]. Moreover, in mouse embryonic fibroblasts (MEFs) about 30% of TLS still occurred although PCNA polyubiquitination was disrupted using a PCNA KR mutant

(Pcnak164R/K164R) [133]. Additionally, Pol η interacts with RAD18 and plays a non- enzymatic role to promote RAD18-mediated PCNA mono-ubiquitination [139]. Finally,

REV1 interacts with ubiquitinated RAD18 to promote RAD18 accumulation to PCNA and subsequent mono-ubiquitination [140].Taken together, this suggests that mono- ubiquitination of PCNA may enhance TLS efficiency but is not necessary for TLS recruitment while REV1 is the main mediator for TLS at a stalled replication fork.

The Interplay between Translesion Synthesis and Homology Directed Repair

Homology directed repair (HDR) is a compensatory pathway to TLS in which cells bypass lesions in an error-free manner using a homologous template [141, 142].

Initially, it was thought that PCNA K63-linked polyubiquitination by HLTF and SHPRH

RING ubiquitin ligase domains mediated the switch between TLS and HDR [143-146].

However, a recent study by Gervai et al. suggested that polyubiquitinated PCNA was 9

unnecessary for efficient HDR [147]. In this study, expression of a PCNA that cannot form polyubiquitin chains (K63R) was able to restore the cell cycle defect of the PCNA

K63R mutants and led to a reduction in translesion synthesis [147]. As such,

SUMOylation of PCNA at K164 by PIAS4 and PIAS1 has been suggested as an alternative mechanism to promote HDR [148]. Consistent with the notion that HDR is compensatory to TLS, POLDIP2, an interactor of Pol ζ, Pol η and REV1 suppresses

HDR and subsequently increases TLS [149]. But the mechanism as to how this occurs is still unknown [149].

Fork reversal and template switching

Nearly 45 years ago it was proposed that stalled replication forks may undergo fork regression to form a structure that will allow error-free bypass through HDR, instead of using an error-prone pathway [150]. 27 years later this structure was identified in vivo by electron microscopy (EM) in response to genotoxic agents [151, 152]. A regressed or reversed fork is formed when the parental strands re-anneal thereby extruding the nascent DNA into a “chicken foot” or four-way structure [37, 150, 151, 153-157]. Fork reversal has been suggested to prevent ssDNA accumulation, allow more time to repair the lesion, re-position the lesion into dsDNA to enable excision repair, or prevent synthesis across SSBs which could result in the formation of deleterious DNA double strand breaks (DSBs) [37, 151, 158]. Furthermore, this structure has been suggested to allow the bypass of the lesion in an error-free manner by allowing the leading strand to use the lagging strand as a template for DNA synthesis [150]. While bypass via the reversed fork has been demonstrated in vitro there has been no in vivo evidence in mammals that bypass occurs on the regressed arm of the fork [51, 53, 159, 160]. 10

While replicating cells have a basal level of reversed forks, exposure to genotoxic stress increases fork reversal due to ATR signaling [37, 151, 161]. Furthermore, rapidly proliferating embryonic stem cells use reversed forks as a way of protecting them from endogenous replication stress [162]. Moreover, fork remodeling is a global response and can occur even at replication forks that are not directly challenged by DNA lesions

[161]. Fork reversal is mediated by the SNF2-family fork translocases HLTF, ZRANB3, and SMARCAL1 along with the recombinase RAD51 and the helicase FBH1 (Figure

1.1, step IV) [37, 153, 154, 156, 163-166].

Surprisingly, RAD51 does not require its strand exchange activity and does not have any fork remodeling activity of its own but has still been demonstrated to catalyze fork reversal in vivo [160, 164]. Therefore, it has been proposed that RAD51 could act like a platform to promote fork reversal [164]. Importantly, the non-enzymatic fork regression function of RAD51 depends upon FBH1 suggesting that FBH1 and RAD51 cooperate to regress forks in vivo [164]. FBH1 is a recruited to replication forks under stressed conditions by its PIP box and AlkB homolog 2 PCNA interaction motif (APIM)

[163, 167, 168]. Notably, FBH1-mediated fork reversal depends on the DNA binding activity of RAD51 [164]. RAD51 requires BRCA2 as a mediator facilitating RAD51 loading [169-173]. However, the fork reversal function of RAD51 does not rely upon

BRCA2 [155, 174]. While RADX has been demonstrated to counteract RAD51 filament assembly on replication forks [175, 176] and RECQ5 has been shown to disrupt RAD51 nucleoprotein filaments assembled on stalled replication forks to prevent fork reversal

[177, 178], there are still open questions about what factors mediate RAD51 loading onto the replication fork and how RAD51 promotes fork reversal. 11

Although the SNF2-family of translocases share fork reversal activity, they play non-redundant functions during replication stress [154, 179, 180]. Notably, only

SMARCAL1 protects telomeres from replication stress and is required for proper telomere replication [179-185]. At the molecular level these remodelers exhibit differences in their respective modes of action. SMARCAL1 associates with the replication fork under unstressed conditions, but becomes enriched during replication stress through its interaction with RPA [154, 155, 186-192]. FBH1, ZRANB3, and HLTF recruitment are PCNA-dependent. ZRANB3 associates to sites of replication stress in a

PCNA-dependent manner through its PIP box and APIM and contains a NPL4 zinc- finger (NZF) motif that binds K63-linked polyubiquitinated PCNA which is promoted by

HLTF and UBC13 [156, 193-196]. Finally, HLTF recruitment has recently been shown to be dependent on PCNA K164-ubiquitination [196], but catalyzes the PCNA K63-linked polyubiquitination that ZRANB3 partially depends on [143, 197]. Along with the different modes of recruitment and activation, the remodelers display distinct substrate specificities. SMARCAL1 primarily acts on fork substrates containing leading strand gaps. Instead both ZRANB3 and HLTF can reverse forks on leading and lagging strand with equal affinity, while HLTF has a strong preference for available 3’-hydroxy groups

[187, 193, 198-201]. ZRANB3 could be preferentially acting upon forks stalled by damaged DNA bases that could be excised through its endonuclease activity [194].

These observations raise the possibility that SMARCAL1, ZRANB3 and HLTF might operate on distinct types of stalled forks induced in response to replication stress.

These structures could arise independently from each other on distinct stalled forks or originate on the same fork during multiple remodeling attempts, especially under 12

conditions where extensive fork remodeling is thought to occur (e.g., BRCA1/2-deficient cells). It has been proposed that, under those circumstances, SMARCAL1, ZRANB3, and HLTF may act cooperatively on their preferred substrates generated on the same fork and that failure at any point in these sequential steps could impair fork reversal

[154, 202]. SMARCAL1 could, for example, generate fork substrates amenable to

HLTF-mediated fork remodeling by pulling the leading strand closer to the fork junction, and HLTF could control ZRANB3-dependent remodeling by regulating the levels of

PCNA polyubiquitination. However, these models remain to be proven and the dynamics between fork remodelers elucidated.

Fork reversal is a double-edged sword. While failure to reverse forks leads to genomic instability by MUS81-mediated cleavage of persistently stalled forks, excessive fork reversal can also lead to DNA damage [175-177, 203-205]. Therefore, fork remodelers need tight regulation to prevent unscheduled fork reversal [151, 175, 176,

203, 205, 206]. Upon DNA binding, SMARCAL1 is negatively regulated by ATR phosphorylation on serine 652 (S652) in between the two lobes of its ATPase domain

[203]. Moreover, RAD52 has recently been demonstrated to prevent SMARCAL1- mediated fork reversal perhaps by competing for the same RPA-binding site [181, 207-

210]. Additionally, HLTF activity at the fork is restrained by the helicase FANCJ [206].

Interestingly, RPA has been demonstrated to impair ZRANB3 fork remodeling activity in vitro, although ZRANB3 does not have any clear RPA binding domains, suggesting

RPA may sterically hinder ZRANB3’s binding to DNA [198]. FBH1 has been demonstrated to negatively regulate RAD51 by promoting RAD51 K58/64 ubiquitination as a part of the SKP1-CUL1-F-box (SCF) complex [163, 211, 212]. Furthermore, the 13

ssDNA-binding protein, RADX, also suppresses RAD51 activity during replication stress by competing with RAD51 for ssDNA binding [175, 176]. In addition to the negative regulators of helicase activity, the EXD2 nuclease counteracts fork reversal by processing stalled forks into a structure refractory for reversal [213]. Similarly, RECQ1 plays an important role by counteracting reversed forks once repair/bypass/pausing is completed allowing fork restart (Figure 1.1, step V) [205]. Subsequently, poly(ADP- ribose)-polymerase 1 (PARP1), has been demonstrated to inhibit RECQ1 and acts like a master switch to prevent premature fork restart [205].

Fork protection

Fork protection is a second mechanism that restricts genomic instability caused by fork reversal. While reversed forks may aid in error-free repair or bypass of a lesion, unprotected nascent DNA extruded from the “chicken foot” structure is vulnerable to excessive degradation by nucleases [154, 155, 165, 174, 214-220]. The reversed arm of the fork is a prime substrate for nuclease degradation by MRE11, WRN/DNA2, and

EXO1 (Figure 1.1, step IV) [153, 154, 165, 221-223]. The endonuclease CtIP has also been demonstrated to have a role in fork resection, however, its role remains controversial [165, 218, 224-226]. In order to prevent degradation, the recombinase

RAD51 forms a nucleoprotein filament on the stalled fork to protect the regressed arm and is stabilized by several factors such as Abro1, WRNIP1, FANCA, FANCD2, BOD1L,

SETD1A, and the RAD51 paralogs (XRCC2 and RAD51C), BRCA1, BRCA2, PALB2, and BARD1 (Figure 1.1, step IV) [214-218, 226-229]. Interestingly, BOD1L and

SETD1A protect cells from DNA2- but not MRE11-mediated resection [218, 228]. But how these factors play a non-redundant role in limiting resection is still unclear. 14

Nuclease recruitment to replication forks depends on the actions of multiple factors including PTIP, PARP1, MLL3/4, CHD4, and SAMHD1 (Figure 1.1, step IV)

[174, 220, 225, 230, 231]. Notably, PP1 is a protein phosphatase that interacts with

RIF1 [232]. This RIF1/PP1 complex dephosphorylates WRN and DNA2 preventing activation and over-resection [222, 223]. Recently the MRE11 interacting protein

(MRNIP) has been identified to prevent MRE11- dependent degradation of the reversed arm of the fork by inhibiting MRE11 exonuclease but not endonuclease activity [233,

234]. Notably, inhibition of MRE11 endonuclease activity by the chemical inhibitor

PFM03 can prevent fork degradation in BRCA2-deficient cells, suggesting that MRE11 uses both its endo- and exonuclease activities to process the stalled replication fork

[233, 235]. Intriguingly, although MRNIP binds to MRE11, inhibition of DNA2-mediated degradation in MRNIP-deficient cells suppresses fork degradation, suggesting that

MRE11 and DNA2 may work together to process stalled forks [233]. On the other hand, it is possible that MRNIP-deficiency generates fork structures that require DNA2- mediated processing [233]. In contrast, the histone methyltransferase, MLL3/4, in association with PTIP, the chromatin remodeler CHD4, and PARP1 play a role in recruiting MRE11 while SAMHD1 stimulates MRE11 exonuclease activity [174, 220,

225, 230, 231, 236].

1.2.2 Fork collapse

Forks irreversibly arrested by DNA lesions undergo MUS81- / SLX4-dependent fork cleavage, which leads to DSB formation and fork collapse (Figure 1.1, step VI)

[165, 237, 238]. MUS81 and SLX4 cleaves the regressed fork structure generating a

DSB that promotes HR-mediated repair [165, 237, 238]. MUS81-depletion prevents 15

DSB formation after fork stalling while loss of either MRE11 or EXO1 also prevents DSB formation, suggesting that MUS81 cleaves replication forks after nucleolytic processing

[165]. Additionally, PARP1 prevents fork collapse by stopping premature fork restart by

RECQ1 [151, 239]. Interestingly MUS81-dependent processing is promoted by the histone methyl transferase EZH2 only in BRCA2-deficient cells suggesting different roles for BRCA1 and BRCA2-mediated protection [240]. A recent study has suggested that WRNIP1 protects replication fork from SLX4 mediated to prevent unnecessary or untimely cleavage of the reversed fork structure [216, 238]. Ultimately, fork processing by MUS81 and SLX4 leads to pathways that repair collapsed forks.

Repair of collapsed forks

After the generation of a DSB either by MUS81-SLX4 processing or replication fork run-off into a SSB, collapsed forks are repaired by an HR-mediated pathway

(Figure 1.1, steps VI-VIII). The HR-mediated fork restart pathway requires the function of RAD51 for strand invasion followed by DNA synthesis aided by the MCM8-9 helicase in complex with MCM8IP (HROB) (Figure 1.1, steps VII-VIII) [241-244]. Additionally, this HR-mediated fork restart pathway relies upon fork convergence and is mediated by the E3 SUMO ligase NSMCE2 [245]. Moreover, unrepaired DSBs that persist into late

G2 and mitosis can be resolved by a RAD52-dependent mutagenic pathway called mitotic DNA synthesis (MiDAS) (Figure 1.1, steps VII-VIII) [241, 246]. In this case,

RAD52 facilitates MiDAS by promoting microhomology mediated strand annealing that is followed by extensive DNA synthesis using a modified replisome containing POLD3

(Figure 1.1, steps VII-VIII)[247, 248]. Notably, a recent study has suggested that in the absence of BRCA1, RNF168-mediated ubiquitination of γH2AX on K13/K15 initiates 16

DNA synthesis by RAD52-POLD3 through the recruitment and activation of the RAD18-

SLF1 complex thereby providing a mechanism for the pathway choice between HR- mediated repair and MiDAS [249]. Interestingly, in yeast a RAD51- and RAD52- dependent pathway called break induced replication (BIR) proceeds through the invasion of the broken end into a homologous template, followed by conservative DNA synthesis until the end of the chromosome [250-258]. While BIR has been extensively studied in yeast, mammalian BIR and the proteins involved are still poorly understood

[257-265]. While MiDAS and the repair of DSBs resulting from oncogene-induced replication stress, or induced during alternative lengthening of telomeres (ALT), have been suggested to be similar to the BIR pathway described in yeast [251, 252, 256,

266], the molecular mechanisms of BIR-like processes in mammals remain to be fully defined.

1.2.3 The role of chromatin in replication fork maintenance

Several studies have now highlighted the importance of chromatin and chromatin bound proteins during replication stress. A study by Feng and colleagues, recently identified that replication fork stalling induces chromatin compaction through the post- translational modification of the histone H2BK33 by the Clr6 deacetylase in the fission yeast Saccharomyces pombe [259]. Additionally, in yeast, failure to generate heterochromatin during replication stress leads to the separation of the CMG helicase from the replicative polymerases and increased the frequency of fork collapse [259].

Similarly, in mammalian cells, replication stress leads to the loss of acetylated histones mediated by ubiquitin-independent proteasomal degradation suggesting that chromatin remodeling occurs in the vicinity of stalled forks [260]. Furthermore, decreased 17

chromatin compaction due to loss of histone H1 leads to an increase in stalled forks caused by transcription-replication conflicts, while loss of the HMGB1 gene leads to faster fork progression [261]. These findings indicate that chromatin conformation is a critical regulator for replication fork dynamics [261]. Interestingly, the histone chaperone

FACT targets the histone variant macroH2A1.2 deposited at sites of replication stress, protecting cells from genomic instability and aberrant senescence [262]. Chromatin remodeling during replication stress is important for mediating the restart of stalled forks. In yeast, chromatin remodeling by the SNF2-family remodelers Chd1 and Isw1 along with RSC and the chromatin modifiers Gcn5 and Set1 enable extensive resection by Exo1 to promote the recruitment of cohesin to stalled replication forks and facilitate fork restart [263]. In mammals, cohesin loading protects stalled replication forks from extensive nucleolytic degradation and promotes the recruitment of fork protection factors: RAD51, BRCA2, and WRNIP1 [219]. On the other hand, improper accumulation of cohesin presents an obstacle for replication fork progression and has been associated with replication fork slowing, fork collapse, and DSB accumulation [219,

264]. Furthermore, SETD1A protects replication forks by promoting histone H3 lysine 4

(H3K4) methylation which enhances the chaperone activity of FANCD2 to prevent degradation suggesting that both histone modification and mobility are important for replication fork protection [228]. Additionally, disrupting global nucleosome deposition by depleting CAF-1 leads to DNA2-mediated fork degradation [196]. Taken together, these studies underscore the importance of replication stress induced chromatin remodeling for preventing genomic instability during replication stress.

1.2.4 The role of SNF2-family remodelers at the fork

The SNF2 family of proteins is structurally characterized by a SWI/SNF2

(switch/sucrose non-fermenting) helicase domain with a bi-lobed RecA-type ATPase configuration, which is present in the yeast protein Snf2 [265]. Functionally, SNF2-family members have been primarily involved in chromatin remodeling. The SNF2 family includes SNF2-like, INO80, RAD54-like, SSO1653-like, RAD5/16-like, and SMARCAL1- like subfamilies [265, 266]. In addition to the previously described fork reversal factors, other members of the SNF2- family have been involved at different stages of the replication stress response. So far, mammalian INO80, ATRX, CHD4 (Mi-2β), CHD1L

(ALC1), SMARCA4 (BRG1), SMARCA2 (BRM), SMARCA5, SMARCAD1, ERCC6L

(PICH) and BTAF1 (MOT1) have been described to be recruited to or act at stalled replication forks.

The SNF2-like group: SMARCA- and CHD-family members

The SNF2-like group of remodelers contain the SNF2-like helicase domain along with domains that recognize histone tail post-translational modifications (Figure 1.2)

[265, 266]. The CHD-type remodelers typically contain tandem chromodomain enabling them to bind to methylated histones [267]. CHD1L (ALC1) is often included in the CHD- type remodelers despite lacking an identifiable chromodomain and instead containing a macrodomain that binds to PAR-moieties [268, 269]. So far only CHD1L and CHD4 have been attributed roles at the replication fork [225, 270]. SMARCA-type remodelers contain a bromodomain to bind to acetylated histone tails (Figure 1.2) [271]. Of these, so far SMARCA4 (BRG1) and SMARCA2 (BRM) have been demonstrated to be required for efficient DNA replication progression [272]. Given their structural and 19

functional features, the role of CHD- and SMARCA- type factors in replication stress responses implicates histone modification and chromatin remodeling. Loss of either

SMARCA4 or SMARCA2 leads to slower replication fork dynamics and a decrease in inter-origin distance suggesting an increase in origin firing in response to the slower replication fork progression of unperturbed replication forks [272]. Additionally,

SMARCA4 colocalizes with ORC1 and PCNA and has been shown to interact with

TOPBP1, BRCA1, and CHD4 [272-274]. CHD4 can bind methylated histones as well as

PAR-moieties and is recruited to IR-induced DSBs in a PARP-dependent manner [275].

CHD4 was demonstrated to promote nucleolytic degradation in BRCA2-mutant through the control of MRE11 recruitment [225]. Mechanistically, CHD4 may promote chromatin decondensation at sites of replication stress facilitating MRE11 accessibility to DNA

[276]. On the other hand, it has been demonstrated that the chromatin decondensation induced by the chemotherapeutic agent camptothecin is dependent on CHD1L function

[270]. Loss of CHD1L led to slower replication fork speed during replication stress [270].

Since CHD1L is recruited for nucleotide excision repair and IR-induced damage repair dependent on PARP1, it is possible that PARP1 may also recruit CHD1L to sites of replication stress [269, 270, 277].

SMARCA1 and SMARCA5 form the ISWI group of chromatin remodelers and contain HAND, SANT, and SLIDE domains which regulate the specificity of the helicase domain [278] (Figure 1.2). The ISWI family of remodelers associate with several accessory proteins in different complexes to mediate the remodeling of chromatin such as the WICH, NoRC, RSF, CHRAC, NURF, CERF, and ACF complexes [279]. Both

SMARCA1 and SMARCA5 are physically enriched at active replication forks, but only 20

SMARCA5 has been functionally implicated [191, 280]. In particular, SMARCA5 associates with BAZ1B (in the WICH-complex) and PCNA to maintain euchromatin during replication [280]. This suggests that nucleosome remodeling is a crucial for proper replication dynamics and response to replication stress agents.

Unlike the other SNF2 chromatin remodeling sub-families, INO80-type remodelers do not have a specific nucleosome binding motif and their ATPase region contains a large insertion (Figure 1.2) [281]. Ino80, the founding member of this subfamily [282-287]. In yeast, the INO80 complex, in which Ino80 is the catalytic subunit stabilizes stalled replication forks, by a still unknown mechanism [288]. In humans, the

INO80 complex is recruited to replication forks by BAP1’s binding to ubiquitinated H2A during unperturbed DNA synthesis [289]. Furthermore, BAP1 promotes replication fork restart through INO80 and RAD51 recruitment [290].

The SSO1653- and RAD54-like factors involved at the replication fork

ATRX is a RAD54-like factor that forms a histone chaperone complex with DAXX to promote H3.3-mediated heterochromatinization at loci including centromeres, telomeres, CpG-islands, imprinted regions, and endogenous retroviral elements [291-

293]. ATRX has been suggested to protect replication forks from MRE11-dependent degradation through MRE11 sequestering, similarly to its role at telomeres [294, 295].

How ATRX sequesters MRE11 and whether ATRX function is limited to heterochromatin sites still needs to be elucidated. On the other hand, RAD54 can regress and restore stalled replication forks in vitro with a preference towards fork restoration [160]. Interestingly, RAD51 can facilitate both the chromatin remodeling and

branch migration functions of RAD54 as well as promote in vitro fork regression by

RAD54 over fork restoration [160, 296, 297].

Finally, a brief mention to ERCC6L (PICH) and BTAF1 (MOT1) from the

SSO1653 family of helicases (Figure 1.2), notable for their preference for non- chromatin substrates [265]. ERCC6L is a translocase that is recruited to kinetochores to monitor tension and facilitate the resolution of sister chromatids during anaphase [298,

299]. ERCC6L activity is critical for the resolution of ultrafine bridges generated by unresolved recombination intermediates, and sister chromatid bridging due to under- replication or unprotected replication forks [300-302]. Therefore, loss of ERCC6L leads to chromosomal aberrations, micronuclei, and cell death [302]. Although BTAF1 has been previously described in the context of transcription regulation [303, 304], it was also identified by Isolation of Proteins on Nascent DNA (iPOND) to be enriched at collapsed replication forks [191]. Interestingly, a PIP box has been found in the BTAF1 protein and loss of BTAF1 described synthetic lethal with the RPA interacting factor

ETAA1 [191, 305]. Considering the role of ETAA1 in maintaining replication fidelity by facilitating proper chromosome alignment and spindle assembly checkpoint it remains possible that BTAF1 functionally resembles ERCC6L [46, 305]. Alternatively, BTAF1 may be enriched at collapsed forks because of transcription-replication collisions.

Overall, several chromatin remodeling factors are recruited to stalled replication forks, but their respective mechanism of actions need to be determined. These studies underscore the importance of chromatin remodeling to replication fork progression and restart. Indeed, more studies should be done to elucidate the epigenetic and chromatin

contexts around stalled replication forks to gain new insights into replication stress- induced dynamics.

1.2.5 Methodologies to study replication fork transactions

The current gold-standard assay used to study replication fork dynamics in response to replication stress relies on DNA fiber and combing techniques (Figure 1.3)

[306-313]. These techniques utilize thymidine analogs to label the progression of the fork, followed by the stretching of the DNA onto a glass coverslip to obtain DNA fibers with a resolution limit of 1 μm, corresponding to 2-4 kb [307, 312-314]. In addition to having resolution constraints that do not enable visualization of fork structures, DNA fiber/combing techniques are unable to discriminate between leading and lagging strand-dependent DNA synthesis. Unlike DNA fiber techniques, EM enables the visualization and quantification of replication fork structures with a higher resolution around ~30-50 base pairs, thereby permitting the discrimination of ssDNA gaps and reversed forks [37, 157, 315-317]. Despite its high resolution, EM only provides a snapshot of replication intermediates captured at a single time point. Interestingly, EM studies have shown that regressed forks contain nucleosomes, indicating that chromatin remodeling may play an important role in fork remodeling [318]. However, EM-based approaches, and also DNA fiber/combing techniques, lack the ability to discern the influence of different chromatin contexts on replication fork metabolism and visualize proteins associated with distinct replication fork intermediates. Current approaches to detect proteins associated with replication forks rely primarily on the iPOND technology, which enables to pull down and identify proteins bound to nascent DNA by western blotting or mass spectrometry (Figure 1.3) [191] . Although highly sensitive for 23

identifying fork-associated proteins, iPOND technologies do not provide spatial information on the location of the identified factors. Recent studies led to the development of proximity ligation assay (PLA)-based approaches that enable the detection of proteins on nascent DNA by standard microscopy in response to replication stress [154, 319]. To improve the resolution of PLA-based applications, these methods could be coupled with super-resolution imaging technologies, which have been shown to enable the mapping of chromatin-associated proteins and chromatin context around a replicating area [239, 320]. Future developments of super-resolution imaging may also allow the direct visualization of reversed forks and other replication fork transactions, thus enabling the study of replication fork dynamics through live imaging-based analyses. These techniques could be coupled to biochemical assays that employ

Xenopus egg extracts [321-324] or purified replication factors [28, 325, 326] to study replication fork progression with and without fork barriers. Alternative biochemical assays could utilize recombinant proteins to monitor enzymatic reactions on fork-like substrates [166, 187, 193] and define DNA transactions using single-molecule technologies, such as DNA curtains [327-329] or optical tweezers (Figure 1.3) [69, 330,

331]. Such approaches would contribute to determine the influence of DNA repair factors on replication fork transactions.

Standard DNA fiber/combing assays can discern global changes in fork progression but lack the ability to detect alterations caused by specific genomic contexts. To overcome this limitation, DNA combing techniques have been combined with fluorescence in situ hybridization to monitor replication fork progression at single loci using SMARD (single molecule analysis of the replicated DNA) (Figure 1.3) [332- 24

334]. SMARD allows the study of a limited number of genomic loci at a time. More recently, the use of optical mapping technologies have enabled the mapping of fluorescently-labeled replication tracts to genomic sites in human cells and Xenopus extracts [335, 336]. In addition, nanopore sequencing-based approaches, such as D-

NAscent or FORK-seq, have been shown to discriminate thymidine analogue incorporation and map DNA replication sites in yeast and mouse pluripotent stem cells

[337-339]. We expect that further developments of optical mapping and nanopore sequencing technologies will enable genome-wide detection of site-specific replication events under normal or replication stress conditions in human cells.

Classical DNA fiber/combing assays are also unable to specifically analyze replication forks that have encountered DNA lesions, since not detectable, thus rendering difficult the study of DNA lesion bypass or repair events. Recent studies have utilized digoxigenin-labeled psoralen to visualize DNA crosslinks on DNA fibers and study the bypass of those lesions (Figure 1.3) [73]. Similar approaches might be possible using antibodies that detect other DNA lesions (e.g., anti-cisplatin modified

DNA antibody). Alternative methods employing the E.coli Tus-Ter system have been employed in yeast and mammalian cells to study the repair of a single, site-specific replication barrier [340, 341]. More recently, a nuclease-dead Cas9 has been demonstrated to be a programmable barrier for replication fork progression [342].

Combining site-specific fork barriers with PLA and next-generation sequencing may enable the discrimination of proteins recruited to a single stalled fork and the detection of repair events induced by fork blockage. Ultimately, the development of new

techniques to study replication dynamics will provide important insights into the mechanisms that control replication fork reversal and other fork transactions.

1.2.6 Genomic consequences

Failure to resolve replication stress can lead to fork collapse, under-replication, and mutagenesis which can contribute to DNA aberrations, DNA damage, and human disease (Figure 1.4). Dysregulation of ATR signaling contributes to DNA breaks and cellular senescence [343, 344]. As such, inhibition of ATR leads to increased ssDNA and RPA accumulation due to increased origin firing by CDK2 [343]. When this occurs, ssDNA exceeds the accumulation of RPA at the fork leading to fork cleavage by nucleases [343]. Cells that do manage to progress through S-phase, arrested at G2-M leading to cellular senescence [343]. On the other hand, persistent ATR signaling, in the absence of DNA damage, leads to persistent CHK1-depedent cell cycle arrest and p53- dependent senescence [344].

While TLS is potentially mutagenic, issues with TLS leads to decreased cellular survival and replication fork blockage [141, 345-347]. Dysregulation of the TLS polymerases is often associated with unresolved replication stress leading to lingering

DNA damage in G1-cells marked by 53BP1, called 53BP1 nuclear bodies, and increased micronuclei formation (Figure 1.4) [348]. Furthermore, loss of PCNA mono- ubiquitination is associated with aberrant TLS recruitment in undamaged conditions leading to an overall increase in mutagenesis [349]. Moreover, overexpression of

SHPRH or loss of HLTF also increases genomic mutations due to aberrant TLS [146].

However, loss of TLS function is also associated with increased mutagenesis [350].

Therefore, it is critical for genome stability that TLS polymerases are well regulated and 26

used appropriately. Besides SPRTN’s role in DPC removal, SPRTN is also an important regulator for TLS activity [351-353]. Therefore, loss of SPRTN has been demonstrated to lead to an increase in mutagenesis, chromosomal aberrations, cellular senescence, increased sister chromatid exchanges, and sensitization to DNA damaging agents [62,

351-355].

As mentioned before, extensive DNA degradation of the reversed arm of the fork leads to chromosomal aberrations and cell death due to replication fork collapse by

MUS81 (Figure 1.4) [153, 155, 165, 174]. Therefore, loss of the replication fork protectors or unrestrained replication fork reversal increases DNA damage accumulation and chromosomal abnormalities [154, 175, 176, 213-216, 226, 227].

Notably, loss of SAMHD1, a factor that promotes MRE11 exonuclease-mediated resection at stalled forks, leads to the accumulation of cytosolic ssDNA generated by

RECQ1-mediated DNA displacement and MRE11 endonuclease-dependent flap cleavage [230]. Cytosolic DNA is sensed by the DNA the cGAS-STING pathway triggering the expression of interferon-stimulated genes and the activation of the inflammatory response [356-358]. Furthermore, the accumulation of cytosolic DNA depends on MUS81 activity [359]. Taken together, this could suggest that extensive replication fork degradation and fork collapse can lead to an aberrant inflammatory response (Figure 1.4). Overall, the replication stress response needs to be finely tuned in order to prevent genomic instability contributing to human disease.

Failure to continue replication during S-phase leads to under-replication and chromosomal breakage (Figure 1.4) [360, 361]. As such, difficult to replicate regions, such as common fragile sites, telomeres, and centromeres, are associated with sister 27

chromatid bridging [361]. Failure to resolve these bridges leads to cell cycle arrest, mis- segregation, and cell death (Figure 1.4) [301, 360-362]. Furthermore, common fragile sites are associated with hypoacetylated histones and chromatin compaction [363].

Increasing histone acetylation in these areas leads to a decrease in chromosome breakage, suggesting that these areas are susceptible to chromosome break because hypoacetylation can prevent proper replication response [363].Therefore, replication stress and fork restart are critical for preventing aberrant mitosis and senescence.

1.2.7 Associated diseases

Defects in replication fork maintenance proteins are associated with human diseases which contain common features such as developmental defects, premature aging, autoimmune disease, and cancer predisposition. The link between cellular senescence and premature aging is clear in several human diseases. As such dysregulation of ATR signaling, mutations in SPRTN, WRN, and FA proteins that contribute to cellular senescence are associated with human diseases featuring premature aging. Hypomorphic mutations in ATR or mutations in ATRIP are associated with Seckel syndrome characterized by microcephaly, developmental delay, and premature aging [364, 365]. Similarly, Ruijs-Aalfs syndrome is a progeroid syndrome characterized by premature aging, hepatoma development, and skeletal abnormalities caused by mutations in SPRTN [355, 366-368]. Furthermore, mutations in WRN are associated with Werner syndrome which is characterized by pre-mature aging but only by their second decade of life and has a high penetrance of age-related disorders such as type II diabetes and bilateral cataracts [369, 370]. Finally, mutations in Fanconi

Anemia (FA)-proteins are associated with FA which is characterized by skeletal 28

abnormalities, developmental delay, pre-mature aging and an increased cancer predisposition [371]. As expected, FA, Werner Syndrome, and Seckel syndrome patient cells all exhibit senescent features and endogenous replication stress markers like cell cycle arrest and micronuclei [372-374].

Similarly, SAMHD1 and Pol η function are also linked to their human disease.

Mutations in SAMHD1 cause the development of Aicardi-Goutières syndrome which is an inflammatory encephalopathy characterized by the increase in type 1 interferon production due to the enhanced cytosolic ssDNA production [375]. Mutations in Pol η are associated with Xeroderma pigmentosum variant (XP-V) characterized by sensitivity to sunlight [376]. Moreover, mutations in proteins ensuring proper replication through difficult genetic sequences can lead to developmental delays. For example, RTEL1 stabilizes telomeric DNA by unwinding secondary structures like G-quadruplexes and prevents trinucleotide repeat expansion and fork breakage [377, 378]. Therefore,

RTEL1 mutant cells grow slower and display shortened telomeres [379]. Notably,

RTEL1 mutations cause while Hoyeraal Hreidarsson syndrome, a disorder characterized by immunodeficiency and developmental defects [380-383]. Additionally, naturally occurring genomic replication blocks are also associated with neurodevelopmental or neurodegenerative diseases [384]. Fragile X syndrome is characterized by expanded CGG repeats which can form fork impediments leading to chromosome breakage [385, 386].

Mutations in ZRANB3 and SMARCAL1 are, also, associated with human diseases. Biallelic mutations in SMARCAL1 cause the autosomal recessive disorder

Schimke immuno-osseous dysplasia (SIOD), a multisystem disorder characterized by 29

the prominent features of renal failure, T cell immunodeficiency, spondyloepiphyseal dysplasia, growth retardation, facial dysmorphism, hyperpigmented macules, and microcephaly [186, 387-390]. More recently, mutations in ZRANB3 were associated with an African-specific type 2 diabetes implicating ZRANB3 in glucose response and insulin secretion in pancreatic β-cells [391].

Increased mutagenesis and chromosomal rearrangements are hallmarks of cancer. Therefore, mutations in several proteins involved with replication fork bypass and reversal are associated with cancer predisposition. For example, several TLS polymerases are associated with cancer progression [392-396]. Dysregulated expression of Pol κ is associated with colorectal and lung cancers [392-394]. While, overexpression of either Pol κ and Pol ι is associated with gliomas, with upregulation of

Pol κ leading to a poorer prognosis [395]. Finally, overexpression of REV1 is associated with carcinogen-induced intestinal adenomas [396-398].

Mutations in factors that promote fork reversal are also associated with cancer.

HLTF is frequently silenced by promoter methylation in human colorectal and gastric cancers [399, 400]. Loss of Hltf is insufficient to induce cancer in wildtype mice but it significantly increases the formation of colon cancer in an Apc-mutant background

[401]. Paradoxically, HLTF is also commonly amplified and overexpressed in esophageal, uterine and several types of squamous cell carcinoma [402]. HLTF overexpression is significantly associated with increased metastasis and poor prognosis in non-small cell lung cancer patients [403]. However, the putative functional role of

HLTF amplification in cancer has not been experimentally investigated. Similarly, only a few studies have investigated the in vivo functions of SMARCAL1 and ZRANB3 in 30

cancer. ZRANB3 was described as a putative tumor suppressor gene in endometrial cancer [397, 404]. SMARCAL1 mutations have been described as candidate drivers in a subtype of glioblastomas that are wildtype for TERT promoter and IDH1/2 and further enriched in ALT positive tumors [182]. In a T-cell lymphoma mouse model, loss of

Smarcal1 impairs survival of hematopoietic cells and significantly reduces lymphomagenesis induced by gamma irradiation [405]. Similarly loss of either Smarcal1 or Zranb3 increases replication stress and cell death induced by Myc overexpression and reduces Myc-driven B-cell lymphomagenesis [406].

1.4 The roles of BRCA1 and BRCA2 during double-strand break repair and fork protection

1.4.1 Sources of double-strand breaks

Endogenous and exogenous sources can cleave the phosphodiester backbone of the two strands of DNA generating a (DSB). Exogenous sources of DSBs include IR, clastogens (bleomycin and neocarzinostatin), and topoisomerase II-inhibitors

(etoposides) [407]. Since the late1930’s, it has been known that IR can break chromosomes [408]. IR is defined as any subatomic particle that contains enough energy to strip electrons from atoms [407]. IR can generate a DSB directly by colliding with the DNA backbone or indirectly by colliding with water generating a reactive hydroxyl radical that can react with the nearby DNA, thus inducing a SSB [409]. Since a single IR particle can generate clustered lesions, two opposing SSBs near each other cause a DSB [410-412]. Hydrogen peroxide (H2O2), similarly generates SSBs through free radical generation that can cause a DSBs [413]. The number of SSBs generated in

this way saturates at intermediate levels of H2O2 suggesting that there is a limiting factor of chromatin-bound metal ions that can be converted into a free radical [413].

Radiomimetic compounds induce a free radical attack on the DNA backbone [414, 415].

Activated bleomycins contain a reduced transition metal like iron or copper, oxygen, and often H2O2 leading to free radical attack [415]. Alternatively, bicyclic enediynes, like neocarzinostatin, removes hydrogens from the deoxyribose sugar on both DNA strands which can then react with water to generate a break in the sugar backbone [414].

Finally, etoposides generate DSBs by inhibiting topoisomerase II [416, 417].

Topoisomerase II generates a DSB transiently to relieve torsional stress from the unwinding DNA helix [418]. Etoposides prevent the re-ligation step of each topoisomerase II homodimer therefore, DSB formation requires the action of two etoposide molecules binding to each topoisomerase II monomers [419].

Endogenous sources include replication stress, cellular metabolism (H2O2), or scheduled DSB generation by nucleases during V(D)J recombination and meiotic recombination [420]. As discussed above, replication fork collision with SSBs can generate a DSB and replication stress can also result in DSB formation upon reversed fork cleavage. The major source for H2O2 is oxidative phosphorylation occurring in the mitochondria [421, 422]. Finally, antigen receptor gene assembly (V(D)J) recombination) and meiotic recombination use programmed DSB formation in order to generate genetic diversity [423-425].

Failure to or faulty repair of DSBs leads to chromosomal abnormalities, mutagenesis, and cell death [50, 426-428]. To avoid such consequences, the cell has evolved several pathways for repairing DSBs. 32

1.4.2 The tumor suppressors BRCA1 and BRCA2

The tumor suppressors BRCA1 (Breast Cancer Type 1) and BRCA2 (Breast

Cancer Type 2) play an important role to maintain genome stability. Mutations in

BRCA1 and BRCA2 together account for 25% of familial breast cancers [429] and predispose carriers to breast and ovarian cancers with a pattern of autosomal dominant inheritance [430-432]. BRCA1 is a large (220 kDa) nuclear protein containing an N- terminal RING (Really Interesting New Gene) domain, a coiled-coil domain, and a tandem BRCT repeat that makes up the BRCT domain (Figure 1.5A) [433-438]. BRCA1 forms an obligate heterodimer with BARD1 through their respective RING-domains that together make a functional E3 ubiquitin ligase (Figure 1.5A) [437, 439]. The BRCT domain of BRCA1 is a phosophoprotein binding domain allowing BRCA1 to exist in 3 different mutually exclusive complexes: BRCA1-A, BRCA1-B, and BRCA1-C (Figure

1.5A) [440]. The A-complex is composed of the ubiquitin binding protein RAP80, the de-ubiquitinating enzymes BRCC36 and BRCC45, MERIT40 (Mediator of Rap80 interactions and targeting 40), and phosphorylated Abraxas and plays an important role in mediating DNA damage-induced ubiquitin signaling and recruitment of BRCA1 to sites of double strand breaks (DSBs) [441-447]. The B complex consists of TOPBP1 and phosphorylated BACH1 (FANCJ), a helicase that promotes replication stress induced cell cycle checkpoint and interstrand crosslink repair [448-451]. Finally, the C complex involves the MRN (MRE11-RAD50-NBS1) complex and phosphorylated CtIP and is critical for DSB repair by facilitating end resection [446, 452]. In addition to binding phosphoproteins, BRCA1 interacts with RAD51 and PALB2 (Partner and localizer of BRCA2). BRCA1 and BARD1 cooperate to bind RAD51 through amino acid 33

residues 758-1064 and 123-261 respectively [453, 454], while BRCA1 interacts with the scaffold protein PALB2 through its coiled coil domain which mediates the interaction between BRCA1 and BRCA2 (Figure 1.5) [434, 455, 456].

BRCA2 is another large nuclear protein (384 kDa) with 8 tandem BRC repeats and 3 OB-folds (oligonucleotide/oligosaccharide-binding) with the helical domain making up the DBD (BRCA2 DNA/DSS1-binding domain) (Figure 1.5B) [457-459]. BRCA2 plays a critical role in sequestering RAD51 monomers through the tandem BRC repeats and facilitating the generation of RAD51 nucleoprotein filaments during the repair of a

DSB (Figure 1.5B) [460, 461]. Furthermore, BRCA2 and PALB2 interact to maintain the

G2 checkpoint independently of their role in HR [462]. Therefore, both BRCA1 and

BRCA2 play an important role in repairing DNA DSBs.

1.4.3 BRCA1/2 and the repair of double-strand breaks

Repair of a DSB is mediated by two main pathways: non-homologous end joining

(NHEJ) and homologous recombination (HR). NHEJ is the predominant repair pathway occurring in all cell cycle phases while HR is restricted to the S and G2-phases of the cell cycle [463-465]. NHEJ is potentially an error-prone pathway that involves the direct

-ligation of the damaged ends [466]. In contrast, HR is a relatively error-free pathway that involves the use of a sister chromatid or homologous sequence to repair the DSB

[467, 468]. BRCA1 plays an important role in the choice between NHEJ and HR while both BRCA1 and BRCA2 facilitate HR.

The role of BRCA1 in DNA repair pathway choice and the S and G2/M checkpoint

Once a DSB is formed, the two ends of the break are rapidly bound by the heterodimer Ku70 and Ku80 (Ku70/Ku80) and DNA-PKcs forming DNA-PK complex 34

[469-473]. During S-phase, the initiating step for HR is the binding of the MRN complex to the DNA end and DNA-PK removal [474]. The MRN complex mediates a 1- dimensional search facilitated by RAD50 for free DNA ends while DNA-PK increases the ability of MRN to remove Ku70/Ku80 from the ends to allow MRN occupancy [474,

475]. MRN then activates ATM leading to the phosphorylation of histone H2A.X on serine 139 (γH2AX) (Figure 1.6) [476]. MDC1 (mediator of DNA damage checkpoint protein 1) binds to γH2AX and recruits more MRN and ATM to further amplify the signal

[477-479]. Activated ATM then phosphorylates MDC1, CHK2 (checkpoint kinase 2), and

BRCA1 leading to the S and G2/M checkpoints while phosphorylation of p53 induces the G1 checkpoint [480-482]. In particular, ATM phosphorylation of BRCA1 on S1387 leads to S-phase checkpoint while S1423 leads to G2/M checkpoint activation [483,

484]. Phosphorylated MDC1 then recruits the E3 ligase RNF8 which polyubiquitinates the linker histone H1, primed by the E3 ligase HUWE1, leading to the recruitment of another E3 ligase RNF168 (Figure 1.6) [485-488]. RNF168 cooperates with RNF8 to ubiquitinate histone H2AK13/K15 leading to the recruitment of RAP80 and 53BP1

(Figure 1.6) [489-492]. Binding of RAP80 on K63-linked H2AX recruits the BRCA1-A complex to DSBs to fine tune HR-mediated repair by preventing the over-accumulation of the BRCA1-B and C complexes [493, 494]. 53BP1 is a scaffold protein that binds to monoubiquitinated H2AK15 and di-methylated H4K20 (H4K20me2) and dimerizes on histones [495-497]. H4K20me2 marks pre-replicative DNA, therefore favoring 53BP1 binding at un-replicated regions [498]. ATM phosphorylated 53BP1 interacts with RIF1 and the Shieldin complex (Shld3, Rev7, Shdl1, and Shld2) [499-502] along with PTIP through distinct phosphorylation sites to prevent HR by blocking end resection (Figure 35

1.6) [503-505]. Therefore, BRCA1-BARD1 prevents NHEJ in two ways: by displacing

53BP1 binding and counteracting RIF1 localization. BARD1 binds unmethylated H4K20, a mark of post-replicative DNA, along with directly binding mono-ubiquitinated H2AK15 thereby excluding 53BP1 binding and localizing BRCA1 to DSBs (Figure 1.6) [506,

507]. Notably, TIP60-mediated acetylation of residues neighboring H4K20 disrupts the ability of 53BP1 to bind H4K20me2 [508, 509]. BRCA1, on the other hand, directly counteracts RIF1 localization at DSBs (Figure 1.6) [510-514]. Additionally, SCAI

(suppressor of cancer cell invasion) has been recently implicated to bind to phosphorylated S/T sites adjacent to proline (S/TP) on 53BP1 to prevent RIF1 binding and promote HR [515]. Ultimately the key step in driving HR is the generation of ssDNA at the break site by end resection.

The roles of BRCA1 and BRCA2 during homologous recombination

End resection is a bidirectional process that is initiated by the MRN complex and

CtIP (Figure 1.7). Specifically, the MRN complex binds next to DSB ends and is activated by CDK-phosphorylated CtIP [516-519]. MRE11 nicks the dsDNA and digests

DNA towards the DSB using its 3’ to 5’ exonuclease activity while the 5’ to 3’ exonuclease EXO1 and the endo/exonuclease DNA2 along with the helicases BLM

(bloom syndrome helicase) or WRN (Werner’s syndrome helicase) degrades the DNA away from the DSB to generate long ssDNA overhangs by long-range end resection

(Figure 1.7) [520-524]. Notably, the CtIP endonuclease activity is only required in part for end resection on complex DNA ends generated by topoisomerase poisons or IR

[525-527]. Additionally, end resection of breaks with modified bases or protein complexes is promoted by the MRN interactor EXD2 [528]. 36

Following end resection, RPA rapidly coats the 3’-overhangs preventing the formation of secondary DNA structures and nuclease degradation (Figure 1.7) [529-

531]. Subsequently, BRCA1 through PALB2 recruits BRCA2 to the resected ends and promotes the loading of RAD51 onto ssDNA leading to strand evasion and repair of the

DSB (Figure 1.7) [172, 434, 456, 532-535]. The BRCA1-BARD1 heterodimer then associate with the presynaptic RAD51 filament to promote DNA strand invasion into the homologous sequence and subsequent D-loop formation (Figure 1.7) [536]. Finally,

HELQ, MCM8-9, and MCM8IP (HROB) facilitate D-loop extension and PCNA, DNA polymerase δ, and RFC-1D synthesize the DNA [242, 244, 537].

1.4.4 BRCA1/2 and the protection of stalled forks from nucleolytic degradation

BRCA1 and BRCA2 also play a HR-independent role in protecting replication forks from nucleolytic degradation after fork reversal [154, 214, 215]. Using the DNA fiber technique, the Jasin lab was the first to show that depletion of either BRCA1 or

BRCA2 led to shorter nascent DNA tracts after replication stress [214, 215]. Abrogation of replication fork reversal prevented the degradation seen in BRCA1 or BRCA2- deficient cells [154]. Interestingly, pretreatment of cells with cisplatin activates

PRIMPOL leading to excessive repriming that prevents SMARCAL1-mediated fork reversal, suggesting that PRIMPOL also protects BRCA-deficient replication forks from degradation [538]. Furthermore, while the BRCT domain of BRCA1 plays a critical role in protecting replication forks and HR, the BRCT domain of BARD1 is only required for protecting replication forks by mediating BRCA1/BARD1 recruitment to stalled forks in a poly(ADP-ribose)(PAR)-dependent manner [217]. RPA phosphorylation, on the other hand, facilitates the recruitment of PALB2 and BRCA2 to stalled replication forks to 37

promote fork protection [533, 539]. BRCA2 promotes replication fork stability through a

RAD51 interaction site that, when mutated, does not alter the ability of BRCA2 to perform HR [215, 540]. Notably, it was identified that CDK phosphorylation of BARD1 at

S114 promotes PIN1 isomerization of BRCA1 allowing enhanced binding of RAD51 and promoting fork protection [229]. It remains, however, to be determined how BRCA1 and

BRCA2 act non-redundantly during fork protection.

1.5 The role of stalled fork protection and homologous recombination in cancer predisposition

Failure to properly repair DSBs or protect replication forks from extensive nascent strand degradation are associated with gross chromosomal aberrations, increased genomic instability, and increased mutational burden which are hallmarks of cancer [154, 209, 214-216, 226, 300, 541]. Germline mutations in the tumor suppressors BRCA1 and BRCA2 confer a lifetime risk for breast (72% and 69%, respectively) and ovarian (44% and 17%, respectively) cancer [542], as well as an increased risk for prostate and pancreatic cancers (predominantly by mutations in

BRCA2) [543]. BRCA1/2-deficient cancers are associated with a broad distribution of base substitutions (mutational signature 3) and non-clustered deletions of less than 100 kb (rearrangement signature 5), while BRCA1-loss is additionally associated with small tandem duplications of less than 10 kb (rearrangement signature 3) [544, 545].

Homozygous loss of BRCA1 or BRCA2 is embryonic lethal suggesting that

BRCA1 and BRCA2 are required for normal embryonic development [546-548]. As previously discussed, BRCA1 and contribute to both replication fork protection and the repair of DSBs by HR. Loss of heterozygosity (LOH) of the second allele is frequently 38

observed in BRCA1/2-mutant carriers indicating that haploinsufficiency increases genomic instability, therefore leading to an increased propensity to lose the wild type allele [549, 550]. While heterozygous mutant BRCA1 (BRCA1mut/+) cells are functional for HR they show defects in stalled replication fork protection [551]. Similarly, heterozygous mutant BRCA2 (BRCA2mut/+) cells also displayed fork protection defects

[552]. However, there is still a debate about the relative contributions of HR-mediated repair and fork protection functions of BRCA1/2 to tumor suppression. Feng and colleagues demonstrated that the HR function of BRCA2 prevents under-replication, while fork protection plays a minor role in the suppression of replication stress [540].

Normally, the p53 pathway becomes activated in response to unrepaired DNA damage and replication stress, leading to cell cycle arrest, senescence or apoptosis [553].

Bypass of senescence appears to be a key step in BRCA1/2-deficient breast and ovarian tumorigenesis. In agreement with this premise, BRCA1mut/+ human mammary epithelial cell lines experience increased DNA damage, telomere erosion and premature senescence that can be bypassed by loss of p53 or pRb [554]. Furthermore, p53, pRb, and p16 are frequently mutated or lost in BRCA-deficient breast cancers [555-558].

Loss of the p53 axis enables the loss of second BRCA1/2 allele. Ultimately, loss of both alleles is associated with defects in both stalled fork protection and HR [214, 215, 535,

559, 560]. These observations support a model in which stalled fork protection and HR contribute to tumor suppression in distinct stages [217, 551, 561]. In this case, heterozygous mutations would drive genomic instability by defective stalled fork protection leading to the generation of genetic lesions that can support the viability of

HR-deficient cells which would drive an increase in genomic instability and drive malignancy [217].

Besides BRCA1/2, several other DNA repair factors are mutated in cancer.

Germline mutations in the BRCA1 partner BARD1 have been identified as a pathogenic lesion in hereditary breast and ovarian cancers [562]. Additionally, biallelic mutations in the BRCA1/2-interacting protein PALB2 predispose carriers to breast and ovarian cancer, Wilms’ tumors and medulloblastoma [563, 564]. Monoallelic germline mutations in the RAD51 paralogs RAD51C and RAD51D predispose carriers to breast and ovarian cancers [565-568]. BLM also plays a role in both promoting HR and fork restart by counteracting RAD51 [218, 569]. This function of BLM to counteract RAD51 is distinct from its ability to prevent elevated sister chromatid exchange by dissolving double

Holliday junctions formed during HR [569, 570]. Loss of function mutations in BLM cause Bloom’s syndrome which is characterized by an increased risk of early onset cancer development for several cancer types including leukemias, lymphomas, and colorectal cancers [571]. These studies underscore the importance between maintaining genomic stability through HR and fork protection and preventing tumor development.

1.6 The role of stalled fork protection and homologous recombination in cancer therapeutics

1.6.1 Chemosensitivity

Targeting the vulnerabilities of cells defective for HR or fork protection is a promising therapeutic strategy for treating several cancer types. A study by Heeke et al. characterized the prevalence of pathogenic mutations in 17 different HR/ DNA damage-

genes and determined that these mutations were detected in 17.4% of all solid tumors tested [572]. Therefore, treatments that can cause lesions that will be repaired by HR in normal tissues but will increase genomic instability and cell death in malignant HR mutant cells are currently being used for treating cancer. For instance, since the 1970’s the main line treatments for HR-deficient ovarian cancers have been platinum-based therapies, such as cisplatin and carboplatin [573]. Furthermore, agents that induce the formation of DSBs, which are typically repaired by HR, are also effective against HR- deficient cancers including ionizing radiation, radiomimetics (bleomycin), topoisomerase

I inhibitors (camptothecin, topotecan), and topoisomerase II inhibitors (doxorubicin, etoposide) [573, 574]. Additionally, the fork stalling agent, gemcitabine, is also an approved cancer treatment in combination with IR and platinum-based therapies [575].

Because gemcitabine is a chain-terminating nucleoside analogue and a ribonucleotide reductase inhibitor, it is possible that the increased DNA damage induced by this agent is due to replication fork stalling and excessive degradation, but this agent has not yet been used in fork degradation experiments.

Notably poly(ADP-ribose) polymerase (PARP) inhibition has been demonstrated to sensitize BRCA1/2-deficient tumors [576-578]. Single strand breaks (SSBs), one of the most common DNA lesions, are typically sensed by PARP1 and PARP2 and stimulate SSB repair [579, 580]. PARP inhibition traps the PARP proteins onto the DNA resulting in both unrepaired SSBs that can be converted into DSBs during replication and replication fork collapse due to replication stalling and premature fork restart by

RECQ1 [581]. Therefore, PARP inhibition highlights the contribution of both HR and fork

protection to genomic stability and cellular viability. In fact, PARP inhibition can be synergistic with inhibitors for either HR or cell cycle checkpoint.

PARP trapping increases replication fork stalling [582]. In response to replication stress the ATR-Chk1 pathway promotes fork slowing and prevents origin firing [583,

584]. Therefore, loss of the ATR-Chk1 checkpoint could increase the sensitivity of HR- deficient cells to PARP inhibitors (PARPis) by increasing DSBs by fork collapse.

Consistent with this idea, ATR or CHK1 inhibition increases the sensitivity of cells to

PARP inhibition [585, 586]. Furthermore, the ATM-CHK2 pathway is activated by DSBs.

ATM inhibition and CHK2 inhibitors also sensitize cells to PARP inhibition [587, 588].

Therefore, clinical trials using checkpoint kinase inhibitors in combination with PARPis is warranted. Moreover, CtIP promotes HR and low levels of CtIP expression is associated with lower survival rates in breast cancer [589]. It has been demonstrated that CtIP depletion increases HR-deficiency in the breast cancer cell line MCF7 and sensitizes them to PARPis [589]. Therefore, CtIP inhibitors in combination with PARP inhibition may be a promising therapy.

In addition to PARPis, other synthetic lethal combinations are currently under investigation. Alternative end-joining by Polθ (Theta-mediated end joining) acts predominantly in HR-deficient tumors [104, 590, 591]. Therefore, inhibition of Polθ is a promising anticancer therapy for HR-deficient tumors [592]. Another synthetic lethal combination was identified between BRCA-deficiency and RAD52 [593-595]. As such,

RAD52 inhibition caused additional sensitivity of BRCA1-deficient cells to PARP inhibition [596]. Interestingly, RAD52 inhibition synergizes with Polθ inhibition to increase sensitivity to cisplatin indicating distinct back up pathways to HR [597]. Finally, 42

a synthetic lethal screen revealed that depletion of FEN1 or APEX2 increase lethality in

BRCA1/2-deficient cells suggesting that inhibition of those factors could be a viable avenue for BRCA1/2-deficient tumors [598]. Additionally, the abrogation of the cell-cycle checkpoints alone sensitizes HR-deficient tumors due to an accumulation of DNA damage leading to cell death and therefore these agents are currently in early clinical trial evaluation for BRCA-mutated cancers as a monotreatment [573, 599]. Ultimately, the characterization of factors that increase genomic instability and cell death when inhibited in HR-deficient tumors will be of clinical importance.

1.6.2 Chemoresistance

Chemotherapeutic resistance is the ability of malignant cells to evade or resist the effects of cancer therapies. Through genomic instability driven rapid evolution, cancer cells can develop mechanisms to survive therapeutics leading to cancer relapse after remission, rapid disease progression, and ultimately poorer survival outcomes

[600-602]. As previously described tumors harboring BRCA1/2 mutations are hypersensitive to DNA-damaging treatments, such as platinum-based chemotherapy and PARPi. However a large fraction of BRCA1/2 mutant patients eventually acquire resistance to these treatments [577, 578, 603, 604]. Specifically, the recurrence rates for ovarian cancer is 85% while the recurrence rate for breast cancer is 30% [605-607].

One explanation is that HR-deficient tumors can establish genomic stability and acquire chemoresistance through the restoration of HR. As such, secondary mutations in HR pathway genes such as the RAD51 paralogs RAD51C and RAD51D or reversion mutations in BRCA1 or BRCA2 have been associated with platinum resistance for ovarian cancers and other cancer types [608-610]. Additionally, elevated expression of 43

HR promoting factors can aid in tumor cell survival to high levels of DNA damage.

Therefore, increased RAD51 expression correlates with chemoresistance [611-614].

Additionally, high levels of MRE11, RAD50, or NBS1 reduce DNA damage and are also correlated with chemoresistance and poor prognosis in gastric cancer [615, 616].

Moreover, HELQ, a helicase that promotes the postsynaptic steps of HR, when overexpressed, is associated with resistance to cisplatin [242, 617, 618]. Finally, increased expression of the histone acetyltransferase TIP60 in nasopharyngeal carcinoma cell lines has been demonstrated to promote both HR and HR gene expression leading to cisplatin resistance [619].

On the other hand, re-establishing end resection can also restore HR. In this case, loss of the shieldin complex or 53BP1 increased resistance of BRCA1-deficient tumor cells to PARP inhibition [620, 621]. Alternatively, aberrant NHEJ in HR-deficient cells can drive genomic instability and lethality. Notably, inhibiting NHEJ (by depleting

Ku80, Artemis, or DNA-PKcs) in BRCA-deficient cells exposed to PARPi or FANCD2- deficient cells exposed to platinum treatment prevents lethality and drives resistance

[622, 623]. In agreement with the hypothesis that inhibiting NHEJ in HR-deficient tumor can drive resistance, loss of 53BP1in BRCA1-mutant mammary tumors led to PARPi resistance in mice [624].

Remarkably, about half of all BRCA2-mutant cancers developed chemoresistance in the absence of HR-restoration suggesting the presence of an HR- independent resistance mechanism [625]. In line with this hypothesis, the restoration of fork protection in addition to or independently from HR functionality in BRCA1/2- deficient tumors has been associated with acquired resistance [225]. As such, FANCD2 44

overexpression which suppresses fork instability has been demonstrated to confer

PARPi resistance [626]. Additionally, loss of the SNF2-fork remodeler SMARCAL1 restores fork stability in BRCA1/2-deficient cells and confers PARPi resistance [154].

Similar observations have been reported for loss of PTIP, the SNF-2 family helicase

CHD4, RADX, and EZH2 which are involved in promoting nascent strand degradation at stalled forks [175, 240, 627]. Notably, decreased CHD4 expression correlates with poorer prognosis for patients with BRCA2-mutant cancers [627]. Activation of alternative and compensatory DNA repair pathways in response to replication stress adds a further layer of complexity. It has been recently reported that BRCA1-deficient ovarian cancer cells adapt to recurrent cisplatin treatments by activating PRIMPOL-mediated repriming of stalled forks to protect BRCA1-deficient cells from nucleolytic degradation [538]. In addition, repriming appears as a general alternative mechanism to SMARCAL1- or

HLTF-mediated fork reversal for the restart of DNA synthesis, at the expense of the formation of ssDNA gaps [538, 628]. These small ssDNA gaps may undergo resection and RAD51-mediated HR for proper post-replicative repair [629]. In line with these observations, HR-defective BRCA1-mutant cancer cells adapt to recurrent cisplatin treatment by activating PRIMPOL-mediated repriming to avoid excessive fork remodeling-induced nucleolytic degradation of nascent DNA at the expense of the formation of ssDNA gaps[538]. Additionally, loss of HLTF promotes discontinuous DNA synthesis by PRIMPOL [628]. TLS has been suggested as a third interconnected mechanism, along with fork reversal and repriming, to promote the restart of stalled forks [628]. Aberrant activation of the TLS pathway by FANCJ mutation, reduces fork reversal and promotes synthesis without leaving ssDNA gaps in the presence of HU 45

[630]. Moreover, in HLTF mutant cells that cannot bind the 3’-end of ssDNA, HIRAN mutants, REV1 promotes fork progression by continuous DNA synthesis [628]. TLS also provides an alternative mechanism to HR for the post-replicative repair of ssDNA gaps generated during replication stress [631, 632]. Interestingly, genomic sequencing and analysis of mutational signatures in syngeneic cells treated with genotoxin agents suggest an up-regulation of TLS after BRCA1 depletion [633]. Taken together these recent findings shed new lights on the complex network of factors involved in remodeling, protection, and repair of stalled forks. Understanding the complex interplay between fork remodeling, repriming, TLS and HR in response to replication stress will be of utmost importance for developing novel cancer therapies.

1.6.3 Immunotherapy

Innate immunity is an evolutionary conserved cell-intrinsic reaction to microbial pathogens, which involves multiple mechanisms, and sensor molecules. Cytosolic DNA sensing recognizes and reacts to foreign DNA found in the cytoplasm (e.g., viral DNA), leading to immune responses. The cGAMP synthase (cGAS)–stimulator of interferon genes (STING) pathway has been identified as a critical component of DNA sensing

[634]. Briefly, cytosolic DNA sensed by cGAS activates STING, which triggers the expression of interferon-stimulated genes (ISGs) and induces the inflammatory response (Figure 1.4) [634, 635].

Recent studies have shown that unresolved DNA damage derived from replication fork instability, telomere fragility, common fragile site expression and chromosome mis-segregation often results in the formation of micronuclei upon mitotic progression, which can be detected by cGAS, thus leading to STING activation and the 46

induction of innate immune signaling [636-638]. Accordingly, micronuclei formed as a result of deficiency in BRCA1/2 or RAD51 activate innate immune signaling, and this response is exacerbated by PARP inhibitor treatment [639-643]. Notably, loss of the

SNF2-family members SMARCAL1 and ERCC6L has been associated with an increase of micronuclei formation [185, 302]. However, it remains to be determined whether deficiency of these factors leads to innate immunity induction.

In addition to micronuclei, short species of genomic DNA released into the cytoplasm during DNA replication and repair can also be recognized by cGAS, thereby triggering the activation of innate immune signaling. In particular, it was shown that cytoplasmic ssDNA derived from replication fork intermediates following HU treatment is degraded by TREX1 [356], a nuclease that suppresses innate immune signaling and maintains immune tolerance [644, 645], thus providing direct evidence that processing of stalled forks can generate cytoplasmic DNA and activate innate immunity. Additional studies have shown that the innate immune response is activated also by cytoplasmic ssDNA generated upon treatment with other replication stress-inducing agents, such as mitomycin C and cisplatin, or in response to IR [646]. Of note, depletion of nucleases such as DNA2 and EXO1, but not MRE11 and CtIP, restrains innate immune signaling, suggesting that ssDNA molecules that induce innate immunity require extensive DNA- end resection activities [646]. In addition, loss of SAMHD1, a factor that stimulates the exonuclease activity of MRE11 at stalled forks, was shown to prevent nascent DNA degradation at reversed forks and stimulate the helicase and endonuclease activities of

RECQ1 and MRE11, respectively, thus resulting in the accumulation of cytosolic ssDNA fragments and the subsequent activation of the cGAS-STING pathway [230]. 47

Cytoplasmic ssDNA fragments deriving from stalled replication intermediates and cGAS-STING-dependent innate immune signaling have also been observed in BRCA1- deficient breast cancer cells, although it remains unclear whether this phenotype is dependent on defective fork protection, as in the case of RAD51-depleted cells [641,

647]. Interestingly, it was reported that prostate cancer cells accumulate cytoplasmic dsDNA derived from nascent DNA in a manner dependent from MUS81, suggesting that it may originate from the cleavage of stalled forks [359]. Further studies are required to understand the relationship of fork remodeling with cytosolic DNA sensing and inflammation and evaluate whether inhibition of SMARCAL1, ZRANB3, and HLTF may stimulate the cGAS-STING pathway. Overall, the above findings demonstrate that replication fork instability leads to aberrant inflammatory response in both malignant and non-malignant cells (Figure 1.4).

cGAS-STING-dependent innate immune signaling enhances tumor immunogenicity, and consequently the response of tumors to immunotherapies, such as immune checkpoint blockade (ICB) therapies that use antibodies inhibiting the immune checkpoint factors PD-1, PD-L1 and CTLA-4 [648-671]. High PD-L1 expression on tumor cells restricts T-cell activity by binding to the PD-1 receptor on tumor infiltrating T- cells representing a tumor adaptive immune response [672-674]. Notably, the expression of PD-L1 depends upon STING activation and is induced in response to replication stress [647]. Importantly, in the context of ICB therapy, higher levels of PD-

L1 on tumor cells correlate with better response to PD-1/PD-L1 antibody-mediated immunotherapy [675]. STING activation in response to replication stress can also result in the expression of CXCL10, a chemo-attractive cytokine that correlates with high 48

tumor-infiltrating cytotoxic T cells [639], as well as type-I interferon signaling, which promotes anti-tumor immunity [676-678]. However, recent studies have also shown that activation of the cGAS-STING pathway in chromosomally unstable tumors drives metastasis [679]. Understanding the connection between replication fork instability and the cGAS-STING pathway will therefore be critical for defining underlying mechanisms of anti-tumor immunity and metastasis and predicting the response to ICB therapy in cancer patients (Figure).

1.7 The chromatin remodeler SMARCAD1

SMARCAD1 (SNF2 family ATPase SWI/SNF-related, matrix-associated actin- dependent regulator of chromatin, subfamily A, containing DEAD/H Box-1) is another member of the INO80 sub-family of SNF2-family chromatin remodelers [265, 266]. SMARCAD1 is a ubiquitously expressed protein involved in DNA repair, heterochromatin maintenance, transcription, and pluripotency [680-688]. SMARCAD1 is conserved from yeast and structurally contains N-terminal tandem CUE (coupling ubiquitin to ER degradation) domains and a C-terminal SNF2-like helicase domain (Figure 1.8) [680].

While full-length SMARCAD1 (1026 amino acids) is expressed in all tissues, a truncated isoform of SMARCAD1 (600 amino acids) lacking the CUE domains is expressed in skin fibroblasts and to a lesser extent in keratinocytes and esophageal tissues [689, 690].

Mutations in the skin-specific isoform causes an autosomal dominant form of adermatoglyphia and Basan syndrome due to altered epidermal differentiation- associated gene expression [689-692]. Strikingly, SMARCAD1 has been demonstrated to promote pancreatic and breast cancer cell growth and metastasis in vitro [693-695] and has been predicted to be associated with bladder cancer [695, 696]. Moreover, 49

SMARCAD1 expression is elevated in mammary, embryonal, and lymphoid tumors

[697]. Additionally elevated expression correlates with poor pancreatic cancer prognosis

[695, 697]. However how mechanistically SMARCAD1 may promote tumorigenesis is still poorly understood.

KAP1 (KRAB (Krüppel associated box)-associated protein 1) is the major binding partner of SMARCAD1 which is pulled-down in equivalent stoichiometric amounts [680].

While CUE domains typically act as ubiquitin binding domains, the tandem CUE domains of SMARCAD1 have recently been implicated to mediate interaction with KAP1

(Figure 1.8) [698]. Both KAP1 and SMARCAD1 play an important role in maintaining heterochromatin during replication (Figure 1.9A) [680]. SMARCAD1 directs histone deacetylation by the coordinated actions of its binding partners histone deacetylases

HDAC1 and HDAC2 and promotes histone methylation by its interactor histone methyltransferase G9a/GLP (Figure 1.9A) [680]. KAP1 interacts with Suv39h1 to reinstate the tri-methylation of histone H3K9 after replication (Figure 1.9A) [699]. Di-and tri-methylated histone H3K9 has been implicated in HP1 binding and heterochromatin formation [700, 701]. SMARCAD1 acts at sites of replication through its interaction with

PCNA, although no canonical PCNA interaction motif has been identified so far [702].

Strikingly, heterochromatin maintenance is a conserved function with the fission yeast ortholog Fft3 [703] and the budding yeast ortholog Fun30 [704]. In addition to

SMARCAD1’s role at the replication fork, SMARCAD1 facilitates DNA repair.

SMARCAD1 also contributes to the completion of mismatch repair (MMR) and

HR. The MMR pathway recognizes and repairs mis-paired bases following replication in a process dependent on lesion recognition by MutSα (MSH2 and MSH6) or MutSβ 50

(MSH2 and MSH3) [705-708]. SMARCAD1 facilitates MMR by promoting nucleosome exclusion in a MSH2-dependent manner in Xenopus egg extracts and HeLa cells

(Figure 1.9B) [680, 685, 688]. This is mediated by SMARCAD1’s interaction with MSH2 and MSH6 [680, 685]. How SMARCAD1 excludes nucleosomes during MMR, however, is still unknown. With regards to HR Fun30 has been shown to play a role in promoting

Exo1-mediated long-range end resection, but the mechanism of SMARCAD1-mediated end resection is less understood (Figure 1.10) [681, 709-712]. Fun30 has been extensively demonstrated to facilitate Exo1 long-range end resection by counteracting the 53BP1 homolog, Rad9, and is recruited to sites of DNA damage through CDK- phosphorylation dependent binding to the TOPBP1 homolog, Dpb11 [681, 709, 713,

714]. Fun30 has a higher affinity for and is activated by ssDNA bound nucleosomes providing a potential mechanism for short-range resection as a means of activating

Fun30 mediated remodeling [712]. Furthermore, purified Fun30 can slide nucleosomes in an ATP-dependent manner in vitro [715]. SMARCAD1 and Fft3 have recently been shown to mediate long-range end resection [681, 682, 716]. In particular, SMARCAD1 has been demonstrated to be recruited to sites of laser or Fok1-mediated DSBs in human osteosarcoma cell lines [681]. Additionally, depletion of SMARCAD1 reduced ssDNA formation after IR and SMARCAD1-depleted cells were defective in gene conversion from an I-SceI-induced DSB [681]. In contrast to the tandem CUE domains binding to KAP1, Densham et al. suggested that the CUE domains recruit SMARCAD1 to DSBs by binding to BRCA1/BARD1 mediated H2A ubiquitination leading to displacement of 53BP1 from the break [682]. While this may be an attractive hypothesis there is substantial evidence that refutes this model. First, the SMARCAD1 CUE 51

domains have a higher affinity for KAP1 than mono-ubiquitin [698]. Second, the enzymatic activity of the BRCA1/BARD1 ubiquitin ligase is not essential for HR or tumor suppression [717, 718]. Finally, SMARCAD1 phosphorylation by ATM and stabilization through RING1-mediated ubiquitination are critical for SMARCAD1 recruitment to and function at DSBs (Figure 1.8) [686]. This indicates that SMARCAD1 does not require histone ubiquitin-binding for recruitment. In fact, ATM phosphorylated-KAP1 has also been implicated in decondensing chromatin during HR in order to facilitate end resection [719, 720]. Taken together, these studies may support a mechanism in which

KAP1 and SMARCAD1 act together to facilitate chromatin de-condensation during HR, thus promoting end resection. How SMARCAD1 and KAP1 decondense the chromatin during HR, however, is still not known.

1.8 Objectives

Replication stress poses a risk to the fidelity of DNA replication and eventually contributes to genomic instability and mutagenesis. Replication fork reversal is a frequent protective mechanism during replication stress that allows forks to resume

DNA replication without chromosomal breakage [154]. HR factors, such as BRCA1,

BRCA2, BARD1, RAD51, have been found to play a protective role by preventing fork degradation on these reversed forks [214, 215, 217]. Recently, Lopes and Penengo have characterized nucleosomes on the reversed arm of the forks [318]. These nucleosomes have been suggested to pose a barrier to resection [228, 318]. Therefore, how nucleases may counteract chromatin deposited on the reversed arm of the fork is still not known.

Another unanswered question is about the possible role of SMARCAD1 during replication stress. Unlike SMARCAD1, Fun30, has been demonstrated to play a role in

DNA damage tolerance after replication stress [721, 722]. fun30Δ yeast are sensitive to replication stress agents like MMS and HU [721, 722]. Furthermore, in yeast lacking the

HLTF/SHRPH ortholog, Rad5, Fun30 contributed to replication stress sensitivity suggesting that Fun30 negatively regulates HR-mediated repair at the fork [721].

Additionally, Fft3 has recently been demonstrated to be required for replication fork resection and restart [723]. Moreover, like Fun30, fft3Δ yeast are sensitive to MMS

[723]. While SMARCAD1 promotes HR through long-range end resection and is associated with the replisome, it is not known whether SMARCAD1 plays a role during replication stress.

Therefore, we hypothesize that SMARCAD1 may promote nucleolytic degradation by remodeling the chromatin on the regressed arm of the replication fork.

This is based on these four observations: 1) nucleosomes are present on the regressed arm of the fork, 2) SMARCAD1 promotes end resection during HR, 3) SMARCAD1 stably associates with the replication fork, and 4) the yeast orthologs, Fun30 and Fft3, play a role during replication stress. To test this hypothesis, we sought to use BRCA1/2- deficient mammary epithelial cell lines. Loss of either BRCA1 or BRCA2 in replication stress leads to replication fork degradation [214, 215]. As such, if SMARCAD1 promotes replication fork resection, then loss of SMARCAD1 can prevent nascent DNA degradation in these cells. Consequently, the objective was to elucidate a new function for SMARCAD1 during replication stress and to determine its mechanism of action.

1.9 Figures

Figure 1. 1 Mechanisms of fork restart upon replication stress. When the replication machinery encounters a replication block (stop sign), replication fork progression stalls, leading to the formation of single- stranded DNA (ssDNA) that is rapidly coated by the RPA trimer (I). Restart of stalled forks can occur through lesion bypass by translesion synthesis or fork repriming (II-III). Translesion synthesis allows the direct bypass of the DNA lesion (II), while fork repriming promotes the restart of DNA synthesis downstream of the lesion (III). Alternative to lesion bypass, stalled forks can undergo reversal mediated by fork remodelers, including the SNF2-family members SMARCAL1, ZRANB3 and HLTF (IV). Fork reversal slows down replication fork progression, thus allowing sufficient time for repair of the lesion. Reset of reversed forks can then promote the restart of DNA synthesis once the lesion has been removed (V). Fork reversal could also directly promote the bypass of the DNA lesion by enabling DNA synthesis on the nascent DNA strand of the sister chromatid (green) through template switching. Following fork reversal, the exposed ends of the regressed fork are stabilized by RAD51 to prevent extensive degradation by the nucleases MRE11, DNA2 and EXO1 and protect the integrity of the fork (IV). BRCA1/2, RAD51 paralogs and Fanconi anemia proteins cooperate with RAD51 to maintain fork protection. Defective fork protection causes extensive fork resection and processing of the regressed fork by the endonucleases SLX4 and MUS81, leading to fork collapse (VI). Fork collapse can also result from MUS81-mediated cleavage of persistently arrested stalled forks or from replication forks encountering single-strand DNA breaks. Collapsed forks are repaired by HR-mediated fork restart pathways dependent on RAD51 and/or RAD52 (VI-VIII). Key factors involved in each pathway and described in the main text are indicated.

Figure 1. 2 The SNF2-family of proteins. SNF2-family sub-family relationships based on the domain alignments of the helicase region adapted from Flaus et al. [265] (not to scale). Alongside the alignments are the schematic representations of the protein domains for the family members (not to scale). All SNF2-family members contain a helicase domain consisting of DEXDc and HELICc motifs. Unlike other SNF2-family members, INO80-like proteins contain a large insertion within the helicase domain. The SMARCA-like group consist of SMARCA2 (1590 aa) and SMARCA4 (1647 aa). The ISWI-like group consists of SMARCA1 (1054 aa) and SMARCA5 (1052 aa). The CHD-like group consists of CHD1 (1710 aa), CHD2 (1828 aa), CHD6 (2715 aa), CHD7 (2997 aa), CHD8 (2581 aa), CHD9 (2897 aa), CHD3 (2000 aa), CHD4 (1912 aa), and CHD5 (1954 aa). CHD3, CHD4, and CHD5 have N-terminal plant homeodomain (PHD) fingers not represented in the schematic. The INO80-like group consists of INO80 (1556 aa) and SMARCAD1 (1026 aa). INO80 has an N-terminal PHD domain, while SMARCAD1 has two coupling of ubiquitin to ER degradation (CUE) motifs not represented in the schematic. The RAD54-like group consists of RAD54L (747 aa), HELLS (838 aa), and ATRX (2492 aa). SMARCAL1 (954 aa), ZRANB3 (1079 aa) and HLTF (1009 aa) (in red) are the only SNF2-family members that have been currently shown to regress stalled replication forks in vivo. HLTF contains a RING domain that mediates the interaction between HLTF and the substrates to ubiquitinate. SMARCAL1 harbors an RPA2-binding motif to directly interact with RPA. ZRANB3 contains a PCNA-interacting protein (PIP) motif, AlkB homolog 2 PCNA interacting motif (APIM) and an NZF domain that cooperate to bind polyubiquitinated PCNA. The HARP, HIRAN, and SRD (substrate recognition) domains of HLTF, SMARCAL1, and ZRANB3, respectively, enable structure specific DNA binding. Finally, the HNH motif of ZRANB3 is a catalytic nuclease domain.

Figure 1. 3 Methods for studying replication fork transactions. Overview of the current techniques used to (1) monitor replication fork dynamics and visualize replication fork structures, (2) identify protein associated with and localized on nascent DNA, (3) monitor locus-specific replication fork progression, (4) study DNA lesion bypass and repair events at replication fork barriers, and (5) reconstitute replication fork transactions in vitro. DNA fibers and DNA combing techniques enable the monitoring of individual replication fork progression, while electron microscopy allows high-resolution detection of DNA intermediates and replication fork structures, such as reversed forks and ssDNA gaps. Protein identification and localization on newly synthesized DNA can be monitored by iPOND (isolation of protein on nascent DNA), PLA-based approaches, and by super-resolution microscopy methods, such as STORM (stochastic optical reconstruction microscopy). Techniques, such as SMARD (single molecule analysis of the replicated DNA), nanopore-based sequencing methods (D-NAscent, Fork-seq), and optical mapping enable the study of replication fork dynamics at genomic loci of interest. Antibody-labeled DNA lesions or programmable barriers, such as the E. coli Tus-Ter system, enable the analysis of events occurring when replication forks encounter obstacles. Xenopus egg extracts can be utilized to study replication fork progression, and bypass and repair of DNA lesions. Similar studies can be conducted using purified replication factors that enable DNA synthesis in vitro. Purified proteins can also be utilized to detect enzymatic activities on fork-like substrates or define DNA transactions at single-molecule level using DNA curtains or optical tweezers technologies combined with fluorescence microscopy.

Figure 1. 4 Cellular effects induced by replication stress. Failure to complete DNA replication due to replication stress results in under-replication and sister-chromatid bridge formation. If the bridges are not resolved during anaphase, the sister chromatids remain attached to each other, resulting in chromosome mis-segregation, lagging chromosome formation, or chromosome breakage. Chromosomal breakage and mis-segregation results in the formation of chromosomal aberrations. Chromosomal fragments or lagging chromosomes that fail to be incorporated into the daughter cell nucleus form micronuclei. Errors during DNA synthesis and DNA repair processes can result in the inaccurate duplication of the genome and mutagenesis. Cytoplasmic DNA fragments and micronuclei originating upon replication stress may be recognized by the cGAS-STING pathway, leading to the induction of interferons, immune-stimulated genes (ISGs), and immune checkpoint factors (e.g., PD-L1). PD-L1 expression inhibits immune cell surveillance, leading to immune evasion and immunosuppression.

Figure 1. 5 Domain organization and binding partners of BRCA1 and BRCA2. Schematic overview of the functional domain organization for BRCA1 and BRCA2 (not to scale). Both BRCA1 and BRCA2 contain nuclear localization signal (NLS) residues to promote their import into the nucleus. (A) BRCA1 interacts with its obligate heterodimer BARD1 through its N-terminal RING (Really Interesting New Gene) finger domain conferring E3 ubiquitin ligase activity. CHK2 phosphorylates BRCA1 on serine 988 (S988) and ATM phosphorylates S1387, S1423, and S1524 to promote BRCA1 function during homologous recombination (HR). A region spanning between amino acids 758-1064 facilitate BRCA1 interaction with RAD51 to promote strand invasion during HR. The coiled-coil domain enables the interaction between PALB2 an associate of BRCA2 (as indicated by the light green box). The phosphor-binding BRCT domain enables BRCA1 association with phosphorylated Abraxas, BACH1, and CtIP to form the BRCA1-A, BRCA1-B, and BRCA1-C complexes, respectively. (B) BRCA2 associates with PALB2 at the N-terminus at amino acid residues 21-39. BRCA2 contains eight BRC repeats that bind RAD51 monomers spanning amino acid residues 1009-2083. The DNA binding domain (DBD) consists of a helical domain (dark orange) and three OB-folds along with a tower domain (light orange) facilitating BRCA2 binding to DNA. The C-terminus of BRCA2 contains a CDK2 phosphorylation site (S3291) and binds RAD51 nucleofilaments.

Figure 1. 6 DNA double strand break repair pathway choice. DNA double strand break (DSB) ends are rapidly coated by the Ku70/Ku80 heterodimer and the MRE11 (M), RAD50 (R), NBS1 (N) complex. Binding of the MRN complex recruits ATM which phosphorylates several targets including itself and H2AX to form γH2AX. γH2AX is bound by MDC1 that bind additional MRN and ATM providing a feedback loop and amplification of the γH2AX signal. RNF8 binds MDC1 and cooperates with HUWE1 to poly- ubiquitinate the linker histone H1 leading to the recruitment of RNF168 and ubiquitination of H2AK15. In G1 and early S-phase there is low CDK-activity and a high amount of histone H4K20me2 marking pre- replicative DNA. While in late S-phase through G2 there is high CDK activity and low H4K20me2. 53BP1 binds both H4K20me2 and H2AK15ub enabling the recruitment of PTIP, RIF1, and the Shieldin complex which restricts resection and promotes non-homologous end joining (NHEJ) mediated by DNA-PKcs and Ku70/Ku80 (DNA-PK), DNA ligase IV and the XRCC4 complex (XRCC4 and XLF). High CDK activity enables the phosphorylation of CtIP. Binding of BARD1 on unmethylated H4K20 and H2AK15ub recruits 59

BRCA1 to DSBs preventing 53BP1 and RIF1 accumulation, thereby promoting end resection. Extensive end resection promotes DSB repair by homologous recombination (HR).

Figure 1. 7 The roles of BRCA1 and BRCA2 during homologous recombination. Double strand breaks (DSBs) are rapidly sensed and bound by the MRN complex (MRE11, RAD50, and NBS1). The MRN complex recruits CtIP which is then phosphorylated by CDK leading to initial end resection by the MRN complex and CtIP. Resection by the nucleases EXO1 and DNA2 in coordination with WRN and BLM 61

generates long 3’-single stranded (ssDNA) overhangs. The ssDNA is then coated by the RPA trimer (RPA1, RPA2, and RPA3) to prevent DNA reannealing. BRCA1-BARD1-PALB2-BRCA2 facilitate the exchange of RPA with the recombinase RAD51. BRCA1/BARD1 binds the RAD51 nucleofilaments to facilitate strand-invasion to enable the completion of homologous recombination by DNA synthesis, Holliday Junction formation and ultimately resolution by structure-specific endonucleases.

Figure 1. 8 Domain structure and binding partners of SMARCAD1. SMARCAD1 is a SWI/SNF remodeler in the INO80-subfamily as characterized by the SNF2 helicase domain that is interrupted by a long insertion (helicase ATP-binding domain and helicase C-terminal domain). The nuclear localization signal (NLS) permits SMARCAD1 import into the nucleus. The region spanning from amino acids 157-294 contains two tandem CUE (coupling of ubiquitin to ER degradation) domains, typically a helix that binds ubiquitin. The CUE domains have been suggested to bind ubiquitinated histone H2A mediated by the BRCA1/BARD1 heterodimer. However, the ubiquitin binding affinity of the CUE domains are weak. Independently of ubiquitination, the first CUE domain mediates SMARCAD1 binding to its constitutive and stoichiometric partner KAP1. SMARCAD1 has been demonstrated to contain a conserved CDK phosphorylation site a threonine 71 (T71) that permits association with TOPBP1 and recruitment to double strand breaks (DSBs) and promote homologous recombination (HR). Phosphorylation of T906 by ATM and ubiquitination by RING-1 on lysine 905 (K905) have also been required for SMARCAD1 localization to DSBs and stabilization to promote HR.

Figure 1. 9 SMARCAD1 maintains heterochromatin through replication and promotes mismatch repair. Along with SMARCAD1’s role in promoting homologous recombination, SMARCAD1 also maintains heterochromatin formation through replication and facilitates mismatch repair. (A) Heterochromatin (silent chromatin) regions are marked by histone H3 lysine 4 tri-methylation (H3K4me3) which is bound by the transcriptional repressor heterochromatin protein 1 (HP1). During replication fork progression, newly deposited acetylated histones are introduced. SMARCAD1 in association with PCNA (not depicted), KAP1, the histone deacetylases HDAC1 and 2 (HDAC1/2) along with the lysine methyltransferases GLP and G9a (GLP/G9a) remove the acetylation and promote the mono- and di- methylation of H3K4. KAP1 in association with the methyltransferase Suv39h1 and PCNA (not depicted) enables the re-establishment of H3K9me3 and HP1 binding. (B) DNA mismatches, like the one depicted of a guanine (G)/adenine (A) mismatch, are recognized by the sliding clamps MutSα (MSH2-MSH6) and to a lesser extent by MutSβ (MSH2-MSH3) (not depicted). SMARCAD1 interacts with MSH2MSH6 and promotes the remodeling of the local nucleosomes to enable the completion of mismatch repair through EXO1 degradation of the mismatch containing area and re-synthesis of the degraded segment leading to the removal of the replication error.

Figure 1. 10 SMARCAD1 promotes long-range end resection during homologous recombination. After the formation of a double strand break (DSB), 53BP1 binds to H4K20me2 and H2AK15ub as indicated in Figure 1.4. DSBs are rapidly sensed and bound by the MRN complex (MRE11, RAD50, and NBS1). The MRN complex then recruits CtIP which is then CDK-phosphorylated leading to the initial end resection of DNA by the MRN complex and CtIP. 53BP1 blocks the progression of the nucleases EXO1 and DNA2 preventing long-range end resection and repair by homologous recombination (HR). Recruitment of SMARCAD1 leads to the remodeling of the nucleosomes in the vicinity of the DSB permitting long-range end resection and HR completion.

Chapter 2: The role of SMARCAD1 in promoting fork

degradation in BRCA1 and BRCA2-deficient cells

This chapter was written with help from Alberto Ciccia.

2.1 Introduction

DNA replication is the process of genome duplication during cellular proliferation.

Failure to accurately replicate the genome causes genomic instability, an enabling characteristic for oncogenesis [1-3]. DNA replication fidelity is often challenged by obstacles, such as DNA lesions, nucleotide pool depletion, and collisions with the transcriptional machinery, that can cause replication fork slowing or stalling, leading to replication stress and eventually contributing to genomic instability and mutagenesis

[724]. Cells have evolved various replication maintenance mechanisms to protect replication fork integrity [51, 725]. One level of replication maintenance that has emerged recently is fork reversal, which stabilizes the replication fork and facilitates the restart of stalled forks.

Replication fork reversal is a protective mechanism commonly occurring during replication stress that remodels the replication fork into a four-way, “chicken foot”-like, structure, thus allowing the resumption of DNA synthesis without chromosomal breakage [159, 726]. Several enzymes have been demonstrated to remodel stalled forks in vivo in response to replication stress, including the DNA helicase FBH1, the recombinase RAD51, and the SNF2-family remodelers SMARCAL1, ZRANB3, and

HLTF [37, 154, 155, 163, 187]. While fork regression is a protective mechanism, the

extruded nascent DNA on the regressed arm is a prime target for nuclease-dependent end resection by DNA2, MRE11, and EXO1 [36, 154, 156, 159, 727]. Limited fork resection by the above nucleases is critical for controlled processing of the regressed arm. Indeed controlled resection may assist in the removal of the chicken-foot structure leading to the generation of a fork structure by which replication can restart, in a similar manner to the E.coli RecBCD heterotrimer, or facilitate HR-mediated restart of the regressed forks [37, 153, 159]. However, uncontrolled fork degradation can lead to chromosomal instability [153, 159]. In order to protect the regressed arm from extensive nucleolytic degradation, RAD51 nucleoprotein filaments are assembled on the regressed arm and stabilized by several DNA repair factors such as: BRCA1, BRCA2,

BARD1, PALB2, ABRO1, WRNIP1, FANCA, FANCD2, BOD1L, SETD1A, and the

RAD51 paralogs (XRCC2 and RAD51C) [155, 214-216, 218, 226-228]. Nucleosomes have been characterized on the reversed arm of the forks and have been suggested to pose a barrier to reversed fork resection [196, 228, 318]. How nucleases may counteract the chromatin deposited on the reversed arm of the fork to mediate resection is still not known. Without fork protection, reversed forks undergo uncontrolled degradation, leading to genomic instability and sensitivity to chemotherapeutic agents

[154, 215, 225, 728]. Notably, re-establishment of fork protection has been implicated in promoting tumor resistance to chemotherapies [154, 175, 240, 626, 627]. Therefore, identifying proteins associated with replication fork protection is crucial for cancer therapy.

SMARCAD1 is a SNF2-family chromatin remodeler that promotes heterochromatin maintenance at the replication fork. Furthermore, SMARCAD1 has 67

been shown to stimulate end-resection at DSBs to promote homologous recombination

(HR) [680-682, 686, 709, 729]. Both DNA end-resection and heterochromatin maintenance functions of SMARCAD1 have been conserved from the yeast orthologs

Fun30 (Saccharomyces cerevisiae) and Fft3 (Saccharomyces pombe) to the human protein [681, 709, 714, 723, 730, 731]. In particular it has been proposed that

SMARCAD1 and its homologs enable end-resection through the mobilization or eviction of nucleosomes [681, 709-714, 716, 730]. Recently, both Fun30 and Fft3 have been characterized to contribute to cellular tolerance in response to the replication stress- inducing agents hydroxyurea (HU) and methyl methane sulfonate (MMS). Specifically,

Fun30 has been characterized to inhibit HR-mediated DNA damage tolerance promoted by the HLTF/SHPRH ortholog Rad5 [721, 722]. Strikingly, Fft3 promotes fork resection and HR-mediated fork restart in response to replication fork stalling [732]. However, while SMARCAD1 promotes HR through long-range end resection and is associated with the replisome, it is not known whether SMARCAD1 plays a role during replication stress.

Here we report that SMARCAD1 promotes resection of nascent DNA in

BRCA1/2-deficient mammary epithelial cells. We demonstrate that its chromatin remodeling domain is critical for promoting resection. Furthermore, we show that

SMARCAD1 promotes fork restart and that loss of SMARCAD1 leads to an increase in

DNA damage induced by replication stress agents. Finally, we demonstrate that the replication-induced DNA damage in SMARCAD1- and BRCA1-deficient cells is dependent on SMARCAL1-mediated fork reversal. Taken together, these findings provide insight into the role of SMARCAD1 at stalled replication forks. 68

2.2 Results

SMARCAD1 depletion prevents nascent DNA from degradation upon fork stalling in

BRCA1/2-deficient cells

A recent study in the fission yeast Saccharomyces pombe has demonstrated that

Fft3 is required for end-resection at stalled replication forks and promotes HR-mediated fork restart [732]. In mammalian cells, replication forks are protected from extensive

DNA degradation by BRCA1 and BRCA2 (BRCA1/2) [214, 215]. Given that SMARCAD1 plays a key role in promoting long range end-resection at DSBs to promote HR [681,

682, 686, 710], we reasoned that SMARCAD1 may resect the nascent DNA of a stalled replication fork unprotected by BRCA1/2. To study the role of SMARCAD1 at the replication fork, we monitored nascent DNA degradation using the DNA fiber technique

[306] in the non-tumorigenic human breast epithelial cell line MCF10A, subjected to shRNA-mediated depletion of SMARCAD1, BRCA1, and/or BRCA2. The MCF10A cells were pulsed sequentially with the thymidine analogs 5-chloro-2’-deoxyuridine (CldU, red) and iodo-2’-deoxyuridine (IdU, green), followed by treatment with 2 mM hydroxyurea (HU) for 5 hours to arrest DNA replication (Figures 2.1A,B). The ratio of

IdU to CldU tract length was assessed to determine fork degradation in response to HU.

As previously demonstrated, depletion of either BRCA1 or BRCA2 significantly reduced the median IdU/CldU length ratio compared to control cells in HU treatment (Figure

2.1C). Upon co-depletion of SMARCAD1 with two independent shRNAs, this reduction in the IdU/CldU length ratio was eliminated (Figure 2.1C). Similarly, SMARCAD1 co- depletion in the human osteosarcoma cell line, U2OS, also prevented the reduction in

IdU/CldU ratio by BRCA1-depletion (Figures 2.2A,B). Taken together this suggests that

SMARCAD1 promotes the resection of the nascent DNA in BRCA1/2-deficient cells.

The ATPase activity of SMARCAD1 is required to promote nascent DNA degradation in

BRCA1-deficient cells

SMARCAD1 depends on its ATPase activity to promote long-range end resection and drive heterochromatin maintenance during replication [681, 686, 710, 729].

Additionally, it has been demonstrated that SMARCAD1 is recruited to DSBs by

TOPBP1 through a conserved CDK phosphorylation site, threonine 71 (T71) [709].

Furthermore, it was shown that SMARCAD1 requires its tandem Cue domains for localization and function [682], and ubiquitination of lysine 905 (K905) by RING-1 [686] to promote end resection. To test whether SMARCAD1 requires its ATPase activity,

TOPBP1 binding, Cue domains, or ubiquitination by RING-1 to promote resection at the replication fork in BRCA1-deficient cells, SMARCAD1 shRNA-depleted cells were complemented with shRNA resistant cDNAs encoding either the wild type (WT),

ATPase-defective (K528R), phosphorylation-defective (T71A), Cue domain-mutant

(CueM), or ubiquitin-defective (K905R) forms of SMARCAD1. Detection of the levels of the complemented SMARCAD1 proteins by western blotting revealed that only the

T71A and K528R mutants were expressed at levels comparable to WT SMARCAD1

(Figure 2.2C). Expression of the WT and T71A forms of SMARCAD1, but not of its

ATPase-defective form, promoted nascent DNA degradation in BRCA1-depleted cells upon HU treatment (Figure 2.1D). This finding suggests that SMARCAD1 promotes nascent DNA degradation in BRCA1-deficient cells in a manner dependent on its

ATPase activity. Future studies are required to determine whether SMARCAD1 promotes MRE11-, DNA2-, or EXO1-mediated stalled fork resection.

SMARCAD1 promotes fork restart

To determine whether SMARCAD1 is required for fork restart similarly to its yeast ortholog Fft3, we monitored fork restart using the DNA fiber technique. To this end, cells were pulsed with CldU and subsequently treated with 2 mM HU for 2 hours followed by an IdU pulse (Figure 2.3A). Individual fibers were assessed for the incorporation of IdU after removal of HU, an indication of successful fork restart. After treatment with HU, depletion of SMARCAD1 alone significantly increased the percentage of stalled forks compared to control cells (Figure 2.3A). However, depletion of BRCA1 did not affect fork restart compared to control cells (Figure 2.3A), suggesting that extensive fork degradation does not prevent fork restart. Interestingly, co-depletion of BRCA1 and

SMARCAD1 led to an increase in the percentage of stalled forks (Figure 2.3A), suggesting SMARCAD1 plays an important role in fork restart regardless of BRCA1/2 status. Future studies are required to test whether SMARCAD1 relies upon its ATPase or other domains for proper fork restart.

SMARCAD1 prevents replication stress induced DNA damage

Blocked reversed forks are cleaved by the endonucleases MUS81 and SLX4, thus generating a DSB, which is repaired by HR [165, 237, 238]. To determine whether loss of SMARCAD1 promotes DSB formation due to failed fork restart, MCF10A cells expressing BRCA1 or SMARCAD1 shRNAs were treated with 100 nM of the topoisomerase inhibitor camptothecin (CPT) and subsequently subjected to an alkaline comet assay for the detection of single-strand DNA breaks (SSBs) and DSBs. As 71

expected, BRCA1-depletion increased the mean comet tail moment after CPT treatment, as compared to control cells (Figures 2.3B and 2.4A). Interestingly, depletion of SMARCAD1 alone also increased the mean tail moment (Figure 2.3B), suggesting that SMARCAD1 protects cells from replication stress-induced DNA damage.

Furthermore, depletion of either BRCA1 or SMARCAD1 led to similar mean tail moment to that observed upon co-depletion of both proteins (Figure 2.3B), suggesting that

SMARCAD1 works in the same pathway as BRCA1 to prevent DSB generation.

Taglialatela et al. previously demonstrated that SMARCAL1-mediated fork reversal is crucial for the DNA damage generated in BRCA1-deficient cells [154].

Therefore, we hypothesized that SMARCAD1-mediated DNA damage in response to replication stress may also be due to fork reversal. To this end, we depleted

SMARCAL1 by siRNA and monitored DSB formation by comet assay (Figure 2.4B). As expected, depletion of SMARCAL1 reduced the mean tail moment in BRCA1-deficient cells (Figure 2.3C) [154]. Interestingly, SMARCAL1 depletion also reduced the mean tail moment for SMARCAD1-depleted cells and SMARCAD1/BRCA1 co-depleted cells

(Figure 2.3C). Taken together, these data suggest that the replication stress induced in

SMARCAD1-depleted cells is mediated by SMARCAL1.

SMARCAD1 promotes nascent DNA degradation in BRCA1/2-deficient cells and fork restart upon replication stress

In this study we characterize a novel role for SMARCAD1 in promoting the resection of the nascent DNA at stalled fork and facilitating efficient fork restart. In particular, we determine that the ATPase activity of SMARCAD1 is essential for promoting resection at the fork (Figure 2.1D ). Additionally, we find that SMARCAD1 72

promotes fork restart and prevents replication stress induced DNA damage (Figures 2.3

A,C).

2.4 Methods and Materials

Cell culture

MCF10A, U2OS and HEK293T cells were obtained from American Type Culture

Collection (ATCC). HEK293T and U2OS cells were cultured in DMEM supplemented with 10% Fetalgro bovine growth serum (RMBIO). MCF10A cells were maintained in a

1:1 mixture of DMEM and Ham’s F12 medium (Thermo Fischer Scientific), supplemented with 5% horse serum (Thermo Fischer Scientific), 20ng/ml human epidermal growth factor (Peprotech), 100ng/ml cholera toxin, 10μg/ml insulin and

0.5μg/ml hydrocortisone (Sigma-Aldrich). Cells were grown in humidified incubators at

o 37 C and 5% CO2.

Plasmids

pDONR223-SMARCAD1 was obtained from the orfeome (ID: 10760). pDONR223-SMARCAD1-T71A, pDONR223-SMARCAD1-K528R, and pDONR223-

SMARCAD1-K905R were generated using inverse PCR with the following DNA oligos respectively: 5’- TGTTCCAGAAgctCCAGATAATG-3’ and 5’-

CTAGAATCTTCTGTTTTTTCAG-3’, 5’- GGGCCTAGGAagaACTATTCAAG-3’ and 5’-

ATTTCATCTGCCAAAATGC-3’, 5’-

ATTAGATGGAcggACTCAGATTTCTGAAAGGATTC-3’ and 5’-

CTGAGGTACCTATGCTGATG-3’. pDONR223-SMARCAD1-CueM was generated using a custom synthesized oligo containing the amino acid changes L168E, F169E,

L195E, L196E, F263E, A285E, and L286E (as described in Densham et al., 2016) 73

cloned into pDONR223-SMARCAD1 plasmid blunted by inverse PCR with the following

DNA oligos: 5’- GAATCTCTAAAAGTGTTTGCAGAAG-3’ and 5’-

TTCCTTCAAAGTCTGAAGTTTAGCA-3’. shRNA resistant pDONR223-SMARCAD1 and mutants were generated using inverse PCR using the oligos 5’- ccacacatgtttagtagtagcaccagtgaaataAGGCGTatgttCAGTAGCaagacaaaatcagcagatgagca

-3’ and 5’- tgctcatctgctgattttgtcttgctactgaacatACGCCTtatttcactggtgctactactaaacatgtgtgg-

3’ prior to Gateway LR recombination using LR Clonase II (Thermo Fisher) into the lentiviral expression vector pHAGE-N-FLAG-HA-DEST-CMV. The following retroviral vectors have been used to express shRNAs in this study: pMSCV-PM vectors containing control firefly luciferase shRNA (CCCGCCTGAAGTCTCTGATTAA) [193],

BRCA1 shRNA (GGCAGGTATTAGAAATGAA), BRCA2 shRNA

(CTCTTAGCTGTCTTAAAGA, SMARCAD1 shRNA #10 (ACGAAGAATGTTTTCCTCT), and SMARCAD1 shRNA#11 (CATAGAGTAGGCCAGACTA). Combinatorial stable shRNA-mediated knockdown of control, BRCA1/2, and/or SMARCAD1 in MCF10A and

U2OS cells was achieved by two sequential rounds of infection and selection using recombinant viral particles derived from the above retroviral vectors carrying resistance to puromycin, blasticidin, or hygromycin. Control, BRCA1-deficient, SMARCAD1- deficient, or BRCA1-SMARCAD1-deficient MCF10A cells were transfected with an siRNA targeting firefly luciferase (control siRNA, GE Dhaemacon) or SMARCAL1 siRNA#1 (GE Dharmacon) using lipofectamine HiPerFect (Qiagen) following the manufacture’s instructions and subjected to comet assays 2 days after transfection. The expression of the lentiviral vectors pHAGE-GFP-FLAG-HA, pHAGE-SMARCAD1, pHAGE-SMARCAD1-T71A, pHAGE-SMARCAD1-K528R, pHAGE-SMARCAD1-CueM, 74

and pHAGE-SMARCAD1-K905R were generated by a sequential round of infection and selection on top of the combinatorial stable shRNA-mediate knockdowns using recombinant viral particles derived from the above lentiviral vectors carrying resistance to puromycin.

DNA fiber analysis

Exponentially asynchronously growing MCF10A and U2OS cells were pulse labelled with two thymidine analogs 25μM 5-cholor-2’-deoxyuridine (CldU, Sigma

Aldrich) and 125μM 5-iodo-2’-deoxyuridine (IdU, Sigma Aldrich) for the indicated times.

To measure DNA degradation, after pulse labeling with the second nucleotide analog, the cells were washed three times with 1X PBS and treated or not with hydroxyurea

(HU, 2mM, Sigma-Aldrich), HU+ mirin (50μM, Sigma-Aldrich) for 5 hours. To measure fork stalling, after the first pulse, the cells were washed 3 times with 1X PBS and treated or not with 2mM HU for 2 hours, washed with 1X PBS three times, and subsequently labeled with the second analog. Labeled cells were trypsinized and resuspended in cold

1X PBS at 4 x 105 cells/mL. Then 2μL of the suspension were spotted onto a pre- cleaned glass slide and lysed with 10μL of spreading buffer (0.5% SDS in 200mM Tris-

HCl pH 7, and 50mM EDTA). After 6 minutes of incubating in a humidity chamber, the chamber was tilted 15-20° relative to the horizontal allowing the DNA to spread. Slides were air dried, fixed with an ice-cold mixture of methanol and acetic acid (3:1) for 2 minutes, rehydrated in 1X PBS, and denatured with 2.5M HCl for 50 minutes at room temperature. Slides were then rinsed in 1X PBS and blocked for 1 hour in blocking buffer (4% BSA and 0.1% Triton X-100 in 1X PBS) for 1 hour at room temperature. Rat anti-Bromodeoxyuridine (BrdU, AbD Serotec, MCA2060T) and mouse anti-BrdU 75

(Becton Dickinson, 347580) were diluted 1:100 each in the blocking buffer and applied to detect CldU and IdU respectively. After 1 hour of incubation in a humidity chamber, slides were washed with 1XPBS and stained with Alexa Fluor 488-labelled goat anti- mouse IgG1 antibody (Thermo Fisher Scientific) and Alexa Fluor 594-labeled goat anti- rat antibody (Thermo Fisher Scientific) diluted 1:300 each in blocking buffer for 30 minutes. Slides were washed with 1X PBS, air dried, and mounted with Prolong Gold

Antifade (Thermo Fisher Scientific) and held at 4°C. Replication tracks were imaged on a Nikon Eclipse 50i microscope fitted with a PL Apo 40X/0.95 numerical aperture (NA) objective and measured using ImageJ software.

Comet Assay

Double and single-stranded DNA breaks were evaluated by alkaline comet assay

(single cell gel electrophoresis). MCF10A cells were plated 1 x105 cells/well in a 12- multiwell plate 2 days after transfection and the following day treated or not with camptothecin (CPT, 100nM, Sigma Aldrich) for 5 hours before collection. Cells were trypsinized, re-suspended in 1X PBS, and mixed with molten LMA agarose and pipetted onto a CometSlide (slide coated with 1% NMA agarose in 1X PBS). The slides were subsequently incubated with a lysis solution at pH 10 (25mM NaCl, 100mM EDTA,

10mM Tris Base) for 16 hours at 4°C, equilibrated in running buffer pH 13 (300mM

NaOH, 1mM EDTA) for 20 minutes. The slides were then placed in a horizontal chamber (FisherBiotech) and alkaline electrophoresis was performed at 19V for 20 minutes. Slides were then washed with a neutralization buffer (0.4M Tris-HCl, pH 7.5) for 15 minutes, fixed with ice-cold ethanol for 5 minutes, and stained with a fluorescent dye (GelRed, Biotium, 1:1,000 in water). At least 75 cells were analyzed for each 76

experimental point using a Nikon Eclipse 50i microscope. The comet tail moment values were determined using the CometScore Software. Apoptotic cells (small comet head with very large comet tails) were excluded from the analysis.

Western Blotting

Cells were collected by trypsinization and lysed in 2X SDS-Page buffer (62.5nM

Tris-HCl pH6.8, 2% SDS, 25% glycerol, 0.01% bromophenol blue, 5% β- mercaptoethanol). Following gel electrophoresis and transfer of cell extracts onto a nitrocellulose membrane, membranes were incubated for 1 hour with blocking buffer

(5% milk in TBS+ 0.1% tween). Membranes were subsequently incubated for 2 hours at room temperature or overnight at 4°C with primary antibodies diluted in antibody buffer

(5% BSA in TBS+ 0.1% tween) and detected using the appropriate horseradish peroxidase (HRP)-conjugated secondary antibody diluted 1:1000 in blocking buffer. The

HRP signal was detected using SuperSignal Western Pico Chemiluminescent Substrate

(Thermo Scientific) and autoradiography films (Research Products International). Anti-

SMARCAD1 (1:20,000, ThermoFisher Scientific, PA5-53482), anti-SMARCAL1

(1:1,000, SantaCruz, sc-376377), anti-BRCA1 (1:75, SantaCruz, sc-6954), anti-BRCA2

(1:1,000, Bethyl, A300-005A), anti-vinculin (1:100,000, Sigma-Aldrich, V9131), anti-

Tubulin (1:100,000, Abcam, ab6160) antibodies were used in western blot experiments.

Antibodies

The following antibodies were used in this study: Anti-SMARCAD1

(ThermoFisher Scientific, PA5-53482), anti-SMARCAL1 (SantaCruz, sc-376377), anti-

BRCA1 (SantaCruz, sc-6954), anti-BRCA2 (Bethyl, A300-005A), anti-vinculin (Sigma-

Aldrich, V9131), anti-Tubulin (Abcam, ab6160), anti-Bromodeoxyuridine (AbD Serotec, 77

MCA2060T), anti-Bromodeoxyuridine (Becton Dickinson, 347580), Alexa Fluor 488- labelled goat anti-mouse IgG1 antibody (Thermo Fisher Scientific), and Alexa Fluor 594- labeled goat anti-rat antibody (Thermo Fisher Scientific).

Recombinant viral production and infection

Recombinant retroviruses and lentiviruses were generated by co-transfecting helper packaging vectors together with retroviral or lentiviral vectors into HEK293T cells using the TransIT-293 transfection reagent (Mirus). Virus-containing supernatants were collected 48 hrs after transfection and utilized to infect MCF10A and U2OS cells in the presence of 8 μg/ml polybrene. 48 hours after viral addition, MCF10A and U2OS cells were selected using 1 μg/ml puromycin, 10 μg/ml blasticidin, or 100 μg/mL hygromycin for 3-5 days.

Quantification and statistical analysis

Statistical differences in the DNA degradation experiments were analyzed by a

Mann-Whitney test. Statistical differences in the comet assays were analyzed by one- way ANOVA. Statistical analysis was performed using the GraphPad Prism software. In all cases: n.s. p>0.05; *p<0.05; **p<0.01; ***p<0.001; ****p<0.0001.

2.5 Figures

Figure 2. 1 Analysis of nascent DNA degradation in BRCA1/2-deficient cells upon SMARCAD1 depletion. (A) Detection by western blot of BRCA1, BRCA2, and SMARCAD1 protein levels in MCF10A cells subjected to shRNA-depletion. Tubulin levels are shown as a loading control. (B) Schematic of CldU/IdU pulse-labeling followed by 5 hours of 2mM hydroxyurea (HU) treatment (top). Representative images of CldU and IdU labelled replication tracks in HU-treated MCF10A cells expressing the indicated shRNAs. The control shRNA is a shRNA against firefly luciferase (bottom). (C) Dot plot of the IdU to CldU track length ratios for individual replication forks for HU-treated MCF10A cells expressing the indicated shRNAs. The median values are indicated by the black lines. Statistical analysis was conducted by Mann- Whitney test (n.s. not significant; ****p<0.0001). Data are representative of 3 independent experiments. (D) Dot plot of the IdU to CldU track length ratios for individual replication forks for HU-treated MCF10A cells expressing the indicated shRNAs with or without expression of wild-type (WT), K5285, T71A -mutant SMARCAD1 protein. The median values are indicated by the black lines. Statistical analysis was conducted by Mann-Whitney test (n.s. not significant; ****p<0.0001). Data are representative of two independent experiments. 79

Figure 2. 2 Supplement to Figure 2.1 — Analysis of nascent DNA degradation in BRCA1/2-deficient cells upon SMARCAD1 depletion. (A) Detection by western blot of BRCA1, BRCA2, and SMARCAD1 protein levels in MCF10A cells subjected to shRNA-depletion. Tubulin levels are shown as a loading control. (B) Dot plot of the IdU to CldU track length ratios for individual replication forks for HU- treated U2OS cells expressing the indicated shRNAs. The median values are indicated by the black lines. Statistical analysis was conducted by Mann-Whitney test (n.s. not significant; ****p<0.0001). Data are representative of two independent experiments. (C) Detection by western blot of BRCA1and SMARCAD1 protein levels in MCF10A cells subjected to shRNA-depletion and expressing either GFP, wild-type (WT), K5285, T71A, CueM, or K905R-mutant SMARCAD1 protein. Vinculin levels are shown as a loading control.

Figure 2. 3 Analysis of replication fork stalling and DNA damage accumulation in BRCA1-deficient cells upon SMARCAD1 depletion. (A) Schematic of CldU pulse-labeling followed by 2 hours of 2mM hydroxyurea (HU) treatment and subsequent IdU-pulse labeling (top). Quantification of fork restart efficiency in MCF10A cells expressing the indicated shRNAs following HU treatment (bottom). Data

arerepresented by three independent experiments as the mean and p-values calculated by a t-test with Welch’s correction (n.s. not significant; *p<0.05; **p<0.01). (B) Dot plot of alkaline comet-tail moments detected in MCF10A cells expressing the indicated shRNAs following a 100 nM camptothecin (CPT) treatment for 5 hr. Data are represented as the mean ± SEM of 75 or more comet tails per indicated condition and p-values were calculated by one-way ANOVA (n.s. not significant; *p<0.05; **p<0.01). Data are representative of two independent experiments. Representative images of the alkaline comet tails are shown. (C) Dot plot of alkaline comet-tail moments detected in MCF10A cells expressing the indicated shRNAs and siRNAs following a 100 nM CPT treatment for 5 hr. Data are represented as the mean ± SEM of 75 or more comet tails per indicated condition and p-values were calculated by one-way ANOVA (n.s. not significant; *p<0.05; **p<0.01). Data are representative of two independent experiments. Representative images of the alkaline comet tails are shown.

Figure 2. 4 Supplement to Figure 2.3 — Analysis of replication fork stalling and DNA damage accumulation in BRCA1-deficient cells upon SMARCAD1 depletion. (A) Dot plot of alkaline comet-tail moments detected in MCF10A cells expressing the indicated shRNAs with no treatment. Data are represented as the mean ± SEM of 75 or more comet tails per indicated condition. Data are representative of two independent experiments. Representative images of alkaline comet tails are shown. (B) Detection by western blot of BRCA1, SMARCAD1, and SMARCAL1 protein levels in MCF10A cells subjected to shRNA-depletion and siRNA-depletion as indicated. The ponceau is shown as a loading control.

Chapter 3: Discussion

This chapter was written with help from Alberto Ciccia.

3.1 Models for the role of SMARCAD1 at stressed replication forks

SMARCAD1 may promote resection as a fork remodeler

SMARCAD1 is a member of the SNF2-family, which contains three known fork remodelers, HLTF, SMARCAL1, and ZRANB3. While no current studies have demonstrated a fork remodeling activity for SMARCAD1, it is still possible that

SMARCAD1 promotes resection indirectly via remodeling of the fork (Figure 3.1A). In order to determine whether SMARCAD1’s role is upstream or downstream of

SMARCAL1-mediated fork remodeling, we have examined the formation of DNA breaks in response to CPT. Indeed, we found that loss of SMARCAL1 mitigated DNA break accumulation in SMARCAD1- and/or BRCA1-deficient cells (Figure 3.1C), suggesting that SMARCAL1 may act upstream of SMARCAD1 and BRCA1. CPT treatment also generates persistent replication-associated DSBs that need to be repaired through HR

[733, 734]. Given the role of SMARCAD1 and BRCA1 in DSB repair [682, 686, 710,

711], we cannot exclude the possibility that the increase in DNA break accumulation in response to CPT observed in SMARCAD1- and/or BRCA1-deficient cells is due to inability to repair DSBs by HR. Interestingly, SMARCAL1 has been demonstrated in one study to promote non-homologous end joining (NHEJ) by stabilizing the heterodimer

Ku70/Ku80 and DNA-PKcs at DSB ends [735]. While a decrease in NHEJ may restore

HR in BRCA1-deficient cells [500, 736, 737], destabilization of the Ku70/Ku80

heterodimer can lead to the repair of persistent DSBs by microhomology-mediated end joining (MMEJ) [738]. In agreement with the latter case, a recent study by Chen and colleagues has determined that 53BP1 loss does not fully rescue the viability of BRCA1 full knockout (KO) mice due to HR deficiency and elevated MMEJ levels [739].

Therefore, more studies will be needed to determine if SMARCAL1 loss increases

MMEJ or prevents the sensitivity of SMARCAD1-deficient cells to PARP inhibitors, ionizing radiation or camptothecin [681, 686].

SMARCAD1 may promote resection as a chromatin reader

SMARCAD1 also plays an important role in regulating global histone deacetylation to promote histone H3 lysine (K) 9 trimethylation (H3K9me3) and heterochromatin formation by HP1 [680]. A recent study by Higgs and colleagues, identified that H3K4 methylation was an important determinant for FANCD2-mediated nucleosome remodeling and fork protection [228]. While loss of SMARCAD1 does not influence the global amount of H3K4 methylation [680], it is possible that loss of H3K9 methylation may protect replication forks (Figure 3.1C). In agreement with this hypothesis, CHD4, a reader of H3K9me3, has already been implicated in promoting nascent DNA degradation in BRCA2-deficient cells [225, 627, 740, 741]. Therefore, an alternative explanation to our findings may be that depletion of SMARCAD1 leads to global loss of H3K9me3 and prevents CHD4 recruitment and nascent DNA degradation.

Further studies are needed to determine if SMARCAD1 loss and, specifically, loss of

H3K9me3, prevents CHD4 recruitment to nascent DNA.

SMARCAD1 may promote resection as a chromatin remodeler

While SMARCAD1 promotes fork degradation in BRCA1/2-deficient cells,

SMARCAD1 function is critical for fork restart (Figure 3.1A) suggesting that fork protection is not linked to fork restart. As such, we propose a mechanism in which

SMARCAD1 is required for moving nucleosomes at the reversed arm of the fork to mediate controlled resection (Figure 3.1B). Interestingly, 53BP1 has been shown to be recruited to stalled forks [742, 743] and has been shown to protect stalled forks from degradation [744]. This suggests that 53BP1 can prevent resection similar to its role in

HR. In this case, loss of SMARCAD1 would facilitate the resection of nascent DNA.

Models for SMARCAD1 fork restart

As discussed in Chapter 1, nucleosomes may physically pose as a barrier to fork restart. Therefore, nucleosomes need to be cleared to permit fork restoration by

RECQ1. Interestingly, it was previously reported that the chromatin remodeling activity of Fft3 is not required for fork restart in fission yeast [723]. This aspect still needs to be determined for SMARCAD1. It is possible that fork restart is mediated indirectly by

SMARCAD1’s or it’s stoichiometric binding partner KAP1’s association with chromatin modifiers. As mentioned in Chapter 1, KAP1 interacts with Suv39h1, but it also acts a scaffold for other histone modification proteins such as SETDB1 and the NuRD complex

[745, 746]. It is possible that SMARCAD1/ KAP1 acts as a scaffold to mediate chromatin modifications that promote fork restart.

Summary

Taken together, loss of SMARCAD1 prevents nucleosome remodeling on the regressed arm, thereby inhibiting fork restoration and restart, thus leading to the accumulation of DNA damage (Figure 3.2A). Loss of BRCA1 leads to extensive DNA 86

degradation and the accumulation of genomic instability (Figure 3.2B). However, because SMARCAD1 is present, forks can restart (Figure 3.2B). However, when

SMARCAD1 is depleted in BRCA1/2-deficient cells, both fork degradation and fork restart are inhibited, leading to fork collapse and DNA damage accumulation (Figure

3.2C). Ultimately, preventing fork reversal by SMARCAL1 prevents fork degradation. It has yet to be determined the effect of SMARCAL1-depletion for fork restart in

SMARCAD1-deficient cells. More studies are needed to mechanistically understand the role of SMARCAD1 at stalled replication forks and provide strong evidence for this hypothesis.

3.2 Inhibition of SMARCAD1 as a chemotherapeutic

As mentioned in Chapter 1, SMARCAD1 promotes breast cancer cell migration, metastasis, and invasion. Notably, loss of SMARCAD1 inhibited IKK-β signaling, an important regulator of the NF-κB pathway [747], and STAT3 phosphorylation [693, 694].

Both the NF-κB pathway and phospho-STAT3 has been shown to promote cellular proliferation and inhibit apoptosis in triple negative breast cancers by upregulating the transcription of its target genes [748-750]. Moreover, phospho-STAT3 drives tumor immune evasion [749]. Therefore, inhibiting SMARCAD1 may prevent cancer cell metastasis and promote anti-tumor immunity. Similarly, MMR-deficiency contributes to a durable anti-tumor response and increases immunotherapy efficacy [751, 752]. MMR- deficiency can cause insertions, deletions, base substitutions, and frameshift mutations which increases tumor mutational burden (TMB) and the likelihood of neoantigen formation [753, 754]. Strikingly, Le and colleagues demonstrate a profound difference in mutational load between MMR-deficient and -proficient tumors, as well as clinical 87

observations supporting anti-PD1 therapy for patients with MMR-deficient tumors, regardless of cancer type [755]. Other results support the notion that an increase in

TMB and neoantigen load may contribute to a durable anti-tumor response during ICB therapy. For instance, Rizvi et al. show that patients harboring somatic mutations in genes involved in MMR (such as MSH2) achieved durable benefit from PD1 treatment

[756]. Therefore, because SMARCAD1 promotes MMR [685, 688], inhibition of

SMARCAD1 will lead to MMR-deficiency and perhaps promote immunotherapy.

Moreover, loss of SMARCAD1 increases DSB formation during replication stress.

Therefore, inhibiting SMARCAD1 may activate cGAS-STING mediated-innate immune response, thus enhancing tumor immunogenicity and response of tumors to immunotherapies. Taken together, inhibition of SMARCAD1 may increase neoantigen production and promote the cGAS-STING inflammatory response while preventing

STAT3-driven immune evasion thereby enhancing tumor immunogenicity. Ultimately, inhibition of SMARCAD1 may be an effective therapeutic in combination with current immunotherapies.

3.3 Conclusions and future perspectives

Our work here has examined the pathways that maintain genomic stability and, in particular, characterized a new role of SMARCAD1 in response to replication stress.

Chapter 1 explored the replication maintenance pathways in response to genotoxic stress and the mechanisms involved in repairing a DSB. Furthermore, chapter 1 highlighted the dual roles of homologous recombination (HR) proteins and their functions at protecting the reversed arm of a stalled replication fork. In chapter 2, we demonstrated a new role for SMARCAD1 at the replication fork. The results presented 88

still need to be mechanistically clarified. While we clearly demonstrate that SMARCAD1 promotes resection at the replication fork, whether this is done by chromatin, fork, or epigenetic remodeling is still not understood. However, our study opens a new avenue of investigation on the role of chromatin context during replication stress. Overall, future studies are needed to elucidate the contribution of SNF2-family fork remodelers to tumor development along with the response to chemotherapies and immune therapies.

Ultimately, the development of small molecules able to efficiently inhibit the activity of specific SNF2 fork remodelers may open new avenues for cancer therapies and increase the therapeutic opportunities for cancer patients.

3.4 Figures

Figure 3. 1 Models for SMARCAD1’s role at a stressed replication fork. (A) SMARCAD1 can remodel stressed replication forks into a reversed fork. (B) Nucleosomes are assembled on the reversed arm of the fork and prevents resection by nucleases. SMARCAD1 subsequently remodels the nucleosomes on the reversed arm of the fork to facilitate fork restoration. (C) SMARCAD1 promotes H3K9me3 deposition leading to CHD4 recruitment. CHD4 then facilitates the recruitment and function of nucleases on the reversed arm of the fork.

Figure 3. 2 Summary of SMARCAD1-, BRCA1-, and SMARCAD1-and BRCA1-deficient phenotypes. (A) SMARCAD1-deficiency during replication stress causes fork stalling and DNA damage accumulation but display no fork degradation. (B) BRCA1-deficiency during replication stress leads to excessive fork degradation resulting in DNA damage accumulation. BRCA1-deifcient cells do not have any fork restart defects. (C) SMARCAD1 depletion in BRCA1-deficient cells restores dork protection but leads to fork stalling and DNA damage accumulation.

Appendix I: Programmable gene editing by CRISPR-Cas9

Chapter A1: Introduction

This chapter was written with help from Tarun Nambiar and Alberto Ciccia.

A1.1 RNA-guided genomic editing by CRISPR-Cas9

A1.1.1 Genome engineering technologies using protein-DNA binding

Direct manipulation of the genome is the gold standard for functional analysis.

Three crucial advancements have enabled precise genome editing and manipulation:

(1) an increased comprehension of DNA transcription and repair mechanisms, (2) innovations in genome sequencing, and (3) the development of target specific programmable endonucleases. While the development of more powerful sequencing technologies have so far generated a wealth of genomic information [757], and increases in computing power ultimately decreased the cost of sequencing [758], the increased comprehension of DNA transcription and repair mechanisms along with the development of programmable endonucleases catalyzed the gene editing revolution.

Initially, the mammalian genome was altered by the insertion of exogenously provided sequences which was discovered to occur by homology-directed integration or gene targeting involving the homologous recombination (HR) repair proteins [759-762].

Unfortunately, gene targeting suffered from low efficiency [760-762]. A key breakthrough was the observation that a double strand break (DSB) could stimulate the homologous introduction of sequences in a site-specific manner [763-767]. These studies were done by integrating a unique 18 base pair non-palindromic sequence into 92

the genome that was recognized by a yeast endonuclease, I-SceI [763-767]. After integrating the I-SceI recognition sequence into the genome, introduction of the I-SceI meganuclease and a homologous DNA donor plasmid increased integration by several orders of magnitude as compared to spontaneous gene targeting or random integration

[763-767]. However, the I-SceI endonuclease could not be modified to recognize other target sequences limiting its usefulness to studying DSB repair mechanisms.

Around the same time, studies of the Flavobacterium okeanokoites restriction endonuclease Fok I revealed that the endonuclease can be separated into a N-terminal

DNA binding domain and a C-terminal non-specific DNA cleavage domain [768]. The modular nature of the Fok I endonuclease led to the next logical step that the endonuclease can be fused to different DNA binding domains to direct the nuclease to a specific site in the genome [769, 770]. Zinc finger proteins are functionally diverse but contain a specific modular DNA binding motif called the zinc finger motif. Each tandem array of the zinc finger motif can interact with a specific triplet of nucleotides in the DNA suggesting that zinc finger motifs can be arranged together to recognize any DNA sequence [771-774]. This chimeric Fok I endonuclease fused to zinc finger motifs was called a Zinc Finger Nuclease (ZFN). In order to efficiently increase gene targeting through the generation of a DSB, two ZFNs with inversely oriented binding domains and no linker sequence are needed [775]. However, ZFN bioengineering is extremely difficult and imperfect specificity of certain ZFNs to off-target sites were found to be cytotoxic and mutagenic limiting its therapeutic usefulness [776, 777].

Similar to zinc finger proteins, transcription activator-like (TAL) effector protein motifs from the plant pathogen genus Xanthaomonas, bind specific DNA sequences 93

[778]. Unlike zinc finger motifs, the TAL effector protein contains individual TAL repeats that bind to a single base pair as determined by the repeat variable diresidue (RVD) code or two specific amino acid residues that interact with the DNA by specific interactions or by special accommodation of the base [778-780]. A TALEN is a chimeric endonuclease consisting of the Fok I nuclease domain fused to a series of TAL repeats to mediate the binding of Fok I to a target site [781-783]. Therefore, bioengineering of

TALENs is easier than with ZFNs. Furthermore, TALENs have higher specificity and lower cytotoxicity than ZFNs [783]. A major limitation to both TALENs and ZFNs are their reliance on protein-DNA interactions. The variability in protein affinity and activity based from the DNA binding motifs fused to the proteins necessitates additional in vitro validations and the construction of multiple TALENs or ZFNs for a given genomic sequence limiting high-through-put screening and precision medicine applications.

A1.1.2 Re-purposing the RNA-guided adaptive response, CRISPR/Cas, for genome editing

An alternative to TALENs or ZFNs, was first discovered as an adaptive immune response in prokaryotes to invading pathogens called the CRISPR/Cas system. This adaptive immune response consists of the integration of foreign nucleic acids into the genome by RNA-guided endonucleases to store a record of the invasion and mount a response upon re-exposure [784-787]. Integration of the viral or foreign DNA into the genome occurs in a polar manner with the newest sequence added to the first position of the repetitive element called CRISPR (Clustered Regularly Interspersed Short

Palindromic Repeat) [786, 788-790]. The CRISPR array consists of variable sequences called spacers that correspond to the captured viral or plasmid DNA and a repeat 94

sequence that is conserved for the given CRISPR array with some repeats predicted to generated a stable RNA hairpin [786, 788-792]. The cognate spacer sequence native to the foreign element is called the protospacer, Adjacent to the CRISPR locus is a variable cassette of genes called the CRISPR associated genes (cas) which encode functional Cas proteins involved with RNA processing and DNA cleavage [784, 787,

793, 794]. In order to confer immunity, the Cas proteins incorporate the foreign DNA into the CRISPR locus and subsequently transcribe them as a CRISPR RNA (crRNA) template for targeted destruction of the protospacer at either the DNA or RNA level

[792, 795-798]. Of these Cas proteins, Cas 1 and Cas2 are highly conserved through the diverse CRISPR-Cas systems [799]. The Cas1 and Cas2 proteins are responsible for integrating the short foreign pieces of DNA into the CRISPR locus as spacers a process known as spacer integration [800-802]. Due to its diversity, the classification of the CRISPR-Cas system into two classes is based on the organization of the CRISPR-

Cas locus, the types of cas genes, sequence similarity between the Cas proteins, the structure of the CRISPR array itself, and the phylogeny of Cas1 [799, 803-806]. Class I

CRISPR-Cas systems (types I, III, and IV) employ a multi-protein effector complex composed of multiple Cas proteins that assemble into a complex containing different numbers of Cas subunits and participates from pre-crRNA processing to target cleavage [803, 804, 807, 808]. Class II systems (types II, V, and VI ) employ a single multidomain effector protein [804, 809]. The effector module of the CRISPR-Cas system binds to the transcribed spacer and repeat sequence, promotes complementary base pairing with the foreign DNA or RNA, and finally cleavage of the phosphodiester bond

[799, 810]. Spacer integration for Types I, II, IV, V, and VI rely upon a 2-6 base pair 95

sequence adjacent to the protospacer, called the PAM (protospacer adjacent motif) [or protospacer flanking sequence, PFS, for type VI], that is recognized by the integration module in a base-specific manner [810-815]. The integrated spacer and mature crRNA are, therefore, complementary to the protospacer but lacks the PAM motif. PAM recognition, is also, a critical part of the effector targeting and is required for ATP- dependent strand separation and crRNA-DNA heteroduplex formation for target interference [816]. The lack of the PAM in the host genome and requirement for effector cleavage prevents auto-cleavage and autoimmunity [817]. Because Class II systems only use a single multidomain protein, it became an attractive tool for genome editing applications. Notably this class is exemplified by the type II Cas9 protein that is currently the most widely used genome editing tool [818].

A1.1.3 The Cas9 effector as a promising molecule for genomic engineering

The Streptococcus pyogenes (S. pyogenes) type IIA CRISPR-Cas system consists of 4 Cas genes (Cas 1, Cas 2, Cas9, and Csn2), two non-coding crRNAs, a trans-activating crRNA (tracrRNA) and the spacer/repeat array (pre-crRNA) [819, 820].

Unlike other CRISPR-Cas systems, type II systems require the tracrRNA for crRNA maturation and crRNA-guided mobile element cleavage by Cas9 [820, 821].

Specifically, the entire CRISPR array is transcribed as a long continuous RNA transcript, pre-crRNA, which is processed by the tracrRNA in complex with RNase III and Csn2 to generate mature crRNAs [820, 821]. The mature crRNAs then base pairs with the repeat sequence of the tracrRNA enabling Cas9 interaction with the crRNA

[820, 821]. Seminal work from the Charpentier and Doudna labs demonstrated that the

mature crRNA and a partial tracrRNA can be fused into a chimeric RNA called the single guide RNA (sgRNA) to target the Cas9 to DNA [821].

Cas9 Activity

Cas9 is a large bi-lobed RNA-activated protein that promotes DNA cleavage in an RNA-directed manner [822-824]. The N-terminal half of the protein contains a RuvC- like domain that is part of the Nuclease (NUC) lobe and a large α-helical arginine rich region called the Helical Recognition (REC) lobe while the C-terminal half contains several RuvC-like domains and an HNH-domain contributing the rest of the NUC lobe as well as a PAM-interaction site (PI) [822, 823]. The RuvC active site cleaves the non- target strand (not complementary to the crRNA sequence) while the HNH-active site undergoes an RNA-dependent conformational change to cleave target strand [819, 821-

827]. The Cas9-RNA complex searches DNA through three-dimensional diffusion and rapidly dissociates from non-PAM sites and non or partially complementary-crRNA sites

[824, 828]. Cas9 recognizes the PAM sequence through major groove interactions between the protein and the DNA facilitating R-loop formation and activating both nuclease domains enabling target DNA cleavage [823, 828, 829].

Genomic engineering using Cas9

The S. pyogenes Cas9 (SpCas9) was the first to be reconstituted in mammalian cells through the expression of a codon optimized Cas9 bearing a nuclear localization signal and the chimeric sgRNA [830-832]. Altering the spacer, seed, sequence in the sgRNA enables Cas9 to be re-directed to any genomic target adjacent to the appropriate PAM making [830]. Different Cas9 homologs recognize unique PAM sequences. For example, the PAM-motif required by SpCas9 is constituted by any base 97

adjacent to two guanines (NGG) while the homolog of SpCas9, SaCas9 from S.aureus, recognizes a sequence of any base adjacent to a guanine (G) next to two base purines

(A/G) and a thymine (T) or NNGRRT PAM [833]. Therefore, using different homologs of

Cas9 enables expanded genetic sequence targeting capabilities. Furthermore, unlike

TALENs and ZFNs, several sgRNAs can be introduced to target different genomic loci simultaneously making it applicable to multiplex screening methodologies [830, 834-

836]. Because the Cas9 molecule relies upon RNA-DNA hybridization dynamics and subsequent conformational changes for accurate gene targeting, the reliability of the

CRISPR technology has been hampered by unintended cleavage of sites with mismatches to the sgRNA [837-840]. As such, there are ongoing efforts to detect and reduce off-target cleavage sites. Strategies to detect off-target sites map Cas9 binding sites or Cas9 activity such as SITE-seq, DIG-seq, CIRCLE-seq, GUIDE-seq, BLISS, and DISCOVER-seq [839, 841-847]. On the other hand, strategies to reduce off-targets include truncating the sgRNA to weaken the sgRNA-DNA duplex stability at off-targets, generating enhanced specificity Cas9 mutants, and computational modeling to predict off-targets of a given sgRNA for optimized guide selection [848]. Taken together, Cas9 is a programmable nuclease that is currently being optimized for precise genome targeting.

Repair of a Cas9-induced break

In addition to homology-directed repair and NHEJ, DSBs can be repaired by an alternative end joining pathway known as microhomology-mediated end-joining (MMEJ).

MMEJ is a DSB repair pathway that occurs in M and early S-phases and shares mechanistic aspects of both HDR and NHEJ [849]. The MMEJ pathway joins broken 98

DSB ends without templated DNA synthesis similarly to NHEJ but requires end resection, like HDR, to expose microhomologous sequences and ultimately generates repair products with sequences removed, deletions, between the microhomologies [849,

850]. Cas9-induced repair products without a homologous sequence are largely determined by the pathway used and Cas9 cleavage pattern. As such, +1 insertions are largely attributed to NHEJ-mediated repair of staggered cleavage products and deletions are due to MMEJ-mediated repair [851, 852]. Therefore, the repair pathways that underlie genome editing after a Cas9-induced break will dictate the types of edited products generated.

A1.2 CRISPR applications

The recent explosion in the development of CRISPR-based technologies [795,

818] has produced an expansive toolbox that enables more comprehensive and systematic assessments protein function, contribution to genome stability, and disease

(Figure A1.1). Because the Cas9 endonuclease generates a DSB, the addition of a homologous sequence can mediate the precise editing of a loci through HDR. HDR can be used to disrupt gene expression (knockout) by the expression of a gene disruption cassettes (Figure A1.1). On the other hand, mutation of either the HNH or RuvC domain can generate a nickase Cas9 (nCas9), which only cleaves a single DNA strand, that can be utilized similarly to precisely edit a locus using HDR by targeting the nickase with dual offset sgRNAs in a manner analogous to ZFNs or TALENs or by a single nCas9 through non-canonical HDR independent of CtIP [821, 831, 853-855]. Paired nCas9 or nCas9 alone can reduce off-target activity while a paired nCas9 has similar efficiencies as the Cas9 nuclease [821, 831, 853, 854, 856]. Mutating both nuclease 99

domains generates a nuclease dead Cas9 (dCas9) that retains its RNA-guided DNA- binding activity allowing site-specific genetic and epigenetic regulation, protein interaction, and imaging applications [857-866]. In particular, transcriptional activators,

VP64, and repressors, the KRAB domain, can be directed to target gene promoters and enhancers through their fusion with dCas9, to either induce expression, CRISPR- activation (CRISPRa), or prevent transcription, CRISPR-interference (CRISPRi) (Figure

A1.1) [863, 867, 868].

CRISPR-Cas9 has enabled forward genetic screening at a genome-wide scale, ideal for studying biological systems in an unbiased manner. Pooled CRISPR screens have enabled the ability to screen thousands of genes in an unbiased manner in only one experiment by delivering a library of sgRNAs such that a single cell receives a single sgRNA and measuring viability, drug survival, or marker-based selection (Figure

A1.1) [835, 868-874]. However, pooled screens are not compatible with identifying mild phenotypes [868]. Unlike pooled screens, arrayed screens use sgRNAs arranged in a multi-well format to examine individual perturbations, but often are more labor intensive

(Figure A1.1) [875-877]. Combinatorial screens enable research into how genes function together in networks. These use dual sgRNAs that are paired with either control sgRNAs (non-targeting/scrambled) so that the effect of the individual sgRNA can be compared to the double sgRNAs to map relevant interactions on viability or survival

[878-881]. This screening method used lentiviral vectors containing the same promoter and repetitive elements, which are prone to recombination leading to a decrease in combinatorial screening efficiency [836, 878, 880]. An alternative approach that was developed uses two orthologous Cas9 molecules (S. pyogenes and S. aureus) along 100

with orthologous promoter sequences to decrease recombination and enable more robust combinatorial screening [881]. Additionally, combinatorial screens enable more intricate combinations including CRISPRa and CRISPRi cell expression platforms with the knockout system to study multifaceted interaction networks [882].

Recent advances made in applying single-cell analyses, such as in situ sequencing in intact cells, to pooled CRISPR screens allow for the acquisition of richer content for phenotypic characterization [883]. As such, differences in transcriptional and chromatin state under different conditions can be identified by coupling CRISPR screening with single cell RNA-seq (Perturb-seq, CROP-seq, TAP-seq) and chromatin accessibility studies (Perturb-ATAC) (Figure A1.1) [884-887]. Importantly, the rich content generated by these particular transcriptomic techniques has demonstrated utility towards characterizing two-way and higher order genetic interactions [878, 884].

The use of high-throughput saturation mutagenesis screens has the potential to uncover novel functional domains, regulatory elements and separation-of-function mutations as well as classify the pathogenicity of variants that occur in the human population. Therefore, CRISPR-mediated HDR can be used to systematically interrogate protein structure-function relationships (Figure A1.1). For example, a saturation mutagenesis screen has already been conducted for BRCA1 using CRISPR- mediated HDR to introduce 4000 variants into 13 coding exons [888]. This proof-of- concept study identified hundreds of novel non-functional variants of BRCA1 and allowed for the potential to predict pathogenicity of variants that were previously of unknown significance. Taken together, CRISPR-Cas9 systems are a useful research tool for functional genomics. Alternatively, Cas9-mediated genomic editing has 101

therapeutic potential to correct or install mutations associated with human diseases for the modeling and treatment of genetic disorders.

A1.2.1 Modulating homologous recombination for increased targeting efficiency

Increasing the efficiency of CRISPR-mediated genomic editing is critical for its therapeutic potential. HDR-mediated repair of a Cas9-induced break can lead to the incorporation of the mutations of interest or precise edit using an exogenous DNA- template, or donor template, making it a versatile choice for modeling or repairing mutations associated with human diseases [889, 890]. However, because HDR is restricted to S- and G2-phases of the cell cycle, efficient HDR is limited to actively dividing cells [891-896]. Additionally, HDR is a considerably slower process as compared to MMEJ and NHEJ and under most conditions NHEJ is more efficient than

HDR [897, 898]. Therefore, even in cultured cell lines or actively dividing cells, HDR efficiency is modest [898]. Several approaches have been used to improve HDR- mediated efficiency including manipulating the cell cycle, small molecules to decrease

NHEJ or enhance HDR, and increasing donor template availability. Lin et al. showed that timed delivery of Cas9 into M-phase synchronized cells using the microtubule inhibitor, nocodazole, enhanced HDR cell editing [899]. Furthermore, fusion of the Cas9 protein with the N-terminal half of the DNA replication inhibitor protein, geminin, led to the expression of the Cas9 protein in a cell-cycle dependent manner and increased

HDR rates [900-902]. Alternatively, inhibiting NHEJ by small molecules such as the ligase IV inhibitor SCR7, and DNA-PKc inhibitors NU7441 and KU-0060648 enhanced

HDR rates and increased genome editing efficiency [903-905]. Furthermore, NHEJ can be prevented by the ectopic expression of a dominant negative mutant form of the 102

NHEJ factor 53BP1 or fusion to Cas9 leading to improved genome editing efficiencies

[906, 907]. Moreover, ectopic expression of RAD52 or an engineered variant of RAD18 prevented 53BP1 binding and increased editing efficiencies [907, 908]. Alternatively, enhancing HDR can also increase HDR efficiencies. As such, the HDR enhancer RS-1 stimulates RAD51 activity to improve HDR efficiency [909, 910]. Finally, increasing the donor template availability by tethering the homologous donor sequence to the Cas9 increases HDR efficiencies [911, 912]. In this case, Cas9 is fused to the Porcine

Circovirus 2 Rep protein which is an HUH endonuclease that can form a covalently bond with unmodified single-stranded DNA (ssDNA) [911, 912].

Despite progress to increase HDR efficiencies, these strategies so far have the potential to be disruptive to the cell by altering DNA repair kinetics or cell cycle progression and are therefore limited to research applications. Moreover, several of these approaches would not be able to enhance editing efficiency in postmitotic cells.

One common limitation to both HDR and NHEJ approaches is the reliance on the generation of a DSB. DSBs are toxic DNA lesions that can cause genomic rearrangements, activate DNA damage checkpoints, and induce cell death [913-916].

Recently, large mono-allelic deletions or insertions have been identified after HDR- mediated CRISPR editing that would be disruptive for clinical applications [917].

Therefore, approaches that can edit the genome without inducing a DSB or relying on cell cycle progression would be beneficial for therapeutic applications.

A1.2.2 Programmable base editing

Single nucleotide polymorphisms (SNPs) or single base changes are the most frequent changes to the genome and half of known inherited disease mutations come 103

from a single point mutation that causes an amino acid substitution leading to alterations in protein folding or function called a missense mutation [918-921]. In fact, over half of all pathogenic SNPs are cytosine (C) or guanine (G) (C•G) to adenosine (A) or thymine (T) (A•T) mutations resulting from the spontaneous formation of uracil and thymine by hydrolytic deamination of cytosine or 5-methylcytosine [918, 922, 923].

Consequently, correcting pathogenic SNPs would treat a substantial fraction of genetic diseases. Furthermore, DSBs are toxic lesions that can lead to genomic rearrangements or cell death so an alternative to HDR or NHEJ-mediated genome editing would be beneficial [913-915, 924]. Therefore, CRISPR-dependent methods have been developed that can directly change DNA bases without generating a DSB

[922, 925-932].

CRISPR-dependent base editing consists of a dCas9 or nCas9 fused to a cytosine deaminase, APOBEC1 or AID, or an engineered tRNA adenine deaminase

(TadA) called cytosine base editors (BE) or adenine base editors (ABE) (Figure A1.2)

[922, 925-932]. Cytosine deamination converts cytosine to uracil (U) while adenine deamination converts adenine to inosine (I) (Figure A1.2) [933, 934]. Base excision repair (BER) is the repair process that typically fixes the aberrant incorporation of U into the genome through recognition and removal by the uracil DNA-glycosylase (UDG)

[935]. Therefore, a more efficient BE is generated by fusing dCas9 or nCas9 to a uracil

DNA glycosylase inhibitor (UGI) from the bacteriophage PBS1 (Figure A1.2A) [925]. In contrast to BEs, I is not as easily recognized and repaired by the BER machinery so

ABEs do not require to be fused to an inhibitor (Figure A1.2B) [922]. The mismatch repair (MMR) pathway is responsible for recognizing mismatched bases in newly 104

synthesized DNA, in particular misincorporation of U, to increase replicative fidelity

[936]. Therefore nCas9 is optimal for base editing since mismatch repair favors the repair of the nicked strand, or the non-edited strand, increasing base editing efficiencies and the chances that the C will be edited to a T or product purity [925]. The resulting

G:U or I:T mismatches are then converted to A:T and G:C respectively by DNA synthesis (Figure A1.2) [922, 925]. Therefore, the CRISPR-dependent cytosine base editor BE3 is a fusion of a rat APOBEC1, a UGI, and the spCas9-D10A nickase mutant

(Figure A1.2A) [925]. ABE7.10, on the other hand, is a fusion of two evolved TadAs to the spCas9-D10A nickase mutant (Figure A1.2B) [922]. The BE3 was then improved for efficiency and product purity by lengthening the linker between nCas9 and APOBEC1 to

32 amino acids making a new chimera called BE4 [932]. Further improvements were made upon ABE7.10 and BE4 by codon optimization and ancestral sequence reconstruction to generate the optimal base editing tool box of AncBE4max, BE4max for

BEs and ABEmax for ABEs [929]. Kim et al. increased the scope of the genome targetable by BE3 by developing new BE chimeras containing either SaCas9, an engineered SaCas9, or an engineered SpCas9 variant with altered PAM specificities

[937]. This expansion in BE3 variety increased the number of C to T (G to A) pathogenic mutations targetable by BE from 34% to 66% [937]. Finally, Grünewald et al. engineered a dual ABE and BE called SPACE that is capable of generating both A to G and C to T transitions on the same target [938]

Unlike HDR-mediated genomic editing, base editing does not have a cell cycle dependency or the need for a homologous donor. So far base editors have been demonstrated to edit the genome of postmitotic sensory cells, in vivo somatic 105

hepatocytes of a mouse liver, and human organoids [930, 939, 940]. Furthermore, base editing has been used to edit the genomes of mice, plants, and human cells [930, 941-

946]. In particular, base editing has been used to treat mouse models of amyotrophic lateral sclerosis (AML) and duchenne muscular dystrophy (DMD) [945, 946].These base editors can alter any C or A within a window 13-17 bases away from the PAM meaning the presence of more than one C or A within the activity window can result in the modification of multiple bases leading to potentially undesirable base substitutions [922,

925, 929, 932]. There have been attempts to improve the base specificity of the base editors but so far only BEs have modified to have a tighter activity window (from 5 to 1-2 nucleotides) [937]. While potentially undesirable base modifications are not favorable for precise base editing, it does not hinder mutations that can disrupt gene expression, nonsense mutation formation, which is the focus of Chapter 2.

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A1.3 Figures

Figure A1. 1 The CRISPR-based technology toolbox. Overview of the tools and technologies used for gene editing and screening. The Cas9 protein is an RNA-guided DNA endonuclease that generates double strand breaks using two endonuclease domains. An engineered Cas9 can be targeted to any site in the genome using a guide RNA (dark blue) to interrogate the genome. Additionally, the catalytic activities of these proteins can be modified without effecting RNA-guided DNA-targeting. As such a Cas9 protein with one endonuclease domain inactivated (nCas9) or both (dCas9) can be fused to different effector proteins to mediate different forms of DNA editing (Base editing or Prime editing) or transcriptional modulation. Which editing technology tool is used depends on the editing strategy. Homology-directed repair requires the addition of DNA donor templates. Current approaches for saturation mutagenesis employ base editors. Gene knockdown by transcriptional repression by CRISPRi. Alternatively, Gene knockout can be accomplished by inducing a stop codon by base editing (iSTOP/ CRISPR-STOP) or integration of a stop cassette/ indel formation leading to a nonsense mutation as mediated by Cas9 cleavage. These strategies can then be employed to interrogate the genome by high-throughput screening approaches (as a pooled screen or an arrayed screen). Recent advances in in situ sequencing of intact cells, imaging, and single cell sequencing has made it possible to identify and couple phenotypes to loss-of-function or structure-function analyses. This overview is not comprehensive and there are several CRISPR tools and strategies not highlighted here. 107

Figure A1. 2 Schematic of the base editors. Overview of the structure and reactions catalyzed by the base editors. (A) Cytosine base editors (BEs) use a catalytically dead Cas9 (dCas9) for base editors BE1 and BE2 or a nickase Cas9 (nCas9) for BE3 which is fused to a cytidine deaminase, APOBEC1 or AID, and uracil DNA glycosylase inhibitor (UGI) (BE2, BE3, and BE4). The cytodine deaminase deaminates cytosine (C) converting it into a uracil (U), which during DNA replication is misread by the polymerase as a thymine (T) and ultimately repaired as a T, leading to the transition of guanine (G) to adenine (A). (B) Representative schematic of the adenine base editor ABE7.10 composed of an nCas9 fused to 2 engineered tRNA adenine deaminases (TadAs). TadA deaminates A converting it into an inosine (I), which during DNA replication is misread by the polymerase as a G and ultimately being repaired to a G leading of the transition of T to C.

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Chapter A2: Inducing stop codons by CRISPR-mediated base

editing

Portions of this chapter have been adapted from:

Billon P*, Bryant EE*, Joseph SA, et al. CRISPR-Mediated Base Editing Enables

Efficient Disruption of Eukaryotic Genes through Induction of STOP Codons. Mol Cell.

2017; 67(6): 1068-1079.e4. doi:10.1016/j.molcel.2017.08.008.

A2.1 Overview

The introduction of a stop codon prematurely into a gene ORF, or a nonsense mutation, can lead to the synthesis of a truncated protein with potentially deleterious gain-of-function or dominant negative activity and is often associated with human disease (11.2% of all mutations known to cause or associate with inherited human diseases) [947-953]. Notably, nonsense mutations are observed in the tumor suppressor genes of 10%-30% of hereditary cancer patients and represent 12% of sporadic non-silent mutations in tumor suppressors reported by the catalog of somatic mutations in cancer (COSMIC) [953-957]. However, strategies to model these disease- associated nonsense mutations have been limited.

Nonsense mediated decay (NMD) is an mRNA quality control pathway that eliminates abnormal mRNA transcripts containing premature stop codons to prevent deleterious protein products [958-960]. If both alleles of a gene contain a nonsense mutation, then NMD-mediated decay of the mRNA would generate a knockout of the

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protein expression. As such, NMD has been utilized to generate efficient gene knockouts through the generation of indels or frameshifts mediated by NHEJ from a

Cas9 induced break and has been utilized in several CRISPR screens to interrogate gene function and uncover new genes responsible for survival, proliferation, and drug resistance [961-963]. Therefore, the ability to efficiently disrupt gene expression without requiring a cytotoxic DSB could open new avenues of study in new model organisms or postmitotic cells.

Here I describe an application of CRISPR-mediated base editing for gene disruption studies published by Pierre Billon and Eric Bryant [964]. Experiments were performed by Pierre Billon with help from Tarun Nambiar, and Samuel Hayward. Eric

Bryant performed the genome-scale analysis and wrote the R packages

(https://github.com/CicciaLab/iSTOP) and (https://github.com/CicciaLab/iSTOP-paper) to reproduce the figures and computational analysis done and enable iSTOP detection respectively. In this study, we show that the CRISPR cytosine base editor (BE3) can efficiently inactivate human genes through the induction of stop codons (iSTOP) by precisely converting four codons (CAA, CAG, CGA, and TGG) into the stop codons

(TAG, TAA, or TGA) and demonstrate that iSTOP-mediated editing in sgRNA- transfected cellular pools or clones can be monitored by a restriction fragment length polymorphism (RFLP) assay using restriction sites targeted by iSTOP. Remarkably,

94%-99% of genes analyzed in eukaryotic genomes can be targeted by multiple sgRNAs for iSTOP (sgSTOPs) while 70% of targetable sites corresponding to 98% of human genes can be monitored by the RFLP assay. We establish that iSTOP can be used to model over 32,000 nonsense mutations observed in human cancer making 110

iSTOP a valuable gene disruption tool to study gene function and model human disease. To facilitate the use of iSTOP I created an online database

(http://www.ciccialab-database.com/istop) of all sgSTOPs for the five BE3 variants

(VQR-BE3, EQR-BE3, VRER-BE3, saBE3, and saKKH-BE3) in humans and sgSTOPs for eight eukaryotic species 3,483,549 targetable gene coordinates expanding the targeting range recently described for iSTOP-based approaches [965].

A2.1.1 CRISPR-mediated base editing efficiently disrupts genes through iSTOP

CRISPR-mediated base editing converts C•G base pairs to T•A [925, 937].

Therefore, base editors (BEs) can convert four different codons (CAA, CAG, CGA, and

TGG) which encode for three amino acids (glutamine [Gln], arginine [Arg], and tryptophan [Trp]) into the TAA, TAG, and TGA stop codons. While CAA, CAG, and CGA can be converted to stop codons when targeted by BEs on the coding strand, TGG can be converted into a stop codon if targeted on the non-coding strand. Therefore, the conversion of a codon into a nonsense codon through base editing has been called iSTOP for induction of stop codons through sgRNAs that target the site for editing

(sgSTOPs). To efficiently monitor iSTOP-mediated edting, targeting cytosines within the recognition sequence of restriction enzymes that if changed could render those restrictions sites refractory to digestion would enable detection of the edit by a RFLP assay. Additionally, iSTOP-mediated editing can create new restriction sites, resulting in the generation of new DNA fragments upon digestion.

In order to generate human knockout cell lines, a previously described marker- free co-selection strategy that increases editing efficiency was used in combination with the sgSTOPs. By targeting the sodium-potassium channel ATP1A1, which when 111

mutated renders cells resistant to oubain treatment [966], percent editing by iSTOP increased by 14.8% in HEK293T cells. Moreover, screening the co-sgSTOP transfected clones by RFLP analysis determined that 52.6% of the 19 clones were heterozygously edited and 10.5% were homozygously edited establishing iSTOP as an efficient approach to generate human knockout cell lines and compatible with co-selection strategies.

To determine if whether this approach can be generalized to disrupt genes on a genome-wide scale, sgSTOPs were identified for all 69,180 ORFs in the human reference genome GRCh38 by using CDS coordinates from the UCSC genome browser to identify all CAA, CAG, CGA, and TGG codons located the correct distance from the

PAM for all validated BE3 variants [925, 937]. This analysis revealed that 62.5% of the

CAA, CAG, CGA, and TGG codons are targetable by iSTOP in the human reference genome, enabling the possibility to convert 523,340 codons into stop codons.

Furthermore, over 80% of ORFs targetable within the first 20% of their sequence,

99.2% of ORFs targetable within the first 100 codons, and 94.7% of ORFs targetable at a position predicted to cause NMD [967]. To aid in selecting sgSTOPs with low off- target sites, we performed a genome search for sequences similar to each sgSTOP and observed that 74% of NGG PAM-based sgSTOPs mapped uniquely in the genome suggesting that a majority of the NGG-based sgSTOPs will have limited to no off- targets. To determine whether the RFLP assay can monitor the efficiency of editing genome-wide, we identified restriction sites of enzymes targetable by iSTOP. This analysis determined that over 70% of targetable sites (corresponding to 98% of human genes) can be monitored by the RFLP assay and predicted that BsrI and BfaI are the 112

two most common restriction enzymes for detecting iSTOP-mediated edits. Collectively, this analysis demonstrates that iSTOP is a promising tool for genome-wide disruption studies in human cells and that RFLP analysis allows for rapid assessment of base editing activity.

Nonsense mutations are frequently associated with human disease and cancer with nonsense mutations accounting for 4%-5% of all observed mutations in cancer. To determine if iSTOP can be used to model cancer-associated nonsense mutations due to cytidine deamination, we determined the prevalence of C to T and G to A base transitions observed in the COSMIC database for the four codons CAA, CAG, CGA, and

TGG. This analysis revealed that 61% of the cancer-associated nonsense mutations could be reproduced using iSTOP when considering each mutation site once suggesting that iSTOP can be used to efficiently model cancer-associated nonsense mutations.

Several eukaryotic species are used as model organisms to understand biological processes. To determine whether iSTOP can be used for genome-wide studies in other eukaryotic species, sgSTOPs were identified for all ORFs of seven eukaryotic model organisms ranging from yeast (S. cerevisiae) to mice (M. musculus).

From this analysis we identified 2,490,476 codons that can be targeted in these species and we found that the percentage of ORFs that can be targeted by iSTOP is between

97% and 99,7% demonstrating that is method is robust for almost any ORF from various model organisms. While iSTOP can target 99.7% of human genes, this meant that 68 genes (0.35%) could not be targeted due to unavailable CAA, CAG, CGA, and

TGG codons or PAM sequences. However, 24/68 of these non-targetable human genes 113

have eukaryotic orthologs that can be targeted by iSTOP, therefore allowing the study of these genes in model organisms.

A2.1.4 The iSTOP database

To facilitate the use of this method, I developed a database that contains information on the number and percentage of alternatively spliced isoforms targeted by each sgSTOP, a prediction of nonsense-mediated decay, counts of putative off-target sites for each sgSTOP and a list of restriction enzymes that can be used to monitor iSTOP-mediated editing for all 3,483,549 gene coordinates. An interface for convenient search of sgSTOPs is available on web and mobile platforms at http://www.ciccialab- database.com/istop (Figure A2.1).This website was developed using a Bitnami stack with a python-based Django web framework that interfaces with the SQL databases generated through the analyses. The source code for this website is available at https://github.com/Sarah-Joseph/iStopWebsite. This website has recently been used to design a library of sgSTOPs [968].

A2.2 Discussion and perspectives

Overall iSTOP is a promising tool for CRISPR-mediated creation of stop codons to facilitate gene disruption and enable the generation of knockout cell lines.

Additionally, we describe an RFLP assay for the easy monitoring of base editing efficiency in cell populations or clones. Finally we provide a convenient resources for searching and selecting sgSTOPs including useful annotations such as off-target propensity, NMD prediction, percentage of isoforms targeted, and a list of restriction

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enzymes that can be used to monitor editing as well as software to design base-editing guides for any annotated genome and any PAM.

A2.2.1 Advantages over DSB-mediated disruption strategies

Given that iSTOP causes the disruption of genes and does not mediated precise genome editing through the creation or correction of missense gene variants, iSTOP is not affected by undesirable cytosine deamination in the BE3 window of activity.

Furthermore, unlike DSB-mediated or CRISPR-mediated base editing techniques, iSTOP induces the exact conversion of four codons into a stop codon. This precision combined with the fact that there are over 3 million targetable codons in the eight eukaryotic species permitting up 97%-99.7% of eukaryotic genes makes iSTOP compatible with genome-scale analyses in multiple model organisms. Additionally, iSTOP does not rely on the formation of DSBs whose formation during organismal development is particularly deleterious thereby potentially facilitating the generation of knockout organisms [969]. Furthermore, iSTOP disrupts genes without relying on

NHEJ-induced frameshift mutations or indels which create a mosaic population. In fact,

CRISPR-mediated base edited mice have been demonstrated to lack mosaicism [970].

Furthermore, iSTOP does not require donor molecules necessary for HDR making iSTOP a simpler disruption strategy and particularly attractive for plant bioengineering in which the use of DNA donor templates is hampered by technical and legislative obstacles [971]. Therefore, recent studies have demonstrated the CRISPR-dependent base editing approaches are highly efficient in rice, wheat, tomatoes, and corn [941-

944].

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A2.2.2 Potential iSTOP applications

Traditional CRISPR-strategies typically interrupt gene structure to study gene disruption. For ORFs that contain both coding and non-coding functions, this will make disrupting these loci extremely difficult. For example, the gene ASCC3 encodes for long and short isoforms as well as a non-coding RNA that have opposite effects on transcription recovery after DNA damage in which the short isoform promotes transcription recovery dependent on its non-coding RNA rather while the long isoform prevents recovery [972]. iSTOP will provide a possible strategy to separate the of coding from non-coding functions of genes. In addition, iSTOP could be employed to incorporate modified or non-natural amino acids into proteins using tRNAs that suppress the newly introduced STOP codons [973]. Moreover, by enabling the modeling of nonsense mutations at a genome-wide level, iSTOP could allow genome-wide studies to investigate eukaryotic gene functions and facilitate the identification of pathogenic variants in cancer through large-scale functional studies of cancer- associated mutations. Recent work utilizing CRISPR-mediated generation of stop codons (CRISPR-STOP) has indeed confirmed the promising potential of iSTOP-related approaches for pooled screening applications [965]. In fact, Zhu and colleagues have proposed using iSTOP in combination with their internal barcode (iBAR) sequences for

CRISPR-pool disruption screens as a better choice for ensuring lower false-positives and false-negatives by increasing the multiplicity of infection (MOI) [835].

Moreover, ABEs have been utilized to silence gene expression by mutating the start codon, a method called i-Silence [974]. It was demonstrated that i-Silence successfully edited the PD-1 gene in mouse embryos for the development of a PD-1 116

knockout mouse with the highest editing efficiency of 90.4% [974]. Similarly to iSTOP, i-

Silence enables genome-wide gene disruption through the substitution of the start codon of about 92.4% of human genes (17,804 genes) as compared to 99.7% by iSTOP and can model 147 human disease [974]. Additionally, a total of 16,730 genes can be targeted by both strategies [974]. Taken together, our work establishes CRISPR- mediated induction of stop codons as a robust and efficient gene disruption technology compatible with genome-wide studies to investigate eukaryotic gene functions, aid plant bioengineering, and model human diseases.

A2.2.3 Limitations to iSTOP

Although iSTOP can mediate substitutions for 523,340 codons, iSTOP is predicted to have reduced editing efficiency at 24% of targetable sites due to the restrictive rules for designing sgSTOPs. Unlike canonical Cas9-mediated editing, which can be programmed to generate DSBs adjacent to any PAM sequence, sgSTOPs require the presence of CAA, CAG, CGA, and TGG codons and a PAM located 13–17 bps away from the targeted base(s). Although ∼60% of all available CAA, CAG, CGA, and TGG codons are targetable by iSTOP, the remainder cannot currently be targeted due to the absence of a nearby PAM sequence. An additional limitation of iSTOP is due to the inability of BE3 to efficiently edit cytosines with a G on the immediate 5’ side, as recently shown [925]. Recently, the engineering of a new SpCas9 variant, called SpRY, created an RNA-guided nuclease with robust activity at NRN sites opening the door for nearly PAMless editing [975]. Further engineering led to the creation of SpCas9-NRRH, spCas9-NRCH, and SpCas9-NRTH compatible with PAMs without guanines [976].

Therefore, the conversion of other RNA-guided DNA nickases, such as SpRY, SpCas9- 117

NRRH, SpCas9-NRCH, and SpCas9-NRTH, into base editors that recognize different

PAM sequences will increase the number of codons targetable by iSTOP. However, it may still be restricted by an immediate G on the immediate 5’-side. Taken together, further engineering is required to enable virtually unrestricted iSTOP-mediated editing.

Similar to other CRISPR-related technologies, the efficacy of iSTOP could be affected by the occurrence of off-target mutations. Notably, initial work that examined the specificity of the cytidine deaminase activity of BE3 using a BE3 variant lacking the

UGI domain (BE3ΔUGI) concluded that BE3 causes fewer off-targets than Cas9 [977].

Additional studies on BE3-dependent off-targets led to the development of a high-fidelity

BE3 enzyme (HF-BE3) with reduced off-target base editing activity [978]. Therefore, more variants are being developed to reduce these off-targets such as Sniper-BE3 which shows 2.4-16.4 fold less off-targets than BE3 and eA3A-BE3 which restricted editing of cytosines to only those adjacent to thymines in the window [979, 980]. Unlike

BEs that have DNA off-targets, ABEs do not make detectable off-targets in the genome

[981-983].

Unfortunately, both BEs and ABEs generate RNA off-target editing and can self- edit their own transcripts leading to heterogeneity [931, 984, 985]. And while efforts are being made to engineer these base editors to prevent RNA-editing, it is imperative to minimize all off-targets to use base editors in clinical settings [931, 984, 985].

A2.2.4 Prime editing: A new precision gene editing tool

Recently the CRISPR toolbox has been expanded to include a breakthrough approach to editing called prime editing. Anzalone and colleagues developed a nickase

Cas9 fused to an engineered reverse transcriptase programmed with a sgRNA 118

containing the donor sequence called a prime editing guide RNA (pegRNA) [986]. The nCas9 is guided to the genomic sequence using the pegRNA generating a D-loop. This loop is then nicked by nCas9 producing a 3’-phosphate followed by primer-binding of the pegRNA to the PAM strand allows the reverse transcriptase to use the rest of the pegRNA as a template to copy the edited donor. This leads to the generation of either a

3’ flap that contains the edited sequence or a 5’ flap that lacks the editing. The 5’ flap is then cleaved by the structure specific endonuclease FEN1 or the 5’ exonuclease EXO1 and 3’ flap ligation drives the incorporation of the edited strand [987, 988]. Unlike base editing, prime editing does rely upon a specific base to be in a window of activity.

Furthermore, this technique enables the incorporation of insertions up to 44 base pairs or deletions up to 80 base pairs as well as nucleotide substitutions. Additionally, unlike base editors, so far prime editing has not been demonstrated to edit off-target RNA- molecules. Taken together, prime editing is a versatile genome editing method that can in principle correct up to 89% of the pathogenic human variations in ClinVar and increases the scope of genome editing that does not rely on the creation of a genotoxic

DSB [986].

A2.2.5 Perspectives

Genetic engineering and genome editing by the CRISPR-Cas system has been revolutionary as a research tool and opened doors for precision medicine and new gene therapies. Monogenic human diseases are disorders that are linked to mutations on a single gene. The World Health Organization currently estimates that there are over

10,000 monogenic human diseases and as of 2005 the “monogenic genome” consists of about 1,500 genes [989, 990]. DMD is a monogenic disease characterized by 119

progressive muscular degeneration that affects one in every 3500 live male births [991].

Approximately, 70% of all DMD-causing mutations are due to a single or multiexon deletion in the Dystrophin gene causing a frameshift resulting in a truncated protein

[991]. Therapies that can restore the frame of Dystrophin has the potential to restore protein function, survival, and improve muscle function [992]. Therefore, CRISPR-Cas- mediated editing of the loci, in particular prime editing, could prevent DMD.

Currently, spinal muscular atrophy (SMA), a monogenic neuromuscular disease, is being treated by a gene therapy from Novartis, called Zolgensma, in which a viral vector delivers a copy of the functional SMN gene to restore SMN protein function kick starting clinical applications of gene therapies [993, 994]. Because gene therapies will cure patients from their disease, these therapies come with a high price tag, for example Zolgensma has a $2.1 million or an annualized cost of $425,000 a year for 5 years. Gene therapies lead to a fast depletion of the total addressable patient population leading to a decline in demand over time, so companies that produce gene therapies need to price their treatment to take in consideration the loss of revenue overtime. Current pricing models take in consideration the life-time cost to the patient if their disorder was not prevented by the therapy making gene therapies the costliest in the market for a one-time treatment. But there can be a case in which a patient’s gene therapy is only partially effective (perhaps due to some irreversible damage accumulated in utero). If that is the case, there is a question of whether the patient should be reimbursed and, if so, how much. It is imperative for gene therapy pricing models to be determined in anticipation for CRISPR-mediate editing as a therapeutic.

Therefore, other pricing models have been proposed, annuity-based payment, 120

outcomes-based payment, outcomes-based rebate, and outcomes-based annuity, but these alternative models face challenges in the current healthcare system [995]. On top of the challenges facing pharmaceutical companies, there are several ethical considerations for using CRISPR methods.

While gene editing as a cure for genetic disorders is wildly favorable, CRISPR- mediated editing could in theory be used to make cosmetic changes or enhance natural ability [996]. Genome editing of the germline, in which these changes could be inherited for generations, is cautiously permitted, although by law United States federal funding cannot be used to conduct research involving human embryos [996]. Cautiously is the key word because current CRISPR technologies still contain a non-significant frequency of off-target editing, and these edits, if in the genome, can be inherited for generations significantly alterin g the future population. For non-clinical uses including changing human features, the territory becomes very murky. While editing an individual to reduce bad cholesterol could be beneficial so would enhancing cognitive factors or muscle mass. As noted above, gene therapies are currently cost prohibitive as a therapeutic.

Therefore, non-therapeutic editing could enable a system in which only the wealthiest individuals can be enhanced and perhaps increasing economic inequality in the United

States. Another risk is the return of eugenics by the editing out “undesirable” traits [996].

These ethical discussions, and others, need to be occurring at the world stage to ensure that no one misuses or weaponizes the potential of CRISPR. Already, in 2018 at the

Second International Summit on Human Genome Editing, a researcher announced the genomic editing of human embryos to prevent HIV infection [997]. Since then, the call for CRISPR ethics has been championed by Dr. Doudna, one of the key researchers 121

who developed CRISPR technologies for genome editing [998]. Hopefully, as the technology evolves research, ethical, and healthcare guidelines will continue to re- examine their current frameworks to maximize the great potential benefits of CRISPR- mediated editing while minimizing the possible risks of abuse.

122

A2.3 Figures

Figure A2. 1 Example of a search performed on the iSTOP website. Representative images of using the iSTOP website to search for SMARCAD1 sgSTOPs. (A) The iSTOP homepage with the advanced search button pressed and SMARCAD1 written in the search box. (B) The first page of results for the search in A.

123

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