Analysis of Human Y-Family DNA Polymerases and Primpol by Pre

Analysis of Human Y-Family DNA Polymerases and PrimPol

by Pre-Steady-State Kinetic Methods

Dissertation

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

E. John P. Tokarsky

Graduate Program in Biophysics

The Ohio State University

2018

Dissertation Committee

Dr. Zucai Suo, Advisor

Dr. Charles Bell

Dr. Jeff Kuret

Dr. Zhengrong Wu

Copyrighted by

E. John P. Tokarsky

2018

Abstract

Eukaryotic genomic DNA is efficiently and accurately replicated to ensure that an exact copy is created before cell division occurs. The complex machinery involved in

DNA replication is tightly coordinated and regulated to ensure it proceeds in a relatively uninhibited manner. The enzymes responsible for copying the genome are known as

DNA polymerases and these are responsible for catalyzing nucleotidyl transfer of the building blocks of DNA, deoxyribonucleotides (dNTPs), onto growing primer strands in the 5′-3′ direction. The active sites of DNA polymerases allow them to facilitate template-dependent nucleotidyl transfer based on Watson-Crick base pairing rules, i.e. adenine:thymine and cytosine:guanine (A:T and C:G). In humans, these enzymes must proceed at an extremely fast rate in order to replicate approximately 6 billion base pairs during each cell cycle. Reactive hydrocarbons, high energy UV-light, or free radicals generated during cellular processes (i.e. electron transport chain), modify DNA bases that can cause DNA polymerases to stall. Specialized DNA polymerases, from the Y-family, catalyze translesion DNA synthesis to replicate through modified DNA bases in order for the replication machinery to continue efficient DNA synthesis. Y-family DNA polymerases are able to accommodate bulky, modified bases into their active sites because they are flexible, and solvent-exposed. This characteristic makes them perfect candidates to bypass many types of DNA damage. However, these flexible active sites

ii make them error-prone and thus, Y-family DNA polymerases must be tightly regulated to ensure that high levels of DNA mutations that lead to genetic disease, are not introduced.

In this dissertation, I will describe my work with four human Y-family DNA polymerases, eta (hPolη), kappa (hPolκ), iota (hPolι), Rev1, and their abilities to bypass an air pollution-generated, bulky DNA lesion. 3-nitrobenzanthrone (3-NBA) is a byproduct of diesel fuel combustion that binds to particulate matter and is subsequently inhaled by humans. 3-NBA undergoes chemical modifications to become a reactive intermediate that subsequently modifies guanine bases producing N-(2′-deoxyguanosin-8- yl)-3-aminobenzanthrone (dGC8-N-ABA) lesions. We show that dGC8-N-ABA inhibits all four

Y-family DNA polymerases in some manner, but hPolη and hPolκ had the ability to bypass the lesion over time, whereas hPolι and Rev1 were unable to bypass it after many hours. An in-depth kinetic analysis was performed with hPolη, to determine the effect of the presence of the lesion on the kinetic parameters of dNTP binding and nucleotidyl transfer rate, at positions upstream, opposite, and downstream from the dGC8-N-ABA.

Directly opposite from the lesion, we found that hPolη had a 100-fold lower efficiency and an approximately 25% lower fidelity (i.e. ability to incorporate the correct nucleotide), with dATP being the highest misincorporation. This result is consistent with what has been found in other publications that show high levels of G→T transversion mutations occurring in human and mouse cells treated with 3-NBA.

A specialized primase-polymerase known as PrimPol, was discovered in humans in 2013. PrimPol exhibits similar properties to Y-family polymerases such as displaying relatively low efficiency and fidelity, and for having the ability to bypass certain types of

iii

DNA damage. However, based on in vitro experiments, the polymerase and primase activities of PrimPol are differentially regulated based on whether it utilizes manganese

(Mn2+) or magnesium (Mg2+) as a divalent metal ion cofactor for catalysis. We sought to determine the effect of divalent metal ions on the polymerase fidelity and sugar selectivity of PrimPol. We found that PrimPol was extremely error-prone (fidelity range

10-1 to 10-2) when utilizing Mn2+, but was ~100-fold more efficient, compared to Mg2+.

Finally, we showed that PrimPol could incorporate the nucleoside analogs and anticancer drugs, cytarabine and gemcitabine, as efficiently as normal dCTP in the presence of either Mn2+ or Mg2+.

Dedication

To my parents, Eugene and Linda, who have supported me through it all.

Acknowledgements

I have so many people to be grateful for during my journey as a graduate student.

I would first like to thank my undergraduate and graduate advisor, Dr. Zucai Suo. He was the first person to give me a chance to succeed in research and always pushed me to do my very best. I am thankful for his advice and constant support over the years. I am thankful to my committee members, Dr. Jeff Kuret, Dr. Charles Bell, and Dr. Justin Wu, for offering helpful advice from the time I began candidacy, to the end of my dissertation defense. It meant a great deal to me to have you all as guides during my graduate career.

My lab mates and friends, Dr. Walter Zahurancik, Dr. Varun Gadkari, Dr.

Anthony Stephenson, Dr. Austin Raper, and (soon to be Dr.) Andrew Reed, I could not thank you enough for making work fun. To be able to work with such smart individuals every day is something that I will miss, and will never take for granted. I know that every single one of you will be successful in your lives, and I hope to be there for all of you.

I want to thank my parents Eugene and Linda for their constant support throughout my entire life. I have had two wonderful parents that I always knew were in my corner no matter what challenges arose. I cannot thank you both enough for being there for me. My brother Joe, and sister Betsy have always been the older siblings that were there to keep me happy and laughing the entire way. I also wanted to show love to

vi my sister-in-law, Lauren, and brother-in-law, Elliot, and to my three beautiful nieces,

Guinevere, Samantha, and Phoebe. Lastly, I want to show my love and appreciation to my incredible fiancé, Kate Gilligan, who has seen me at my best and at my worst. I cannot wait to see what life has in store for us.

vii

Vita

2009-2013 B.S. Biology

The Ohio State University, Columbus, OH

2013-2018 Ph.D. Biophysics

The Ohio State University, Columbus, OH

2013-2018 Graduate Teaching Associate,

Department of Chemistry and Biochemistry

The Ohio State University, Columbus, OH

Publications

1. Tokarsky, E.J., Wallenmeyer, P.C., Phi, K.K., Suo, Z. (2017) Significant impact of

divalent metal ions on the fidelity, sugar selectivity, and drug incorporation efficiency

of human PrimPol. DNA Repair. 49, 51-59.

2. Tokarsky, E.J., Gadkari, V.V., Zahurancik, W.J., Malik, C.K., Basu, A.K., Suo, Z.

(2016) Pre-Steady-State Kinetic Investigation of Bypass of a Bulky Guanine Lesion

by Human Y-family DNA Polymerases. DNA Repair. 46, 20-28.

3. Patra, A., Politica, D.A., Chatterjee, A., Tokarsky, E.J., Suo, Z., Basu, A.K., Stone,

M.P., Egli, M. (2016) Mechanism of Error-Free Bypass of the Environmental

viii

Carcinogen N-(2- Deoxyguanosin-8-yl)-3-aminobenzanthrone Adduct by Human

DNA Polymerase η. ChemBioChem. 17 (21), 2033 –2037.

4. Vyas, R., Efthimiopoulous G., Tokarsky, E.J., Malik, C.K., Basu, A.K., Suo, Z.

(2015) Mechanistic Basis for the Bypass of a Bulky DNA Adduct Catalyzed by a Y-

Family DNA Polymerase. J Am Chem Soc. 137 (37), 12131-12142.

5. Vyas, R., Reed, A.J., Tokarsky, E.J., and Suo, Z. (2015) Viewing DNA Polymerase

β Faithfully and Unfaithfully Bypass an Oxidative Lesion by Time-Dependent

Crystallography. J Am Chem Soc. 137 (15), 5225-5230.

6. Gadkari, V.V., Tokarsky, E.J., Malik, C.K., Basu, A.K., and Suo, Z. (2014)

Mechanistic Investigation of the Bypass of a Bulky Aromatic DNA Adduct Catalyzed

by a Y-Family DNA Polymerase, DNA Repair. 21, 65-77.

Fields of Study

Major Field: Biophysics

Table of Contents

Abstract ...... ii Dedication ...... v Acknowledgements ...... vi Vita ...... viii Table of Contents ...... x List of Schemes ...... xiii List of Tables ...... xiv List of Figures ...... xv Chapter 1. Introduction ...... 1 1.1 Introduction to DNA replication ...... 1 1.1.1 Origin licensing and helicase assembly ...... 2 1.1.2 Action of essential polymerases and other enzymes at the eukaryotic replication fork ...... 3 1.2 DNA damage tolerance and repair ...... 6 1.2.1 Translesion DNA synthesis...... 7 1.2.2 Replication fork restart catalyzed by PrimPol ...... 10 1.2.3 Base excision repair of oxidative DNA damage ...... 11 1.3 Focus of this dissertation ...... 14 1.4 Tables ...... 16 1.5 Figures...... 17 Chapter 2. Pre-Steady-State Kinetic Investigation of Bypass of a Bulky Guanine Lesion by Human Y-family DNA Polymerases ...... 21 2.1 Introduction ...... 22 2.2 Materials and Methods ...... 24 2.2.1 Materials ...... 24 2.2.2 DNA substrates ...... 24 x

2.2.3 Reaction buffers and assay analysis ...... 25 2.2.4 Running start assays ...... 25 2.2.5 Electrophoretic mobility shift assays ...... 26 2.2.6 Standing start assay for hRev1 ...... 27 2.2.7 Single-turnover assays ...... 27 2.2.8 Biphasic assays ...... 28 2.3 Results ...... 28 2.3.1 Measuring relative polymerase efficiency during dGC8-N-ABA bypass ...... 28 2.3.2 DNA binding affinity estimated through electrophoretic mobility shift assays ...... 31 2.3.3 Determination of kinetic parameters for nucleotide incorporations upstream, opposite, and downstream from dGC8-N-ABA ...... 32 2.3.4 Determination of biphasic kinetic parameters at polymerase pause sites ...... 35 2.4 Discussion ...... 36 2.4.1 Kinetic basis for hPolη pausing caused by dGC8-N-ABA ...... 38 2.4.2 Kinetic mechanism for dGC8-N-ABA bypass and subsequent extension of the lesion bypass product ...... 39 2.4.3 Error-prone bypass of dGC8-N-ABA and subsequent extension of the lesion bypass product ...... 42 2.5 Schemes ...... 44 2.6 Tables ...... 46 2.7 Figures...... 53 Chapter 3: Significant impact of divalent metal ions on the fidelity, sugar selectivity, and drug incorporation efficiency of human PrimPol ...... 58 3.1 Introduction ...... 59 3.2 Materials and Methods ...... 60 3.2.1 Materials ...... 60 3.2.2 Expression and Purification of Human PrimPol ...... 61 3.2.3 Radiolabeling and annealing DNA substrates ...... 62 3.2.4 Fluorescence anisotropy titration ...... 63 3.2.5 Single-turnover kinetic assays ...... 63 3.3 Results ...... 64 3.3.1 Binding affinity of human PrimPol to DNA in the presence of Mn2+ or Mg2+ 64 3.3.2 Correct nucleotide incorporation efficiency in the presence of Mn2+ or Mg2+ 66 xi

3.3.3 Substitution fidelity of human PrimPol in the presence of Mn2+ or Mg2+ ...... 67 3.3.4 Sugar selectivity of human PrimPol in the presence of Mn2+ and Mg2+...... 69 3.3.5 Incorporation efficiencies of the triphosphates of four cytidine analog drugs gemcitabine, cytarabine, emtricitabine, and lamivudine by human PrimPol ...... 70 3.4. Discussion ...... 72 3.4.1 The polymerase activity of human PrimPol is error-prone in the presence of Mn2+ ...... 75 3.4.2 Human PrimPol displays moderate sugar selectivity in the presence of Mn2+ and Mg2+ ...... 76 3.4.3 Human PrimPol was able to incorporate the triphosphates of gemcitabine and cytarabine, but not emtricitabine and lamivudine ...... 77 3.4.4 Conclusion ...... 78 3.5 Tables ...... 80 3.6 Figures...... 86 Chapter 4. Conclusion ...... 92 4.1 Introduction ...... 94 4.2 Lesion bypass of dGC8-N-ABA and dG1,8 by Sulfolobus solfataricus Dpo4 ...... 94 4.3 Structural basis of dGC8-N-ABA bypass by hPolη ...... 99 4.4 Future directions for Y-family DNA polymerases and PrimPol ...... 100 4.5 Concluding Remarks ...... 103 4.6 Figures...... 104 References ...... 108

xii

List of Schemes

Scheme 2.1: Bioactivation of 3-NBA to form a bulky dGC8-N-ABA lesion...... 44 Scheme 2.2: Proposed kinetic mechanism for dGC8-N-ABA bypass and extension...... 45

xiii

List of Tables

Table 1.1:A list of DNA replicating enzymes found in humans, their families, and their major function(s) in human cells...... 16 Table 2.1: Sequences of DNA oligonucleotides ...... 46 Table 2.2: dGC8-N-ABA bypass halftimes of human Y-family polymerases at 37 °C ...... 47 Table 2.3: DNA binding affinity of hPolη at 23 °C ...... 48 Table 2.4: Kinetic parameters for dNTP incorporation into dGC8-N-ABA-containing DNA at 37 °C ...... 49 Table 2.5: Kinetic parameters for dNTP incorporation into undamaged DNA catalyzed by hPolη at 37 °C ...... 50 Table 2.6: Kinetic parameters from biphasic kinetic assays with hPolη at 37 °C ...... 51 Table 2.7: Kinetic parameters from biphasic kinetic assays with hPolη and undamaged DNA at 37 °C...... 52 Table 3.1: 21/41-mer DNA substrates ...... 80 Table 3.2: Pre-steady-state kinetic parameters for nucleotide incorporation onto 21/41- mer DNA substrates catalyzed by PrimPol in the presence of 5 mM Mn2+ at 37 C...... 81 Table 3.3: Pre-steady-state kinetic parameters for dNTP incorporation onto 21/41-mer DNA substrates catalyzed by PrimPol in the presence of 5 mM Mg2+ at 37 C...... 82

Table 3.4: Observed rate constants (kobs) for incorporation of a correct dNTP (1 mM) or a matched rNTP (1 mM) onto a DNA/DNA substrate (Table 3.1) in the presence of 5 mM MgCl2 catalyzed by human PrimPol at 37 C...... 83 Table 3.5: Pre-steady-state kinetic parameters for incorporation of dCTP analogs onto D- 6 (Table 3.1) catalyzed by human PrimPol at 37 C...... 84 Table 3.6: Comparison of the kinetic parameters for dGTP incorporation opposite dC by PrimPol and other human DNA polymerases in the presence of Mg2+...... 85

xiv

List of Figures

Figure 1.1: DNA polymerization reaction (two-metal ion mechanism) ...... 17 Figure 1.2: DNA polymerase structure...... 18 Figure 1.3: Watson Crick and Hoogsteen Base Pairing ...... 19 Figure 1.4: Repair of 8-oxoG via the BER pathway ...... 20 Figure 2.1: Running start assays at 37 °C ...... 53 Figure 2.2: Standing start assays for hRev1 at 37 °C ...... 54 Figure 2.3: Electrophoretic mobility shift assay for 21/26-mer dGC8-N-ABA binding by hPolη at 23 °C...... 55 Figure 2.4: Determination of the kinetic parameters for dCTP incorporation into 20/26- mer-dGC8-N-ABA at 37 °C...... 56 Figure 2.5: Biphasic kinetics of dNTP incorporation in the presence of a DNA trap at 37 °C...... 57 Figure 3.1: Determination of the binding affinity of PrimPol to DNA in the presence or absence of a divalent metal ion...... 86 Figure 3.2: Determination of kinetic parameters for dATP incorporation opposite dT in the presence of Mn2+ at 37 C...... 87 Figure 3.3: Gel image of time courses of dNTP misincorporation catalyzed by PrimPol in the presence of Mn2+ at 37 C...... 88 Figure 3.4: Gel image of time courses of dNTP misincorporation opposite dT catalyzed by PrimPol in the presence of Mg2+ at 37 C...... 90 Figure 3.5: Chemical structures of dC, rC, and their analogs used in this study...... 91 Figure 4.1: Bulky DNA lesions...... 104 Figure 4.2: Running Start Assays ...... 105 Figure 4.3: Space filling model of the hPolη active site with dGC8-N-ABA...... 106 Figure 4.4: NMR structure of the 12-mer DNA duplex containing a C8-dG-ABA adduct ...... 107

Chapter 1. Introduction

1.1 Introduction to DNA replication

The ability to perform DNA replication is a fundamental process that defines all living organisms. The successful copying of a genome, even in the most primitive of species, is a complex process that requires the coordinated actions of many protein complexes. Decades of research have led to the development of elegant models on the mechanisms that are involved in DNA replication, such as replication origin licensing,

DNA unwinding, and genome copying. With the ability to reconstitute large protein complexes and the rapid enhancement in cryo-electron microscopy (cryo-EM) technologies, it is possible to determine true mechanisms of important replication processes. Coupling protein complex reconstitution and cryo-EM with X-ray crystallography, single-molecule techniques, kinetics, and thermodynamic methods, the mechanistic details of the processes of DNA replication and repair are greatly elucidated.

In this chapter, I will give an overview of the individual steps of eukaryotic DNA replication and define the most important proteins that are involved. Later, I will discuss the processes that are involved in DNA repair that allow replication processes to function normally when DNA damage is present.

1.1.1 Origin licensing and helicase assembly

The process of DNA replication is carried out in a similar manner across all forms of life, and many of these protein components are homologous. In eukaryotes, the process known as origin licensing is the initial step of DNA replication where components necessary to unwind double stranded DNA must be recruited to specific locations in the genome. Origin licensing is initiated by the origin recognition complex

(ORC), which consists of six subunits (Orc1-6) and requires adenosine triphosphate

(ATP) as a cofactor for high affinity binding to DNA1; 2; 3. Once bound to DNA, ORC recruits Cdc6 to the origin4; 5, followed by Cdt16 and mini-chromosome maintenance complex (MCM2-7)7; 8; 9. MCM2-7 is the main component of the eukaryotic replicative helicase, and all six subunits are related members of the AAA+ superfamily of

ATPases10. MCM2-7 is loaded onto origin DNA via an ATP-dependent reaction where

Cdc6 and ORC facilitate the process of bringing together the Mcm2 and Mcm5 subunits, essentially closing the MCM2-7 ring and tethering it to the origin DNA11; 12; 13. Two

MCM2-7 rings are loaded at the same origin DNA in a head-to-head fashion, to facilitate unwinding bi-directionally. The two hexameric MCM2-7 rings mark potential sites of replication initiation and only become activated by cell-cycle dependent kinases. MCM2-

7 does not contain helicase activity on its own, however, it associates with Cdc4514; 15, and the hetero-tetrameric GINS16, to form the active Cdc45-MCM2-7-GINS (CMG) helicase. The presence of Cdc45 and GINS alter the molecular architecture of MCM2-7, enhance its DNA binding affinity, and stimulates its ATPase activity, thus allowing it to unwind partially duplexed DNA in a 3ʹ-5ʹ manner17; 18; 19. In order to form the active 2

CMG complex in vivo, the actions of two protein-dependent kinases, cyclin-dependent kinase (CDK) and Dbf4-dependent kinase (DDK) are upregulated at the onset of S- phase20; 21; 22. Specifically, phosphorylation of Mcm4 or Mcm6 subunits by DDK causes recruitment of Sld323 which subsequently binds to Cdc45. Next, CDK phosphorylates two proteins Sld2 and Sld3 which causes assembly of these two proteins with Dpb11,

GINS, and DNA polymerase epsilon (Polε)24; 25; 26; 27. Finally, one additional protein

MCM10 binding to the assembled CMG helicase, initiates DNA unwinding, though the specific molecular action of this protein is unclear at present28.

1.1.2 Action of essential polymerases and other enzymes at the eukaryotic replication fork

The process of genomic replication is carried out by enzymes known as DNA polymerases that catalyze the metal-ion dependent addition of deoxyribonucleotide triphosphates (dNTPs) onto nascent DNA strands in a template-dependent manner

(Figure 1.1). DNA polymerases have a right-handed architecture, and the polymerase subdomains are named accordingly and the Thumb, Palm, and Fingers subdomains are common to all DNA polymerases. The Thumb subdomain binds DNA, the Palm subdomain contains the catalytic residues that are necessary for catalysis, and the fingers subdomain aid bind to dNTPs (Figure 1.2). Other accessory domains are the exonuclease subdomain, which are common in replicative DNA polymerases and the Little Finger subdomain which is are typically found in Y-family DNA polymerases and may help in lesion bypass. Requiring only a small DNA primer, DNA polymerases catalyze DNA synthesis in the 5ʹ-3ʹ direction. Three DNA polymerases divide the work load of 3 eukaryotic replication: alpha (Polα), delta (Polδ), and Polε29. Polα was originally thought to be the only DNA polymerase required for DNA replication in eukaryotes, however, many more were subsequently discovered30; 31. Polα forms a hetero-tetrameric complex with human primase, which is responsible for generating short RNA primers. Eukaryotic primase contains two subunits, p49, the catalytic subunit and p58, an accessory subunit32.

Primase functions by binding to single stranded DNA, and two NTP molecules, then catalyzes dinucleotide formation and extends it to generate RNA primers of 8-10 nts33.

Subsequently, Polα exchanges positions with primase and extends the RNA primers by approximately 30 nucleotides (nts), creating a hybrid RNA/DNA primer. Polα exchanges on the primer-terminus with either Polε or Polδ, which are B-family polymerases that exhibit efficient and faithful DNA synthesis34. At physiological pH and temperature,

Polε and Polδ are capable of synthesizing dNTPs onto growing DNA strands at rates of

200- to greater than 500-nts per second, for Polδ35 and Polε36, respectively. Also, Polε and Polδ are highly faithful, i.e. select for correct dNTPs over incorrect dNTPs, in the range of 10-4 to 10-5 for Polδ and 10-4 to 10-7 for Polε. Furthermore, Polε and Polδ contain

3ʹ-5ʹ exonuclease activity which serves as a proof-reading mechanism to correct mismatches that may occur during DNA synthesis37. These characteristics make them ideal enzymes to copy the ~6 billion bases, in an approximate 24-hour cell cycle, for eukaryotes.

After CMG helicase unwinds double stranded DNA and a primer is generated, leading and lagging strand synthesis occur simultaneously but are catalyzed by different enzymes, with Polε acting at the leading strand and Polδ acting at the lagging strand38; 39.

Leading strand DNA synthesis occurs in the same direction of replication fork movement. Polε is generally accepted as the polymerase that performs this process based on genetic studies by Pursell38, and by reconstitution of the eukaryotic replisome by

Yurieva and O’Donnell40. Polε is proposed to form a complex with the eukaryotic CMG helicase to efficiently replicate DNA and also potentially enhance its processivity, i.e. catalyze many dNTP additions before dissociation from DNA41; 42; 43. By remaining coupled to the helicase, the action of Polε prevents long stretches of single-stranded DNA from being exposed to potentially reactive molecules in the cellular environment that modify DNA. In contrast, lagging strand DNA synthesis is considered discontinuous and must occur in the opposite direction of replication fork movement. Lagging strand DNA synthesis produces stretches of DNA (100-200 base pairs) known as Okazaki fragments44. After DNA is unwound by CMG, stretches of single stranded DNA on the lagging strand is bound by replication protein A (RPA) molecules. RPA is a single- stranded binding protein that has a DNA binding footprint of about 30 nts, and binds its substrate extremely tightly (~1 nM)45. RPA acts to protect single-stranded DNA from environmental factors that may cause the fragile single-stranded DNA to be modified or break46. Polδ displaces RPA molecules as it replicates until it meets the 5ʹ end of the preceding Okazaki fragment. The 5ʹ end contains a short stretch of RNA (generated by primase), that must be removed before ligation can occur. This process is facilitated by the actions of both Polδ, which actively displaces the short stretch of RNA (replacing it with DNA) and flap endonuclease 1 which catalyzes the removal of 1-2 ribonucleotide monophosphates molecules at a time until all RNA molecules are removed47. Once RNA

5 is removed from the 5ʹ end of the Okazaki fragment, the two strands of DNA are joined together by the action of DNA ligase I48.

1.2 DNA damage tolerance and repair

In order to maintain the integrity of the genome, the cell utilizes multiple pathways during and after replication that either allow the replication fork to proceed in an efficient manner, or to directly remove the damage from the DNA strand. Translesion

DNA synthesis (TLS)49 and replication fork restart50 are two pathways that are utilized by the cell to bypass DNA damage, allowing replication to continue in a timely manner.

These DNA damage tolerance pathways are catalyzed by one or more polymerases for

TLS and a single primase-polymerase (PrimPol), for replication fork restart. Base excision repair (BER)51, mismatch repair and nucleotide excision repair are pathways that identify, remove, and replaced damaged bases with undamaged bases before the cell divides. Another form of DNA damage that can have devastating consequences if unrepaired, are double-stranded breaks46. Double stranded breaks are generally caused by UV-light, and the cell utilizes pathways known as non-homologous end joining, and homologous recombination to repair this type of damage. This section will focus primarily on the DNA damage tolerance and repair processes of TLS, replication fork restart, and BER repair of the most common oxidative lesion, 7,8-dihydro-8-oxo-2′- deoxyguanosine (8-oxoG).

1.2.1 Translesion DNA synthesis

Replicative polymerases have tight and selective active sites that cannot tolerate modified bases, and therefore stall during incorporation of dNTPs opposite damaged bases52. Polymerase stalling can lead to replication fork uncoupling, where the helicase continues unwinding DNA causing replication fork collapse and cell death53. To prevent these outcomes, the cell utilizes specialized DNA polymerases that are capable of bypassing DNA damage and allowing replicative polymerases to continue efficient and faithful DNA replication downstream from DNA damage. The specialized DNA polymerases are commonly from the Y-family, and are able to bypass many types of

DNA damage. In eukaryotes, four Y-family polymerase(s) eta (Polη), kappa (Polκ), iota

(Polι), and Rev1, and a B-family polymerase zeta (Polζ) are responsible for catalyzing the majority of TLS49. Two other polymerases that may be involved in TLS, polymerases theta (Polθ) and nu (Polν) are newly discovered and will not be discussed in this review.

A list of human DNA polymerases and their functions are located in Table 1.1.

The variant form of xeroderma pigmentosum (XPV) is caused by mutations in the

POLH gene (encoding for Polη), that leads to sensitivity to UV light causing painful blisters and early-onset cancer in XPV patients54. XPV cells are incapable of replicating through cis-syn TT-dimers, and likely the loss of Polη activity leads to the observed phenotype. Polη efficiently and faithfully incorporates A’s opposite the two T’s in a TT- dimer55, whereas no other polymerase is capable of this activity in an error-free manner.

Polη is also responsible for bypassing, O6-methylguanine (m6G)56, 8-oxoG57, and other bulky lesions such as dGAP58, and dGC8-N-ABA (discussed below)59.

Polκ is a Y-family DNA polymerase from the DinB subfamily and is related to

E.coli DinB (Pol IV), and the archaeal enzymes, Dpo4 and Dbh. Polκ, is often unable to incorporate dNTPs directly opposite from a lesion (i.e. bypass step). Since the bypass step can often be error-prone, Polκ has the unique ability to efficiently extend from mismatched primer-termini to allow TLS to proceed60; 61. However, this process can lead to buried mismatches that can become mutations if the mismatch repair pathway fails.

Polκ generates frame-shift mutations by a process called misinsertion misalignment where the templating base of a mispair is repositioned (looped out), and the primer- terminal nucleotide base pairs with the next complementary templating base49. Polκ may play a role during extension from m6G and cys-syn TT-dimers albeit in an error-prone manner61. Additionally, Polκ can catalyze the bypass and extension of other DNA

62 63 64 adducts such as -HOPdG , 8-oxoG , cisplatin intrastrand cross-links , and N2-CH2(9- anthracenyl)G65. Polκ may also play a role in nucleotide excision repair to repair UV damage66.

Polι shows interesting sequence dependence during incorporation of dNTPs by displaying higher fidelity and efficiency with a template dA, rather than dC, dG, or dT.

For example, incorporation of dTTP is 1,000- to 100,000-fold more efficient opposite a templating dA67 compared to incorrect dNTPs. However, for template dC, dG, or dT, correct incorporation by Polι may only be 10- to 100-fold more efficient than incorrect dNTPs, displaying its error-prone nature68. Based on crystallographic analysis of Polι, it has been shown that its molecular architecture facilities the generation of Hoogsteen base pairs, rather than canonical Watson-Crick base pairs69 (Figure 1.3). During dTTP

8 incorporation opposite dA, the purine base adopts a syn conformation to avoid steric clash with residues Leu62 and Gly59 in the fingers domain. The incoming dTTP forms two base pairs on its Watson-Crick edge with dA on its Hoogsteen edge. The mechanism by which Polι forms Hoogsteen base pairs may aid in its ability to bypass certain types of

DNA damage such as minor groove lesions such as -HOPdG70.

Similar to Polι, Rev1 does not utilize Watson-Crick base pairing to synthesize

DNA but utilizes a distinct mechanism from any other DNA polymerase. Rather than facilitating interactions between the templating base and incoming dNTP. Rev1 utilizes

Arg357 to hydrogen bond with dCTP, and the extrahelical template base dG is accommodated in a hydrophobic pocket while L358 rests in the conventional location of a templating base to form preferable contacts with dCTP71. Thus, Rev1 has been classified as a deoxycitidyl transferase as it preferably incorporates dCTP, regardless of the identity of the templating base. Interestingly, Rev1 has lower polymerase fidelity than other Y-family polymerases, and is typically unable to bypass DNA lesions on its own72. However, Rev1 but may act as a protein scaffold for other DNA polymerases, such as Polη, Polι, and Polζ to stably bind primer-termini opposite DNA lesions73.

Specifically, Polη, Polι, and Polζ have been shown to associate with Rev1 during co- immunoprecipitation assays by forming interactions with a region in the C-terminus of

Rev174; 75. Therefore, Rev1 may not be directly involved during the bypass or extension of DNA lesions, but may be involved in coordinating other polymerases to catalyze TLS.

Polζ, a B-family polymerase that lacks proofreading activity, is similar to Polκ in its ability to catalyze the extension step of TLS. Interestingly, Polζ is unable to

9 incorporate dNTPs opposite from DNA lesions but is extremely efficient during the extension step of TLS. This likely means that Polζ will overtake a primer-terminus after another Y-family polymerase catalyzes the bypass step76. As was discussed above, Rev1 may be crucial for the coordination of the two polymerases to perform the process of TLS efficiently. Polζ is also a highly efficient and catalyzes error-free polymerase activity during extension. It is responsible for catalyzing TLS of TT-dimers, (6-4) photoproducts,

8-oxoG, m6G, and thymine glycol77; 78. Unfortunately, the mechanistic basis for DNA synthesis is currently unknown, and the only structural details of Polζ solved with cryo-

EM at 23 Å that describe only its molecular architecture79.

1.2.2 Replication fork restart catalyzed by PrimPol

The human gene CCDC111, originally identified from in silico analysis of the superfamily of archaeo-eukaryotic primases (AEPs)80, encodes for a bifunctional enzyme that catalyzes both primase and polymerase activities and is named PrimPol. Human

PrimPol is classified as member of the nucleocytoplasmic large DNA virus (NCLDV) clade of AEPs, and is found in both nuclear and mitochondrial cellular compartments80;

81. PrimPol has two distinct domains, an AEP domain that is responsible for catalyzing 3′-

5′ nucleotidyl transfer, and a UL52-like zinc finger domain that binds single stranded

DNA (ssDNA) and is necessary for its primase activity82; 83; 84; 85. Importantly, the loss of

PrimPol in human cells (PrimPol-/-) leads to slowed replication fork rates due to the formation of stalled replication forks86, and severely diminished mitochondrial DNA replication82. PrimPol has been implicated in DNA damage tolerance pathways based on

10 its recruitment to DNA by replication protein A (RPA)83; 87 and its association to chromatin during UV-induced cellular stress85; 86 A unique aspect of PrimPol that is relatively understudied is the role it plays in replication fork restart based on its ability to reprime downstream of DNA damage85; 86, G-Quadruplex DNA structures88, and chain terminated replication forks89. Unlike most DNA primases, PrimPol selectively utilizes deoxyribonucleotide 5′-triphosphates (dNTPs), rather than ribonucleotide 5′- triphosphates (rNTPs) for its primase activity. This preferential utilization of dNTPs has been proposed to be advantageous during replication fork restart because little or no RNA will be left, and further processing of the primers would not be needed3. Interestingly, replication fork restart catalyzed by PrimPol is an important process in mitochondria that provides a mechanism to cope with genotoxic stress, and may enable origin-independent initiation of lagging-strand DNA synthesis79.

1.2.3 Base excision repair of oxidative DNA damage

Although TLS and replication fork restart are necessary to maintain replication fork progression, these process leave behind damaged DNA bases that will persist during subsequent rounds of replication. In order to truly repair the damaged bases, they must be removed from the DNA strand completely. DNA damage occurs at a rate of approximately 10,000 lesions per cell per day in humans90 and more than 100 different types of damaged bases91 resulting from oxidative stress caused by normal cellular processes have been identified. BER is the primary pathway involved in processing oxidative DNA damage and must be tightly coordinated and regulated to prevent

11 cytotoxic DNA intermediates, such as gapped and nicked DNA, from causing harm to the cell. Notably, inactivation or mutation of various BER proteins can lead to cancer formation. Below, I discuss in detail the BER pathway involved in the repair of the most common, and well-studied oxidative lesion, 8-oxoG.

Reactive oxygen species are free radicals generated as byproducts of normal cellular metabolism that react with and alter DNA bases, generating lesions such as 8- oxoG. 8-oxoG is potentially mutagenic due to its dual-coding potential that arises from its ability to stably form both Watson-Crick and Hoogsteen base pairs (Figure 1.3).

Human DNA polymerase beta (hPolβ), the BER enzyme responsible for gap-filling DNA synthesis, inserts either a correct dCTP or an incorrect dATP opposite 8-oxoG, forming

Watson-Crick (dC:8-oxoG) or Hoogsteen (dA:8-oxoG) base pairs, respectively92; 93.

Formation of dA:8-oxoG base pairs, if unrepaired, can lead to G→T transversion mutations which are found in many forms of cancer13. The process of BER typically involves four enzymes that are responsible for catalyzing the individual steps in the BER pathway (Figure 1.4). Importantly, coordination of BER enzymes is essential to protect unstable DNA intermediates from the cellular environment, and failure to do so properly can lead to base substitutions, genome instability and cell death94.

First, DNA glycosylases rapidly search the DNA strand, identify a damaged base, and catalyze the removal of the base by cleaving the N-glycosidic bond. Various DNA glycosylases exist to combat the many forms of DNA lesions that arise daily51; 91. Two important DNA glycosylases responsible for recognizing 8-oxoG are human 8-oxoG glycosylase (hOGG1) and MutYH95. hOGG1 recognizes a dC:8-oxoG base-pair and

12 removes the 8-oxoG, whereas MutYH recognizes an 8-oxoG:dA mismatched base-pair and removes the adenine base opposite from 8-oxoG51. The action of the glycosylase leaves behind an apurinic/apyrimidinic (AP) site, characterized by a deoxyribose sugar lacking a base at the C1ʹ position. AP-sites that are not repaired properly can lead to base substitutions or insertion/deletion mutations during subsequent rounds of DNA replication. AP endonuclease 1 (APE1) binds to the AP-site and catalyzes strand scission, nicking the DNA backbone. APE1 begins by ‘flipping out’ the AP-site into its active site and stabilizes it with four loop structures and an α-helix, thus forming a

APE1•DNA complex96; 97. APE1 then catalyzes cleavage of the DNA backbone by utilizing two aspartate residues that coordinate a single Mg2+, thus activating a nucleophilic water molecule to attack the 5ʹ-phosphorous atom of the AP site deoxyribose, leaving a single-nucleotide gap with a 3ʹ-OH and a 5ʹ-deoxyribosephosphate

(5ʹ-dRP)98. Next, the DNA substrate is bound by hPolβ, which catalyzes dRP lyase and gap-filling DNA synthesis activities. The dRP-lyase activity of hPolβ removes the 5ʹ- dRP99 and is a necessary step to allow ligation of the DNA strand. For gap-filling DNA synthesis, hPolβ binds the DNA substrate in an ‘open conformation’ and transitions to a

‘closed conformation’ upon dNTP binding, thus positioning catalytic groups for efficient nucleotide incorporation. The reaction is facilitated by three catalytic metal ions (Mg2+), that coordinate active site residues and stabilize the buildup of negative charge associated with pyrophosphate formation92; 93. Once the gapped DNA is filled, the resulting nicked

DNA substrate is sealed by DNA ligase III-α (Lig3α) and X-ray repair cross-

13 complimenting protein 1 (XRCC1) complex (Lig3α•XRCC1) which joins the 5ʹ- phosphate and 3ʹ-OH groups thus completing the BER pathway100.

1.3 Focus of this dissertation

This dissertation focuses on aspects of the activities of human Y-family polymerases and PrimPol. Chapter 2 focuses on the ability of human Y-family polymerases to bypass a bulky DNA lesion, dGC8-N-ABA. dGC8-N-ABA is the major product that forms that forms when a small polycyclic aromatic hydrocarbon, 3-

Nitrobenzanthrone (3-NBA), is inhaled and subsequently metabolized in the cell101; 102. 3-

NBA is formed by the incomplete combustion of diesel fuels and is found in high concentrations in industrial areas. 3-NBA is a known mutagen and potential carcinogen, and high levels of 3-NBA were found in the collected urine samples of occupational workers. In human cells, 3-NBA undergoes nitroreduction followed by either N,O- acetyltransferase or sulfotransferase, to form intermediates that react with guanine bases, thus forming dGC8-N-ABA (Scheme 2.1). Thus, understanding the mechanistic basis of mutation formation caused by 3-NBA could ultimately lead to the development of therapeutics that could cure genetic diseases. We sought to determine the capabilities of each human Y-family DNA polymerases to catalyze TLS of dGC8-N-ABA, in vitro.

Furthermore, we determined that Polη was one of two polymerases to bypass dGC8-N-ABA and we performed an in-depth kinetic characterization for nucleotide insertion upstream, opposite, and downstream from the lesion. Lastly, we determined a plausible kinetic mechanism for Polη to catalyze TLS of dGC8-N-ABA.

PrimPol, as discussed above is an enzyme that is implicated in the DNA damage tolerance pathway known as replication fork restart. PrimPol is similar to Y-family DNA polymerases based on its relatively low efficiency and low fidelity while also exhibiting

DNA lesion bypass capabilities. However, during its initial characterization it was observed that PrimPol is much more efficient during DNA synthesis when utilizing Mn2+ as a metal ion cofactor, compared to Mg2+. Mn2+ is a known mutagen and this may be based on its ability to adopt different coordination geometries103. This may ultimately allow PrimPol, and other DNA polymerases, to facilitate nucleotidyl transfer without having the proper contacts for Watson-Crick base pairing. We used pre-steady-state kinetics to study the fidelity of dNTP incorporation catalyzed by PrimPol in the presence of either Mn2+ or Mg2+ as the catalytic metal ion cofactor. Furthermore, we determined the ability of PrimPol to discriminate from rNTPs and other nucleoside analogs, such as anticancer drugs (gemcitabine and cytarabine) and HIV reverse transcriptase inhibitors

(emtricitabine and lamivudine).

1.4 Tables

Human DNA Polymerase Family Major Function (s) Pol α (alpha) B Replicative, primer formation Pol δ (delta) B Replication of lagging strand Pol ε (epsilon) B Replication of leading strand Pol γ (gamma) A Mitochondrial DNA replication Pol θ (theta) A DNA replication timing, translesion DNA synthesis

Pol ν (nu) A Translesion DNA synthesis Pol β (beta) X Base excision repair Pol λ (lambda) X Base excision repair Pol µ (mu) X Non-homologous end joining Pol η (eta) Y Translesion DNA Synthesis Pol κ (kappa) Y Translesion DNA Synthesis Pol ι (iota) Y Translesion DNA Synthesis Rev1 Y Translesion DNA Synthesis Pol ζ (zeta) B Translesion DNA Synthesis TdT X V(D)J Recombination PrimPol AEP Replication Fork Restart

Table 1.1:A list of DNA replicating enzymes found in humans, their families, and their major function(s) in human cells.

1.5 Figures

Figure 1.1: DNA polymerization reaction (two-metal ion mechanism). D705 and

D882 are from E. coli DNA polymerase I. Two metal ions are present in the active site that stabilize the resulting pentacoordinated transition state of the reaction. Metal ion A is responsible for activating the 3′-OH of the primer for attack on the α-phosphate of the incoming dNTP. Metal ion B stabilizes the negative charge that builds up on the leaving oxygen and also chelates the β- and γ-phosphates. Figure is adapted from Brautigam and

Steitz, 2002104.

Figure 1.2: DNA polymerase structure. The structure presented is of RB69

Polymerase, A B-family DNA polymerase (PDB code: 1IG9). The common DNA polymerase subdomains are named above. The Thumb subdomain binds to DNA, the

Palm subdomain contains catalytic residues, and the Fingers subdomain binds to dNTP.

Figure 1.3: Watson Crick and Hoogsteen Base Pairing. (A) Watson-Crick base pairing for A-T and G-C base pairs. (B) Hoogsteen base pair formation between A-T, and

G-C. Figure adapted from Johnson, 2005105.

Figure 1.4: Repair of 8-oxoG via the BER pathway. HoGG1 removes the damaged 8- oxoG via glycosylase activity. APE1 catalyzes strand scission, and hPolβ inserts dGTP opposite the templating dC. LigIII•XRCC1 ligates the DNA strand, completing BER.

Chapter 2. Pre-Steady-State Kinetic Investigation of Bypass of a Bulky Guanine Lesion by Human Y-family DNA Polymerases

Reproduced in part with permission from Tokarsky, E.J., Gadkari, V.V.,

Zahurancik, W.J., Malik, C.K., Basu, A.K., Suo, Z. (2016) Pre-Steady-State Kinetic

Investigation of Bypass of a Bulky Guanine Lesion by Human Y-family DNA

Polymerases. DNA Repair. 46, 20-28. The full article is available at https://doi.org/10.1016/j.dnarep.2016.08.002. E. John Tokarsky planned and performed the kinetic experiments with assistance from Varun V. Gadkari. E.J.T., Walter J.

Zahurancik and Zucai Suo analyzed the results. Modified DNA substrates were provided by Chachal K. Malik and Ashis K. Basu. E.J.T. wrote the initial draft of the manuscript.

Z.S. conceived the research and modified the manuscript. This work was supported by the National Institutes of Health grant ES009127 to both A.K.B. and Z.S.

2.1 Introduction

DNA damage caused by prevalent chemical agents in the environment poses significant threats to cell survival and proliferation and represents a major concern for occupational and public health. One molecule in particular, 3-nitrobenzanthrone (3-

NBA), a nitro-polycyclic aromatic hydrocarbon, was discovered in diesel exhaust as bound to particulate air matter 101. 3-NBA is ubiquitously present in the environment, especially in areas exposed to high levels of diesel emissions, such as cities or areas of heavy construction 106. It has one of the most mutagenic profiles of known pollutants tested in the Ames Salmonella typhimurium (TA98) assay and is responsible for inducing mostly G→T transversions 101; 107. Upon inhalation, 3-NBA is metabolized into various mutagenic and potentially carcinogenic metabolites that can react with purines in DNA

108. These reactions lead to the formation of bulky, aromatic DNA lesions which act as roadblocks to genomic replication. One such product, N-(2′-deoxyguanosin-8-yl)-3- aminobenzanthrone (dGC8-N-ABA), is of particular interest because it is a major lesion formed in mice after intraperitoneal treatment with 3-NBA 102. The bulky lesion dGC8-N-

ABA, produced by the electrophilic attack of deoxyguanosine (dG) by a 3-NBA metabolite

108 (Scheme 2.1), has been found to severely block replication fork progression 109.

During normal DNA replication, highly efficient and faithful DNA polymerases perform the bulk of the work. It is known that replicative DNA polymerases possess tight and selective active sites that allow them to stringently choose against incorrect

22 nucleotides but also limit their abilities to catalyze translesion synthesis (TLS) through damaged template nucleotides 38; 110. When a damaged site is encountered, replicative polymerases are stalled due to a significantly slowed nucleotide incorporation activity and a competing 3′→5′ exonuclease activity, resulting in futile cycles of DNA synthesis and excision111. In contrast, Y-family DNA polymerases, which lack proofreading activity and possess comparatively flexible and solvent accessible active sites, are more suited to accommodate DNA lesions and catalyze TLS49. Notably, the human genome encodes four Y-family members: DNA polymerase eta (hPolη), kappa (hPolκ), iota

(hPolι), and hRev1. These human polymerases vary in their lesion bypass capabilities and are differentially recruited to damaged DNA to catalyze TLS depending on the lesion that is present 49. In this study, we sought to determine which of the human Y-family DNA polymerase(s) are able to efficiently bypass dGC8-N-ABA in vitro. Previously we utilized pre-steady-state kinetic methods to study the bypass of dGC8-N-ABA catalyzed by a model

Y-family polymerase, Sulfolobus solfataricus DNA Polymerase IV (Dpo4)112. We observed that Dpo4 is significantly less efficient and faithful during dGC8-N-ABA bypass and the subsequent extension compared to undamaged DNA, implicating the mutagenic effect of this lesion in vivo. Here we applied similar methods to determine the most active human Y-family polymerase for dGC8-N-ABA bypass and establish a kinetic mechanism for

TLS of dGC8-N-ABA. Initially, running start assays were performed to study the effect that a site-specifically placed, bulky dGC8-N-ABA lesion had on each human Y-family polymerase. Our results indicate that hPolη is a likely candidate to catalyze TLS of dGC8-

N-ABA in vivo. Subsequently, we performed single-turnover kinetic assays to determine the

DNA binding affinities (Kd, DNA), maximum nucleotide incorporation rate constants (kp), apparent nucleotide binding affinities (Kd, dNTP), and the substrate specificities (kp/Kd, dNTP) for nucleotide incorporations by hPolη at sites upstream, opposite, and downstream from the lesion. Finally, biphasic kinetic assays were performed to establish a minimal kinetic mechanism for nucleotide incorporation opposite dGC8-N-ABA and the subsequent extension of the lesion bypass product.

2.2 Materials and Methods

2.2.1 Materials.

All reagents were purchased from the following companies: OptiKinase from

United States Biochemical, [γ-32P] ATP from Perkin-Elmer, Biospin-6 columns from

Bio-Rad Laboratories, and dNTPs from Bioline. Full-length hPolη and truncated fragments of hPolκ (9-518), hPolι (1-420), and hRev1 (341–829) were overexpressed and purified as described previously113.

2.2.2 DNA substrates.

A DNA template (26-mer) containing a dGC8-N-ABA lesion at the 21st position was synthesized as described previously 112. DNA containing the dGC8-N-ABA lesion was kept in dark conditions to preserve the light-sensitive substrate. All other synthetic oligonucleotides were purchased from Integrated DNA Technologies (IDT) and gel purified by 17% denaturing polyacrylamide gel electrophoresis (PAGE). Primers were

5′-radiolabeled by incubating with [γ-32P]-ATP and OptiKinase for 3 hr, followed by heat

24 inactivation (95 °C for 2 min), and isolation of radiolabeled DNA using a Biospin-6 column to remove excess, unreacted [γ-32P]-ATP. A primer was annealed to a template at a 1:1.35 ratio. Annealing solutions containing undamaged DNA and dGC8-N-ABA- containing DNA were heated to 95 and 72 °C, respectively, for 5 minutes, followed by slow cooling to room temperature. All DNA substrates used in this study are listed in

Table 2.1.

2.2.3 Reaction buffers and assay analysis.

All chemical quench assays were performed with a KinTek Rapid Chemical

Quench-Flow apparatus at 37 °C in buffer R (50 mM HEPES, pH 7.5 at 37 °C, 50 mM

NaCl, 5 mM MgCl2, 0.1 mM EDTA, 10% glycerol, 5 mM DTT, and 0.1 mg/mL Bovine

Serum Albumin (BSA)). Electrophoretic mobility shift assays (EMSAs) were performed at 23 °C in buffer S (50 mM Tris-HCl, pH 7.5 at 23 °C, 50 mM NaCl, 5 mM MgCl2, 0.1 mM EDTA, 10% glycerol, 5 mM DTT, and 0.1 mg/mL BSA). All concentrations listed were final upon mixing and all gels were scanned using a TyphoonTrio (GE Healthcare).

Quantification was carried out using ImageQuant (GE Healthcare) and Kaleidagraph

(Synergy) software.

2.2.4 Running start assays.

A pre-incubated solution of hPolη (100 nM), hPolκ (100 nM), hPolι (1 μM), or hRev1 (1 µM) and 5′-[32P]-17/26-mer (100 nM) was rapidly mixed with all four dNTPs

(200 µM each). After various incubation periods, the samples were quenched with the

25 addition of 0.37 M EDTA and separated by 17% PAGE (8 M Urea, 1X TBE). Analysis and quantification of gels were performed as described in 2.2.3 Reaction buffers and assay analysis. The relative lesion bypass efficiencies (bypass %) were calculated for each polymerase as a function of reaction time. For each time point t, the bypass % was calculated as a ratio of bypass to encounter events (Eq. 2.1) as described previously 58; 113;

114.

dGC8-N-ABA bypass % = (B/E) x 100 = [B/([20-mer] + B)] x 100 Eq. 2.1

Quantification of running start assays (Figure 2.1A-H) was achieved by treating the total encounter (E) events as equivalent to the concentration of 20-mer, and bypass (B) events as equivalent to the sum of the concentrations of all products with sizes greater than or equal to the 21-mer. The bypass % was calculated using Eq. 2.1 then plotted against reaction times, t. To compare the relative amount of time it took for each polymerase to

C8-N-ABA bypass bypass dG , the t50 was determined as the time to bypass of 50% of the total lesions encountered. In a similar fashion, the t50 was determined as the time to bypass

50% of the undamaged dG base.

2.2.5 Electrophoretic mobility shift assays.

hPolη (2 – 650 nM) was titrated into solutions containing 5′-[32P]-labeled DNA

(10 nM) and allowed to equilibrate in buffer S for 15 min at room temperature (23 °C).

The samples were separated using 4.5% native PAGE which were analyzed and quantified as described in 2.2.3 Reaction Buffers and Assay Analysis. The concentration

26 of hPolη•DNA complex was plotted against the total concentration of hPolη and the data were fit to a quadratic equation (Eq. 2.2):

2 1/2 [E•DNA] = 0.5(Kd, DNA + E₀ + D₀) – 0.5[(Kd, DNA + E₀ + D₀) -4E₀D₀] Eq. 2.2

Where E₀ is the initial enzyme concentration, D₀ is the initial DNA concentration, and

Kd, DNA is the apparent equilibrium dissociation constant for the hPolη•DNA complex.

2.2.6 Standing start assay for hRev1.

A pre-incubated solution of hRev1 (200 nM) and 5′-[32P]-labeled undamaged

(20/26-mer) or damaged (20/26-mer-dGC8-N-ABA) DNA (200 nM) in buffer R was mixed with dATP, dCTP, dGTP, dTTP, or all four nucleotides (200 µM each). After 10 min, the reaction was quenched with the addition of 0.37 M EDTA and the samples were separated using 17% denaturing PAGE.

2.2.7 Single-turnover assays.

A pre-incubated solution of 5′-[32P]-labeled DNA (20 nM) and hPolη (130 nM) in buffer R was rapidly mixed with varying concentrations of dNTP (10 - 1200 µM). After various incubation times, the reaction was quenched with the addition of 0.37 M EDTA.

The samples were then analyzed using 17% denaturing PAGE. Product concentration was plotted against time and the data were fit to a single-exponential equation (Eq. 2.3).

[Product] = A[1 - exp (- kobst)] Eq. 2.3

Where kobs is the observed rate constant of nucleotide incorporation and A is the reaction amplitude. The kobs values obtained using Eq. 2.3 were plotted against the respective dNTP concentration and the data were fit to a hyperbolic equation (Eq. 2.4).

kobs = kp[dNTP]/([dNTP] + Kd, dNTP) Eq. 2.4

Where kp is the maximum nucleotide incorporation rate constant and Kd, dNTP is the apparent equilibrium dissociation constant for the binding of dNTP to hPol•DNA.

2.2.8 Biphasic assays.

Biphasic assays were performed by rapidly mixing a pre-incubated solution of hPolη (120 nM) and 5′-[32P]-labeled DNA (30 nM) with various concentrations of dNTP

(25-1,000 µM) and excess, unlabeled DNA trap (5 µM). After various incubation periods, reactions were quenched with the addition of 0.37 M EDTA and samples were separated using 17% denaturing PAGE. Product concentration was plotted against time and the data were fit to a double-exponential equation (Eq. 2.5)

[Product] = Af[1-exp (-kft)] + As[1-exp(-kst)] Eq. 2.5

Where Af and As are the reaction amplitudes of the fast and slow phases, respectively, and kf and ks are the rate constants of the fast and slow phases, respectively.

2.3 Results

2.3.1 Measuring relative polymerase efficiency during dGC8-N-ABA bypass.

To test if each individual human DNA polymerase could bypass a single dGC8-N-

ABA lesion, running start assays were performed by rapidly mixing pre-incubated hPolη,

28 hPolκ, hPolι, or hRev1 and undamaged 17/26-mer or damaged 17/26-mer-dGC8-N-ABA containing a site-specifically placed dGC8-N-ABA (Table 2.1) with all four dNTPs. The lesion stalled DNA synthesis by each of the Y-family polymerases, but the degree of stalling as well as the positions at which stalling was observed varied between each polymerase. For hPolη, moderate pausing was observed at the 19-mer (-2), 20-mer (-1),

21-mer (0), and 22-mer (+1) positions. Significant accumulation of the 20-mer product indicated that incorporation of nucleotide at the bypass step (Position -1) was the most problematic for hPolη (Figure 2.1B). Despite this stalling, the full-length product appeared after 6 s, which was only 2-fold longer than the time required for hPolη to synthesize the full-length product with the undamaged 17/26-mer. hPolκ stalled significantly during the bypass and extension steps, indicated by the 20-mer and 21-mer accumulation at these positions (Figure 2.1D). hPolκ produced the full length products within 1 s with the undamaged 17/26-mer and took only 3-fold longer (3 s) with the

17/26-mer-dGC8-N-ABA substrate, indicating that hPolκ was similarly efficient as hPolη when bypassing dGC8-N-ABA. The observed accumulation of the 25-mer product is likely caused by polymerase slippage and template repositioning within the 5′-dC-rich region of the template (Table 2.1), resulting in the “-1” frameshift 115; 116. Additionally, the full length product (25-mer) produced by this mechanism was likely further extended via a blunt-end addition 117; 118. Interestingly, the bulky lesion caused significant stalling of hPolι and hRev1 with no appearance of the full-length product even after long reaction periods (3 and 10.5 hr, respectively). With 17/26-mer-dGC8-N-ABA, hPolι synthesized the bypass and extension products after 300 and 3,900 s, respectively, which are 30- and 65-

29 fold, respectively, longer than with the undamaged 17/26-mer (Figure 2.1E-F). In comparison, hRev1 synthesized the lesion bypass product within 1 hr but was unable to extend it even after 10.5 hr. The failure of hRev1, a dCTP polymerase, to efficiently catalyze TLS of dGC8-N-ABA was likely caused by multiple template-independent misincorporations of dCTP by hRev1 72, resulting in DNA distortion. This is supported by the fact that hRev1was not able to synthesize the full-length product even with the undamaged DNA substrate (Figure 2.1H). The results of the running start assay with hRev1 prompted us to perform standing start primer extension assays to qualitatively evaluate the efficiency and fidelity of hRev1 to incorporate a nucleotide directly opposite from dGC8-N-ABA. A pre-incubated solution of hRev1 and 20/26-mer-dGC8-N-ABA (or

20/26-mer) was mixed and reacted with a solution containing either dATP, dCTP, dGTP, dTTP, or all four dNTPs. Interestingly, hRev1 showed greater selectivity with damaged than with undamaged DNA by predominantly incorporating dCTP (Figure 2.2). With undamaged DNA, hRev1 incorporated all four nucleotides with a preference of dCTP > dATP = dGTP = dTTP.

To provide a comparison for the amount of time it took for each human Y-family polymerase to bypass dGC8-N-ABA, gels from the running start assays (Figure 2.1A-H)

bypass were quantified and Eq. 2.1 was used to calculate values for t50 and t50 which represent the times it took to elongate 50% of the 20-mer to the 21-mer or longer

bypass products with undamaged and damaged DNA, respectively. The t50 and t50 times measured for each of the Y-family polymerases are summarized in Table 2.2. Notably, hPolι and hRev1 did not bypass 50% of the dGC8-N-ABA lesion in the allowed reaction time

bypass for the running start assays. In contrast, the relatively short t50 times for hPolη (13 s) and hPolκ (7 s) suggest a potential role involving these two polymerases in the bypass and/or extension of dGC8-N-ABA lesions in vivo. Consistently, a recent publication reports

~25 and ~30% efficiency reductions in TLS of dGC8-N-ABA in human embryonic kidney

(HEK293T) cells when hPolη and hPolκ, respectively, were subject to gene knockdown

119. Based on the slightly smaller perturbation of DNA synthesis by hPolη in the presence of dGC8-N-ABA, we decided to use pre-steady-state kinetic methods to initially characterize the efficiency and fidelity of TLS of dGC8-N-ABA catalyzed by hPolη. However, it is apparent based on both the running start and HEK293T cell-based assays that hPolκ is also potentially involved in TLS of dGC8-N-ABA in vivo. Accordingly, we are currently performing similar kinetic studies to determine the effect of dGC8-N-ABA on nucleotide incorporation kinetics with hPolκ.

2.3.2 DNA binding affinity estimated through electrophoretic mobility shift assays.

To determine the apparent equilibrium dissociation constant for the binding of

C8-N-ABA hPolη to DNA containing dG (Kd, DNA), EMSAs were performed. A representative gel image and a plot for an EMSA experiment are shown in Figure 2.3 for hPolη binding

C8-N-ABA to 21/26-mer-dG . For all DNA substrates tested, the Kd, DNA values were within a

2-fold range for the corresponding undamaged and damaged DNA substrates (Table 2.3).

Interestingly, the binding affinities of hPolη for 19/26-mer-dGC8-N-ABA and 20/26-mer-

C8-N-ABA dG (Kd, DNA =12 ± 2 nM for both) were 2-fold tighter than the corresponding undamaged DNA substrates (Kd, DNA =25 ± 5 nM; 23 ± 2 nM). Based on our recently

31 published ternary crystal structure of hPolη in complex with dGC8-N-ABA-DNA and dCTP

120, the slight increase in binding affinity likely arises from positioning of the ABA moiety of dGC8-N-ABA within a hydrophobic cleft in the hPolη active site thereby stabilizing the conformation of the damaged base. Similar effects are observed in the crystal structures of Dpo4 with dGBPDE and dGAP 121; 122. Overall, hPolη binds to dGC8-N-

ABA-containing DNA tightly enough such that it does not significantly contribute to the polymerase pausing pattern observed in the running start assay (Figure 2.1B). Similar results have been obtained for dGAP bypass by hPolη 58 as well as dGAP, dGC8-N-ABA, and abasic site bypass by Dpo4 112; 123; 124.

2.3.3 Determination of kinetic parameters for nucleotide incorporations upstream, opposite, and downstream from dGC8-N-ABA.

Since DNA binding did not significantly affect TLS of dGC8-N-ABA by hPolη, it is possible that the stalling of hPolη during TLS (Figure 2.1B) was caused by decreased nucleotide binding affinity and incorporation efficiency. To examine this hypothesis, we utilized single-turnover assays to measure the apparent equilibrium dissociation constant for nucleotide binding (Kd, dNTP) and the maximum rate constant of nucleotide

C8-N-ABA incorporation (kp) at positions upstream, downstream, and opposite from dG . By varying the primer length, we were able to measure the kinetic parameters at specific positions along the template strand relative to dGC8-N-ABA, and thus determine the effect of dGC8-N-ABA on the efficiency and fidelity of nucleotide incorporation during each step of

TLS catalyzed by hPolη. Briefly, a pre-incubated solution of 5′-[32P]-labeled DNA (20

32 nM, Table 2.1) and hPolη (130 nM) was rapidly mixed with a single dNTP at varying concentrations. Reactions were analyzed as described (2.2.3 Reaction Buffers and Assay

Analysis), and the data were fit to Eq. 2.3 to yield an observed rate constant of incorporation (kobs). The kobs values were then plotted against the respective dNTP

concentrations and fit to a hyperbolic equation (Eq. 2.4) to yield the kp and Kd, dNTP

(Figure 2.4). hPolη showed a decrease in nucleotide incorporation efficiency (kp/Kd, dNTP) at positions upstream, opposite, and downstream from the lesion (Position -3, -2, -1,

0, +1). With 18/26-mer-dGC8-N-ABA at Position -3, hPolη correctly incorporated dCTP

-1 with a kp of 66 ± 1 s , although the dCTP binding affinity was weakened by 4-fold (Kd, dCTP =200 ± 11 µM) relative to undamaged 18/26-mer. As a result, the polymerase

-1 -1 -1 efficiency (kp/Kd, dCTP = 3.3 x 10 s µM ) was only slightly reduced compared to the

-1 -1 value with 18/26-mer (kp/Kd, dCTP = 1.5 s µM ). Similar experiments were carried out with the 19-mer (Position -2), 20-mer (Position -1), 21-mer (Position 0), and 22-mer

(Position +1) primers and the kinetic parameters are listed in Tables 2.4 and 2.5, for dGC8-N-ABA-containing and undamaged templates, respectively. The most significant stalling occurred during incorporation opposite dGC8-N-ABA (Position -1), caused by a 25-

-1 fold decrease in kp (1.9 ± 0.1 s ) and a 4-fold increase in Kd, dCTP (325 ± 44 µM) which resulted in an overall reduction in efficiency by approximately 100-fold (kp/Kd, dCTP = 5.8

-3 -1 -1 C8-N-ABA x 10 s µM ) with 20/26-mer-dG compared to undamaged 20/26-mer (kp/Kd,

-1 -1 -1 dCTP = 5.6 x 10 s µM ). At the extension step (Position 0), hPolη displayed a kp that was 49-fold slower (0.8 ± 0.03 s-1) but surprisingly bound the incoming dGTP more

C8-N-ABA tightly (Kd, dGTP = 28 ± 5 µM). With 21/26-mer-dG , hPolη showed an approximate

-2 -1 -1 5-fold increase in efficiency (kp/Kd, dGTP = 2.8 x 10 s µM ) compared to the lesion bypass step (Position -1). With 22/26-mer-dGC8-N-ABA (Position +1), the efficiency of

-2 -1 -1 hPolη was similar (kp/Kd, dCTP = 4.2 x 10 s µM ) to that measured at position -2 (Table

2.4) which is indicated by similarly moderate accumulation at these positions (Figure

2.1B).

Previously, it has been shown that dGC8-N-ABA bypass carried out by hPolη is error- prone in HEK293T cells119. Thus, we investigated the effect of dGC8-N-ABA on the fidelity of hPolη during dGC8-N-ABA bypass and the subsequent extension step. The fidelity of

hPolη, defined as (kp/Kd, dNTP)incorrect/[(kp/Kd, dNTP)correct + (kp/Kd, dNTP)incorrect], was calculated for the bypass (20/26-mer-dGC8-N-ABA) and extension (21/26-mer-dGC8-N-ABA) steps by using the single-turnover kinetic data of both correct and three incorrect nucleotide incorporations (Table 2.4). At these two steps, the fidelity of hPolη (Table

2.4) was determined to be in the ranges of 4.8 x 10-2 to 1.6 x 10-1 and 4.9 x 10-2 to 1.9 x

10-1, respectively. In comparison, the fidelity of hPolη with undamaged DNA was previously determined to be in the range of 10-2 to 10-3 (Table 2.5) by pre-steady-state kinetics 58 and this range is consistent with the values determined using phage plaque assays 125; 126. Thus, the presence of dGC8-N-ABA significantly reduced the fidelity of hPolη at both the lesion bypass and subsequent extension steps, especially during misincorporations of dGTP (29-fold) and dATP (33-fold) opposite dGC8-N-ABA (Table

2.4). For these two TLS steps, nucleotide incorporation probabilities, given by [(kp/Kd, dNTP)damaged DNA/Σ(kp/Kd, dNTP)damaged DNA] x 100, were also calculated. The probability of correct incorporation was reduced from 98% and 96% with undamaged DNA (Table 2.5)

34 to 77% and 74% with damaged DNA (Table 2.4) for dGC8-N-ABA bypass and the subsequent extension step, respectively. Notably, with damaged DNA, the probability of dATP misincorporation was 15 and 17% at the bypass and extension steps, respectively, and these probabilities are the highest among all misincorporations (Table 2.4).

Consistently, the G→T transversion was found to occur at a high rate during TLS of dGC8-N-ABA in vivo 102; 119; 127.

2.3.4 Determination of biphasic kinetic parameters at polymerase pause sites.

Nucleotide incorporation during TLS has often been shown to follow biphasic kinetics. Thus, we performed biphasic kinetic assays as described previously58; 123; 124; 128 to test if hPolη followed the same trend during dGC8-N-ABA bypass and the subsequent extension step. A pre-incubated solution of 5′-[32P]-labeled DNA (30 nM) and hPolη (120 nM) was rapidly mixed with increasing concentrations of a correct nucleotide (25 to

1,000 µM) and an unlabeled trap DNA substrate 20/26-mer (5 µM). The trap DNA was added to a 150-fold molar excess over the labeled DNA so that any enzyme that had dissociated from the radiolabeled DNA substrate during the incubation time would be sequestered by the trap DNA. Thus, only labeled DNA product formed during a single enzyme binding event would be observed. The trap DNA concentration (5 µM) was determined to be sufficiently high enough to sequester any hPolη that had dissociated from 5′-[32P]-labeled DNA123. During the incorporation of dCTP (1,000 µM) onto 20/26- mer-dGC8-N-ABA, hPolη indeed followed biphasic kinetics (Figure 2.5A). The plot of reaction time versus product concentration was fit to Eq. 2.5 (2.2.8 Biphasic Assays) to

35 yield reaction amplitudes of Af = 5.8 ± 0.8 nM (19%) and As = 8.6 ± 0.8 nM (28%) with

-1 -1 rate constants of kf = 7.2 ± 1.4 s and ks = 0.42 ± 0.06 s for the fast and slow phases, respectively (Table 2.6). To test if the kinetic parameters were affected by dCTP concentration, similar biphasic kinetic experiments were performed individually with 300 and 600 µM dCTP. Indeed, as the dCTP concentration increased, the reaction rate constants (kf and ks) also increased while the reaction amplitudes remained relatively unchanged (Table 2.6). In addition, the biphasic kinetic experiments were also performed with 21/26-mer-dGC8-N-ABA. As with 20/26-mer-dGC8-N-ABA, a similar biphasic kinetic trend and a dNTP concentration-dependent increase in the rate constants were observed with 21/26-mer-dGC8-N-ABA during the extension step (Figure 2.5B and Table 2.6).

Interestingly, the dNTP concentration-dependent increase in the rate constants (kf and ks) had been observed previously with Dpo4 when it bypassed the dGC8-N-ABA 112 and dG1,8 lesions 129. However, our current study is the first one to report this concentration- dependence for lesion bypass catalyzed by a human Y-family DNA polymerase. Notably, kinetic parameters for the DNA trap experiments performed with undamaged 20/26-mer and 21/26-mer DNA are reported in Table 2.7, showing that even with undamaged DNA substrates, hPolη also displays biphasic kinetics of nucleotide incorporation.

2.4 Discussion

In this chapter, we performed running start assays to determine which of the four human Y-family DNA polymerases were able to most efficiently catalyze TLS of dGC8-N-

ABA in vitro. Both hPolη and hPolκ were able to bypass dGC8-N-ABA, while hPolι and

36 hRev1 were completely stalled by the lesion (Figure 2.1). Consistently, small interfering

RNA (siRNA)-mediated knockdown of hPolι had a minimal effect on overall TLS of dGC8-N-ABA in HEK293T cells, suggesting that hPolι plays a very minor role during dGC8-

N-ABA bypass in vivo 119. Surprisingly, siRNA-mediated knockdown of hRev1 proved to be detrimental to dGC8-N-ABA bypass in HEK293T cells. Based on its inability to catalyze the bypass of dGC8-N-ABA in vitro, hRev1 likely serves as a scaffold protein that synergistically interacts with proliferating cell nuclear antigen and other TLS polymerases to aid in TLS of dGC8-N-ABA and likely other DNA lesions in vivo 58; 130; 131;

132.

In contrast, hPolη and hPolκ were able to bypass dGC8-N-ABA with similarly small

bypass C8-N-ABA t50 values (Table 2.2), indicating potential roles for these polymerases in dG bypass in vivo. Consistently, knockdown of these two polymerases in HEK293T cells significantly reduced the efficiency of TLS of dGC8-N-ABA 119. Knockdown of hPolκ resulted in no change in the percentage of mutagenic dGC8-N-ABA bypass events, suggesting that hPolκ carries out relatively error-free TLS of dGC8-N-ABA 119. Conversely, knockdown of hPolη resulted in a 39% decrease in mutation frequency, indicating that the bypass of dGC8-N-ABA by hPolη in vivo is error-prone 119. Taken together, the bypass of dGC8-N-ABA by hPolη is likely a mechanism through which 3-NBA promotes lesion- induced mutagenesis and tumorigenesis. To determine a kinetic basis for mutagenic bypass of dGC8-N-ABA by hPolη, we utilized pre-steady-state kinetic methods to investigate the effect of a site-specifically placed dGC8-N-ABA on hPolη-catalyzed nucleotide incorporation at and near the lesion.

2.4.1 Kinetic basis for hPolη pausing caused by dGC8-N-ABA.

Overall, the binding affinity of hPolη to DNA (Table 2.3) is insignificantly affected by the presence of dGC8-N-ABA and therefore does not significantly contribute to the polymerase stalling observed in Figure 1B. Instead, the decrease of nucleotide incorporation rate constants and the weakened nucleotide binding affinities are likely responsible for the observed accumulation of intermediate products. A similar kinetic trend is seen for the conversion of 18-mer→19-mer→20-mer as well as 19-mer→20- mer→21-mer. Based on the kinetic data in Table 2.4, the first conversion step has a higher rate constant (kp) and also a higher efficiency ratio (kp/Kd, dNTP) than the second, less efficient conversion step, leading to the observed accumulation of the 19-mer and 20- mer products (Figure 1B). The 19-mer and 20-mer accumulation was absent at later time points (30-1200 s) due to their eventual conversion to the 20-mer and 21-mer products, respectively. For the reaction series of the conversion of 20-mer→21-mer→22-mer and

21-mer→22-mer→23-mer, the first conversion step was less efficient than the second conversion step. This kinetic pattern led to accumulation of 20-mer and 21-mer as seen in Figure 1B. Taken together, it is clear that the conversion of 20-mer→21-mer is less efficient than both the preceding and subsequent reactions leading to the strong accumulation of the 20-mer product. For 23-mer, 24-mer, and 25-mer, their accumulation appeared the same in Figure 1B, indicating that the nucleotide incorporation in the consecutive conversion steps associated with these intermediate products were similarly efficient, and that hPolη likely resumed its normal polymerase efficiency after being

38 several positions downstream from the lesion. Thus, the conversion rate and efficiency of

23-mer→24-mer, although not measured here, are likely as high as the conversion of 18-

-1 -1 -1 mer→19-mer (kp = 66 s , kp/Kd, dNTP = 0.33 µM s ) but higher than the conversion of 22-

-1 -1 -1 mer→23-mer (kp = 9.1 s , kp/Kd, dNTP = 0.042 µM s ), contributing to the observed 22- mer accumulation (Figure 1B).

2.4.2 Kinetic mechanism for dGC8-N-ABA bypass and subsequent extension of the lesion bypass product.

Our biphasic kinetic assays revealed that two distinct phases exist when hPolη incorporated correct dNTPs opposite dGC8-N-ABA (Position -1) and one base pair downstream (Position 0) from the lesion. The fast phases were characterized by relatively small reaction amplitudes and fast rate constants while the slow phases had larger reaction amplitudes and slower rate constants (Table 2.5). Individual contributions of the amplitudes (Af and As) and rate constants (kf and ks) were used to calculate the overall nucleotide incorporation rate constant in the presence of the DNA trap. For example, in the presence of 20/26-mer-dGC8-N-ABA and 1,000 µM dGTP (Table

2.5), the sum of the individual contributions of the fast phase [(7.2 s-1) x 19%] and slow phase [(0.42 s-1) x 29%] gave an overall rate constant of 1.5 s-1. As expected, 1.5 s-1 is

-1 close to the kobs of 1.4 s , estimated using Eq. 2.4, 1.0 mM dCTP, and corresponding

C8-N-ABA measured kp and Kd, dCTP values (Table 2.4), for dCTP incorporation opposite dG under single-turnover reaction conditions. Similarly, in the presence of 21/26-mer-dGC8-

N-ABA and 50 µM dGTP, the sum of the individual contributions of the fast phase [(5.2 s-1)

39 x 15%] and slow phase [(0.13 s-1) x 30%] gave an overall nucleotide incorporation rate

-1 -1 constant of 0.8 s , which is similar to the kobs of 0.5 s , calculated using Eq. 2.4, 50 µM dGTP, and corresponding measured kp and Kd, dGTP values (Table 2.4), for dGTP incorporation during the extension step. Thus, the biphasic kinetic feature of nucleotide incorporation at Positions -1 and 0 under single-turnover conditions was successfully deconvoluted into a fast and a slow phase through the DNA trap experiments (Figure

2.5). These two kinetic phases prompted us to propose a kinetic mechanism for dGC8-N-

ABA bypass and the subsequent extension by including the formation of productive

P N (E•DNAn ) and non-productive (E•DNAn ) complexes (Scheme 2.1A). The portion of

P the hPolη population bound in the E•DNAn will bind correct dNTP and rapidly elongate

DNA with a fast rate constant of kf. The fast phase kf should be dNTP concentration- dependent and can reach its maximum value of k1. The remainder of hPolη bound in the

N N E•DNAn complex will elongate DNA only after the slow conversion (ke) of E•DNAn

P → E•DNAn . The slow phase rate constant of ks is a function of both ke and k1 but should

AP be limited by ke. The bypass of a bulky dG lesion by hPolη as well as the bypass of an abasic site, a cisplatin-d(GG) adduct, and a dGAP lesion by Dpo4 have previously been proposed to follow this mechanism 58; 123; 124; 128. If this mechanism is correct, the

N P E•DNAn → E•DNAn conversion rate (ke) and reaction amplitudes should not depend on dNTP concentration. However, both kf and ks increased with dNTP concentration at

Positions -1 and 0 while both the fast and slow phase reaction amplitudes remained unchanged. To accommodate these observations, an alternative kinetic mechanism

(Scheme 2.2B) was proposed for hPolη to bypass dGC8-N-ABA and to subsequently extend

40 the lesion bypass product. The same mechanism has been proposed by us for dGC8-N-ABA

112 bypass by Dpo4 . This mechanism shows that ks is actually a function of an exchange rate (ke) and a slow nucleotide incorporation rate constant (k2). If ks had remained the same as dNTP concentration was increased, this would indicate that ks were solely a function of ke, and the mechanism could be simplified to reflect Scheme 2.2A. Because ks increased with higher dNTP concentration, especially during the extension step, ks was

C8-N-ABA predominantly a function of k2 during TLS of dG by hPolη. Notably, the total reaction amplitudes for the bypass and extension steps (48% and 46%, Table 2.6) are significantly smaller than the corresponding values with undamaged DNA (88% and

86%, Table 2.7). These differences indicate that hPolη may have bound to damaged

D DNA in a catalytically incompetent complex (E•DNAn ) where conformational change(s) of hPolη would not be able to produce a catalytically competent complex (Scheme 2.2B).

In instances when any molecules of the E•DNAD complex dissociated, the enzyme would bind to excess, unlabeled trap DNA resulting in no detectable product formation. Thus, the presence of such a dead-end binary complex can only be inferred by the sub-maximal product formation observed over the course of TLS of dGC8-N-ABA by hPolη. Similar results have been observed in previous kinetic studies of lesion bypass under comparable conditions 58; 123; 124; 128.

Although the hPolη•dGC8-N-ABA-DNA binary structures are not available yet, some of their binding conformations could be inferred from an NMR structure of a 12-mer

C8-N-ABA 133 DNA duplex containing an embedded dG :dC basepair and crystal structures of yeast Polη (yPolη) in complex with DNA containing a bulky 2-acetylaminofluorine

41 lesion (dGAAF) 134. In the NMR structure, the ABA moiety of dGC8-N-ABA is intercalated in the DNA duplex and takes the place of a base pair, with the damaged guanine in the syn conformation and extruded into the major groove, and with the opposing cytosine also pushed into the major groove 133. Similarly, one yPolη•dGAAF-DNA structure shows that the bulky adduct stacks on the primer/template junction base pair, occupies the polymerase active site, and precludes the incoming nucleotide from binding 134. If such a

C8-N-ABA D binding conformation also exists in hPolη•dG -DNA, it likely represents E•DNAn in Scheme 2.2. Interestingly, in the other structure of yPolη•dGAAF-DNA 134, the AAF adduct is partially rotated out of the DNA helix to allow some excess for incoming dCTP

N and the structure likely represents the binding conformation of E•DNAn (Scheme 2.2).

Lastly, a recently solved ternary crystal structure of hPolη•dGC8-N-ABA-DNA•dCTP shows that the adducted guanine forms a standard Watson-Crick pair with the incoming dCTP, with the ABA moiety accommodated inside a hydrophobic cleft to the side of the active site of hPolη120. Such a binding conformation of hPolη•dGC8-N-ABA-DNA is considered to

P be E•DNAn (Scheme 2.2).

2.4.3 Error-prone bypass of dGC8-N-ABA and subsequent extension of the lesion bypass product.

During dGC8-N-ABA bypass, hPolη showed a 10-fold reduced fidelity (4.8 x 10-2 –

1.6 x 10-1) relative to undamaged DNA (2.0 x 10-3–1.2 x 10-2, Table S1) with nucleotide incorporation preference following the order of dCTP > dATP > dGTP > dTTP (Table

2.4). Thus, the presence of dGC8-N-ABA decreased the ability of hPolη to discriminate

42 against incorrect nucleotides, especially dATP. Our kinetic studies show that hPolη will misincorporate dATP 15% of the time when bypassing dGC8-N-ABA (Table 2.4).

Interestingly, our previous studies have shown that in NER-proficient HEK293T cells,

G→T transversions were the most common mutation resulting from dGC8-N-ABA bypass, followed by G→A transitions and G→C transversions, albeit at much lower rates 119.

Furthermore, our previous studies have also demonstrated that siRNA-mediated knockdown of hPolη leads to a 39% decrease in mutation frequency in HEK293T cells, and strongly suggest that hPolη plays a major role in the error-prone bypass of dGC8-N-ABA in vivo 119. Therefore, these observations from published cell-based assays 119 are consistent with the results of our kinetic studies with hPolη (Table 2.4).

2.5 Schemes

Scheme 2.1: Bioactivation of 3-NBA to form a bulky dGC8-N-ABA lesion. 3-NBA is nitroreduced to form an intermediate (N-OH-ABA) which is further converted to N-AcO-

C8-N- ABA or N-OSO3H-ABA species that react with deoxyguanosine bases to form dG

ABA.

Scheme 2.2: Proposed kinetic mechanism for dGC8-N-ABA bypass and extension.

(A) Kinetic mechanism for nucleotide incorporation from a productive conformation. (B)

Kinetic mechanism for nucleotide incorporation from a productive or non-productive conformation. Abbreviations: E, hPolη; DNAn, annealed primer-template DNA (Table

2.1); DNAn+1, DNAn extended by 1 nucleotide; dNTP, correct deoxyribonucleotide;

D P E•DNAn , hPolη bound to DNA in a dead-end complex; E•DNAn ; hPolη bound to

N DNA in a productive conformation; E•DNAn ; hPolη bound to DNA in a non-productive complex; PPi, pyrophosphate; k1 and k2, nucleotide incorporation rate constants; ke, rate constant of exchange between non-productive and productive complexes of hPolη bound to DNA.

2.6 Tables

Primers (Position)a Sequence

17-mer (-4) 5′-AACGACGGCCAGTGAAT-3′

18-mer (-3) 5′-AACGACGGCCAGTGAATT-3′

19-mer (-2) 5′-AACGACGGCCAGTGAATTC-3′

20-mer (-1) 5′-AACGACGGCCAGTGAATTCG-3′

21-mer (0) 5′-AACGACGGCCAGTGAATTCGC-3′

22-mer (+1) 5′-AACGACGGCCAGTGAATTCGCG-3′

Templates

26-mer 3′-TTGCTGCCGGTCACTTAAGCGCGCCC-5′

26-mer-dGC8-N-ABA 3′-TTGCTGCCGGTCACTTAAGCGCGCCC-5′

G indicates the dGC8-N-ABA lesion at the 21st position in the template. aPosition of primer 3′ terminus relative to the templating dGC8-N-ABA.

Table 2.1: Sequences of DNA oligonucleotides

a bypass b bypass Polymerase t50 (s) t50 (s) t50 /t50

hPolη 0.7 13 19

hPolκ 0.3 7 23

hPolι 200 >10,800 >55

hRev1 1,800 >37,000 >20 aCalculated as the time required for traversing 50% of undamaged dG. bCalculated as the time required for bypassing of 50% of dGC8-N-ABA.

Table 2.2: dGC8-N-ABA bypass halftimes of human Y-family polymerases at 37 °C

a b DNA Substrate Kd, undamaged DNA (nM) Kd, damaged DNA (nM) Affinity Ratio

18/26-mer 28 ± 3 32 ± 5 1.1

19/26-mer 25 ± 5 12 ± 2 0.5

20/26-mer 23 ± 2 12 ± 2 0.5

21/26-mer 26 ± 1 26 ± 7 1.0

22/26-mer 17 ± 6 24 ± 4 1.4 aDamaged DNA refers to template 26-mer containing a dGC8-N-ABA lesion at the

21st position. b Calculated as Kd, damaged DNA/Kd, undamaged DNA.

Table 2.3: DNA binding affinity of hPolη at 23 °C

K k /K Efficiency Fidelity Probability e dNTP d, dNTP k (s-1) p d, dNTP Fidelityc (µM) p (µM-1s-1) Ratioa,b ratiod (%)

18/26-mer-dGC8-N-ABA - Template dG (Position -3)

dCTP 200 ± 11 66 ± 1 3.3 x 10-1 5 - - - dATP 369 ± 48 2.1 ± 0.1 5.7 x 10-3 0.6 1.7 x 10-2 7.4 -

19/26-mer-dGC8-N-ABA - Template dC (Position -2)

dGTP 150 ± 34 6.6 ± 0.4 4.4 x 10-2 23 - - - dCTP 90 ± 19 0.093 ± 0.0001 1.0 x 10-3 5 2.2 x 10-2 4.8 -

20/26-mer-dGC8-N-ABA - Template dGC8-N-ABA (Position -1) dCTP 325 ± 44 1.9 ± 0.1 5.8 x 10-3 97 - - 77 dATP 161 ± 38 0.18 ± 0.01 1.1 x 10-3 3 1.6 x 10-1 33 15 dGTP 39 ± 5 0.014 ± 0.0003 3.6 x 10-4 3 5.8 x 10-2 29 5 dTTP 475 ± 85 0.14 ± 0.01 2.9 x 10-4 23 4.8 x 10-2 4 4

21/26-mer-dGC8-N-ABA - Template dC (Position 0) dGTP 28 ± 5 0.8 ± 0.03 2.9 x 10-2 29 - - 74 dATP 473 ± 25 3.2 ± 0.10 6.8 x 10-3 2 1.9 x 10-1 11 17 dCTP 201 ± 20 0.4 ± 0.02 2.0 x 10-3 9 6.4 x 10-2 3 5 dTTP 392 ± 108 0.6 ± 0.01 1.5 x 10-3 3 4.9 x 10-2 9 4

22/26-mer-dGC8-N-ABA - Template dG (Position +1) dCTP 215 ± 67 9.1 ± 1.0 4.2 x 10-2 4 - - - dGTP 41 ± 8 0.028 ± 0.001 6.8 x 10-4 0.6 1.6 x 10-2 6.5 - a Calculated as [(kp/Kd, dNTP)undamaged DNA/(kp/Kd, dNTP)damaged DNA]. bKinetic parameters for undamaged DNA from Table S1 of Sherrer et al.58 c Calculated as (kp/Kd, dNTP)incorrect/[(kp/Kd, dNTP)incorrect) + (kp/Kd, dNTP)correct]. d Calculated as Fidelitydamaged DNA/Fidelityundamaged DNA. e Calculated as [(kp/Kd, dNTP)damaged DNA/Σ(kp/Kd, dNTP)damaged DNA] x 100.

Table 2.4: Kinetic parameters for dNTP incorporation into dGC8-N-ABA-containing

DNA at 37 °C

b kp/Kd, dNTP Probability dNTP K (µM) k (s-1) Fidelitya d, dNTP p (µM-1s-1) (%) 18/26-mer - Template dG (Position -3) dCTP 54 ± 14 83 ± 6 1.54 - - dATP 374 ± 63 1.3 ± 0.1 3.5 x 10-3 2.3 x 10-3 - 19/26-mer - Template dC (Position -2) dGTP 59 ± 6 60 ± 2 1.01 - - dCTP 103 ± 37 0.49 ± 0.01 4.7 x 10-3 4.6 x 10-3 - 20/26-mer - Template dG c (Position -1) dCTP 85 ± 11 48 ± 2 5.6 x 10-1 - 98 dATP 350 ± 36 0.95 ± 0.04 2.7 x 10-3 4.8 x 10-3 <1 dGTP 37 ± 8 0.04 ± 0.002 1.0 x 10-3 2.0 x 10-3 <1 dTTP 494 ± 70 3.3 ± 0.2 6.6 x 10-3 1.2 x 10-2 1 21/26-mer - Template dC c (Position 0) dGTP 46 ± 8 39 ± 2 8.4 x 10-1 - 96 dATP 242 ± 25 3.4 ± 0.1 1.4 x 10-2 1.7 x 10-2 2 dCTP 119 ± 21 2.2 ± 0.1 1.8 x 10-2 2.1 x 10-2 2 dTTP 712 ± 73 3.3 ± 0.2 4.6 x 10-3 5.4 x 10-3 1 22/26-mer - Template dG (Position +1) dCTP 261 ± 88 43 ± 4 1.7 x 10-1 - - dGTP 47 ± 9 0.02 ± 0.001 4.3 x 10-4 2.5 x 10-3 - a Calculated as (kp/Kd, dNTP)incorrect/[(kp/Kd, dNTP)incorrect) + (kp/Kd, dNTP)correct]. b Calculated as [(kp/Kd, dNTP)undamaged DNA/Σ(kp/Kd, dNTP)undamaged DNA] x 100. c Kinetic parameters for undamaged DNA from Table S1 of Sherrer et al. 58

Table 2.5: Kinetic parameters for dNTP incorporation into undamaged DNA catalyzed by hPolη at 37 °C

-1 -1 [dNTP] Af (nM) kf (s ) As (nM) ks (s ) 20/26-mer-dGC8-N-ABA

300 µM dCTP 5.6 ± 0.8 (19%) 4.6 ± 1.3 8.3 ± 0.8 (28%) 0.32 ± 0.06

600 µM dCTP 5.5 ± 0.7 (18%) 6.3 ± 1.5 8.5 ± 0.7 (28%) 0.36 ± 0.05

1000 µM dCTP 5.8 ± 0.8 (19%) 7.2 ± 1.4 8.6 ± 0.8 (29%) 0.42 ± 0.06

21/26-mer-dGC8-N-ABA

25 µM dGTP 4.4 ± 0.7 (15%) 3.8 ± 1.5 9.2 ± 0.8 (31%) 0.082 ± 0.002

50 µM dGTP 4.4 ± 0.8 (15%) 5.2 ± 1.7 9.1 ± 0.6 (30%) 0.13 ± 0.02

90 µM dGTP 4.3 ± 0.6 (14%) 11 ± 4 9.3 ± 0.7 (31%) 0.16 ± 0.03

Af and As are the reaction amplitudes of the fast and slow phases, respectively, while kf and ks are the rate constants of the fast and slow phases, respectively.

Table 2.6: Kinetic parameters from biphasic kinetic assays with hPolη at 37 °C

-1 -1 Substrate Af (nM) kf (s ) As (nM) ks (s )

20/26-mer 22.4 ± 0.9 (74%)a 76 ± 7 4.2 ± 0.9 (14%)a 4.6 ± 1.7

21/26-merc 7.8 ± 0.3 (39%)b 72 ± 5 9.5 ± 0.3 (47%)b 4.4 ± 0.5

a Calculated as (reaction amplitude/30 nM) x 100.

b Calculated as (reaction amplitude/20 nM) x 100.

c Values from Table S5 from Sherrer et al. 58

Table 2.7: Kinetic parameters from biphasic kinetic assays with hPolη and undamaged DNA at 37 °C.

2.7 Figures

Figure 2.1: Running start assays at 37 °C. A pre-incubated solution of 100 nM 5′-[32P]- labeled 17/26-mer DNA (A, C, E, and G) or 17/26-mer-dGC8-N-ABA (B, D, F, and H) and

100 nM hPolη (A and B), 100 nM hPolκ (C and D), 1 μM hPolι (E and F), or 1 μM hRev1 (G and H) was rapidly mixed with all four dNTPs (200 µM each) for various incubation periods before being quenched with the addition of 0.37 M EDTA. Sizes of important products are indicated. The dGC8-N-ABA lesion is located at the 21st position from the 3′ terminus of the DNA template.

A B

Figure 2.2: Standing start assays for hRev1 at 37 °C. A pre-incubated solution of 200 nM hRev1 and 200 nM control or damaged DNA was mixed with dATP, dCTP, dGTP, dTTP, or all four nucleotides (200 μM each) at 37 °C for 10 min. The DNA substrates used were 20/26-mer (A) and 20/26-mer-dGC8-N-ABA (B).

Figure 2.3: Electrophoretic mobility shift assay for 21/26-mer dGC8-N-ABA binding by hPolη at 23 °C. Solutions containing 21/26-mer dGC8-N-ABA DNA (10 nM) were incubated with varying amounts of hPolη (0-650 nM) for 15 min at room temperature (23

°C). The amount of hPolη•DNA complex formed was plotted against the total concentration of hPolη and the data were fit to Eq. 2.2 (2.2.5 Electrophoretic mobility

shift assays) to yield a Kd, DNA of 25 ± 6 nM. The inset shows the corresponding 4.5% native PAGE gel.

A B

Figure 2.4: Determination of the kinetic parameters for dCTP incorporation into 20/26- mer-dGC8-N-ABA at 37 °C. (A) A pre-incubated solution of hPolη (120 nM) and 5′-[32P]- labeled 20/26-mer-dGC8-N-ABA (20 nM) was rapidly mixed with increasing concentrations of dCTP (25 µM, ●; 50 µM, ○; 100 µM, ■; 200 µM, □; 400 µM, ♦; 600 µM, ◊; and 800

µM,▲) before being quenched by the addition of 0.37 M EDTA at the indicated time points. Product formation was plotted against time and the data were fit to Eq. 2.3 to yield the observed rate constant of product formation (kobs). (B) Each kobs value was plotted against its respective dCTP concentration and the data were fit to Eq. 2.4 to yield

-1 the maximum dCTP incorporation rate constant (kp) of 1.9 ± 0.1 s and dCTP equilibrium dissociation constant (Kd, dCTP) of 325 ± 44 µM.

A B

Figure 2.5: Biphasic kinetics of dNTP incorporation in the presence of a DNA trap at 37 °C.

A pre-incubated solution of 120 nM hPolη and 30 nM 20/26-mer-dGC8-N-ABA (A) or

21/26-mer-dGC8-N-ABA (B) was rapidly mixed with 5 µM unlabeled 20/26-mer trap DNA and an indicated concentration of dCTP (A) or dGTP (B). After various incubation periods, the reaction was quenched with the addition of 0.37 M EDTA. Product formation was plotted against time and the data were fit to Eq. 2.5 to yield Af and As, which are the reaction amplitudes of the fast and slow phases, respectively, as well as kf and ks, which are the rate constants of the fast and slow phases, respectively. The insets are magnifications of shorter time points to show the fast phase of product formation.

Chapter 3: Significant impact of divalent metal ions on the fidelity, sugar selectivity, and drug incorporation efficiency of human PrimPol

Reproduced in part with permission from Tokarsky, E.J., Wallenmeyer, P.C., Phi,

K.K., Suo, Z. (2017) Significant impact of divalent metal ions on the fidelity, sugar selectivity, and drug incorporation efficiency of human PrimPol. DNA Repair. 49, 51-59.

The full article is available at https://doi.org/10.1016/j.dnarep.2016.11.003. E. John

Tokarsky planned and performed the kinetic experiments with assistance from Petra C.

Wallenmeyer, and Kenneth K. Phi. E.J.T. and Zucai Suo analyzed the results. E.J.T. wrote the initial draft of the manuscript. Z.S. conceived the research and modified the manuscript. This work was supported by National Institutes of Health (grants ES026821,

ES024585, and ES009127) to Z.S.

3.1 Introduction

Since the discovery of human PrimPol in 2013 82; 83; 85, there has been extensive research to determine its precise role during genome replication in vivo. PrimPol is a bifunctional enzyme that is able to prime single-stranded DNA and subsequently catalyze primer extension via a polymerase-like, nucleotidyl transfer activity. PrimPol is only the second primase to be identified in humans and contains two distinct domains: an evolutionarily conserved Archaeo-Eukaryotic Primase (AEP) domain, and a UL52-like zinc finger domain that is required for its primase activity 84. Interestingly, PrimPol is found in both the nucleus and the mitochondria and has been implicated in replication fork progression and restart, as well as DNA lesion bypass 82; 85; 86; 135. Gene silencing of

PrimPol in human cells causes profound arrest in mitochondrial DNA (mtDNA) synthesis, decreased replication fork progression rates, and increased replication protein

A (RPA) foci, which is indicative of replicative stress 82; 85; 86; 136. In PrimPol-/- derived mouse embryonic fibroblasts, the presence of chromosomal aberrations such as chromatid breaks 83; 85 and micronuclei 86 indicate that PrimPol is essential in the maintenance of genomic integrity. PrimPol is only the second enzyme with DNA polymerase activity to be identified in mitochondria, the other being DNA polymerase γ which is responsible for rapid and faithful mtDNA replication 137; 138. Furthermore,

PrimPol is found to play a key role in UV damage resistance, as cells lacking PrimPol have increased sensitivity to UV-C radiation 85 and GFP-tagged PrimPol was shown to be rapidly recruited to chromatin in cells treated with UV-A radiation 86. Consistently, 59

PrimPol can bypass common UV-induced lesions such as cis-syn thymine-thymine cyclobutane pyrimidine dimers and (6-4) photoproducts in vitro 85; 86. PrimPol has also been shown to bypass common single-base lesions including 8-oxoguanine 135 and abasic sites 82; 86; 135. During DNA synthesis with undamaged DNA templates, PrimPol makes one error per 104-105 nucleotide incorporations in the presence of Mg2+ as measured by lacZ, HSV-tk, and M13mp2 forward mutation assays and the errors are primarily base insertions and deletions, not substitutions 84; 87; 139. In vitro, the primase and polymerase activities of PrimPol 84; 135; 139 are shown to be differentially influenced by Mg2+ and

Mn2+, the known divalent metal ion cofactors utilized by other primases and polymerases for catalysis. However, such an effect of the divalent metal ion cofactor has not been quantitatively and rigorously analyzed. To fill the void, we employed fluorescence anisotropy and pre-steady-state kinetic assays to determine the effect of the divalent metal ion cofactor on the DNA binding affinity and nucleotide incorporation efficiency of human PrimPol. In addition, we used pre-steady-state kinetics to determine the ability of human PrimPol to discriminate against ribonucleotides (rNTPs) and to incorporate the triphosphates of four nucleoside analog drugs in the presence of Mn2+ or Mg2+.

3.2 Materials and Methods

3.2.1 Materials.

Reagents were purchased from the following companies: OptiKinase from USB corp., [γ-32P]-ATP from PerkinElmer, deoxyribonucleotides (dNTPs) and rNTPs from

Bioline, 2′-deoxy-2′,2′-difluorodeoxycytidine 5′-triphosphate (GemCTP) and 2′-

aracytidine 5′-triphosphate (AraCTP) from TriLink BioTechnologies, 5′-triphosphate of lamivudine ((-)3TC-TP) and emtricitabine ((-)FTC-TP) from Gilead Sciences, and oligonucleotides from Integrated DNA Technologies.

3.2.2 Expression and Purification of Human PrimPol.

Human PrimPol containing an N-terminal 6x-histidine tag was subcloned as previously described85, and transformed into Escherichia coli Rosetta (DE3) competent cells. A single colony was used to inoculate 100 mL of LB media (30 µg/mL kanamycin and 34 µg/mL chloramphenicol) and the culture was grown at 37 °C overnight to an

OD600 of 1.5. The starter culture was then used to inoculate 6 x 1 L of fresh LB media (30

µg/mL kanamycin and 34 µg/mL chloramphenicol) and the overexpression cultures were grown at 37 °C to an OD600 of 0.8 followed by rapid cooling on ice. The cultures were induced at an OD600 of 1.0 with 0.1 mM Isopropyl β-D-1-thiogalactopyranoside and allowed to grow at 16 °C for an additional 15 hours following induction before pelleting by centrifugation. The cell pellet was re-suspended in buffer A (50 mM Tris-HCl pH 7.5,

300 mM NaCl, 10% glycerol, 0.1% β-mercaptoethanol, 10 mM imidazole, 0.01 mM

EDTA, and 0.1% IGEPAL) and supplemented with EDTA-free Protein Inhibitor Cocktail tablets (Roche) and 1 mM phenylmethanesulfonylfluoride. Cells were lysed by three passages through a French pressure cell press at 15,000 psi. The soluble fraction was isolated by ultracentrifugation at 40,000 rpm for 40 min. The cleared lysate was then incubated with charged nickel nitrilotriacetic acid (Ni-NTA) resin for 3 hr at 4 °C. The

Ni-NTA beads were packed into a tricorn FPLC column and were washed with 20

column volumes (CV) of buffer A and further washed with 10 CV of 4% buffer B (buffer

A containing 500 mM imidazole). Protein was eluted with a linear gradient of 4 to 100% buffer B over 15 CV and fractions were analyzed by SDS-PAGE. Protein-containing fractions were pooled and then loaded onto a HiTrap Heparin HP column (GE

Healthcare). The column was washed with 5 CV buffer C (buffer A without imidazole) and then with 10 CV of 10% buffer D (buffer C containing 1 M NaCl). The protein was eluted with a gradient of 10 to 100% buffer D over 10 CV. Stepwise dialysis was performed overnight at 4°C to decrease the NaCl concentration of the eluted protein solution from 700 mM to 125 mM. The purest samples were pooled and concentrated to

500 µL using an Amicon Ultra-15 Centrifugal filter (Millipore). The protein sample was further purified in a Superdex 200 size exclusion chromatography column (GE

Healthcare) to isolate full-length human PrimPol (66.5 kDa). Fractions were analyzed via

SDS-PAGE and the most pure samples were pooled and dialyzed against storage buffer

(50 mM Tris-HCl, pH 7.5, 400 mM NaCl, 50% glycerol, 1 mM DTT, and 0.1 mM

-1 -1 EDTA). Using the predicted extinction coefficient (ε280 = 77,655 M cm ), the final concentration of purified PrimPol was determined by UV-Vis spectroscopy at 280 nm and the final yield of protein was 1 mg/L of culture.

3.2.3 Radiolabeling and annealing DNA substrates.

All oligonucleotides were purified via PAGE and reverse-phase chromatography

(Sep-Pak classic C-18 cartridges). The 21-mer primer used for the single-turnover assays was 5′-[32P]-labeled by incubating it with [γ-32P]-ATP and OptiKinase (USB) for 3 hr at

37 °C. The reaction was terminated by heating at 95 °C for 2 min to denature Optikinase.

The radiolabeled primer was purified from any unreacted [γ-32P]-ATP using a Bio-spin 6 column (Bio-Rad). The primer was then annealed to a DNA template (Table 3.1) in a

1:1.35 molar ratio by first heating the reaction mixture to 95 °C for 5 min and then slowly cooling the mixture to room temperature overnight.

3.2.4 Fluorescence anisotropy titration.

The Cy3-labeled DNA substrate 17/30-mer (Figure 3.1A, 30 nM) was titrated with increasing amounts of PrimPol and the anisotropy was monitored using a

FluoroMax-4 (Horiba). Assays were carried out at 25 °C in buffer E [50 mM Tris–HCl

(pH = 7.5 at 25 °C), 50 mM NaCl, 0.01 mM EDTA] without divalent metal ions or in the presence of 5 mM MnCl2, or 5 mM MgCl2 where indicated. Excitation and emission for the Cy3 fluorophore were set to 540 and 568 nm, respectively, with a 10 nm slit width and 2 s integration time. The data obtained from anisotropy measurements were fit to Eq.

3.1:

2 ½ ΔA = (ΔAT /2D0) x [(Kd,DNA + D0 + E0) – [(Kd,DNA + D0 + E0) − 4 E0D0] ] Eq. 3.1

Where ΔA is the change in anisotropy, ΔAT is the maximum anisotropy change, D0 and

E0 are the initial concentrations of DNA and PrimPol, respectively, and Kd,DNA is the equilibrium dissociation constant of the PrimPol●DNA binary complex.

3.2.5 Single-turnover kinetic assays.

PrimPol (300 nM) and 5′-[32P]-labeled DNA (30 nM) were pre-incubated at 37 °C for 5 min in buffer E (pH = 7.5 at 37 °C) containing 5 mM DTT and 0.1 µg/ml BSA, before mixing with increasing concentrations of a single dNTP. Reaction mixtures (10

µL) were quenched with EDTA to a final concentration of 0.37 M at increasing time points. Reaction products were separated via PAGE (17% polyacrylamide, 8 M urea) and visualized using a PhosphorImager plate (Amersham Biosciences) and TyphoonTrio scanner (GE Healthcare). The product was quantified using ImageQuant software

(Molecular Dynamics). The plot of product concentration versus time from each time course was fit to a single-exponential equation (Eq. 3.2) using non-linear regression software KaleidaGraph (Synergy Software) to determine the observed nucleotide incorporation rate constant (kobs).

[Product] = A[1 - exp(- kobst)] Eq. 3.2 where A is the reaction amplitude, which is equal to the initial concentration of the

PrimPol●DNA binary complex. The kobs values obtained from Eq. 2 were plotted against the respective concentrations of dNTP and the data were fit to a hyperbolic equation (Eq.

3.3): kobs = kp[dNTP]/([dNTP] + Kd) Eq. 3.3

Where Kd is the apparent dissociation constant of dNTP from the PrimPol●DNA●dNTP ternary complex and kp is the maximum rate constant of dNTP incorporation.

3.3 Results

3.3.1 Binding affinity of human PrimPol to DNA in the presence of Mn2+ or Mg2+.

Human PrimPol was expressed in Escherichia coli and purified to >98% purity through column chromatography (3.2.2 Expression and Purification of Human PrimPol).

To investigate the effect of divalent metal ions on the binding affinity of human PrimPol to DNA, we employed a fluorescence anisotropy assay (3.2.4 Fluorescence anisotropy titration) and determined the dissociation equilibrium constant (Kd,DNA) for the

PrimPol●DNA binary complex in the presence, or absence, of 5 mM MnCl2 or MgCl2.

Similar concentrations of the divalent metal ions have been used in published studies of human PrimPol 84; 135; 139. In the absence of any supplemental divalent metal ions, the

Kd,DNA for the binding of PrimPol to the Cy3-labeled DNA substrate 17/30-mer (Figure

3.1A) was measured to be 41 ± 5 nM (Figure 3.1B). In the presence of 5 mM Mn2+ or

2+ Mg , the Kd,DNA value was determined to be 29 ± 5 or 979 ± 119 nM, respectively

(Figures 3.1B and 3.1C). Thus, the affinity (Kd,DNA = 979 nM) of our prepared PrimPol from E. coli to DNA in the presence of 5 mM Mg2+ is ~24-fold lower than in the absence

2+ (Kd,DNA = 41 nM) while the presence of 5 mM Mn slightly enhanced the affinity (Kd,DNA

= 29 nM). This suggests that the presence of a molar excess of Mg2+ likely disrupted the interaction between the Zn2+ and its coordinating amino acid residues (Cys-His-Cys-

Cys), which subsequently altered the binding of the zinc finger domain to the single- stranded template region in the DNA substrate 84 (Figure 3.1A) and weakened the overall

DNA binding affinity of PrimPol (Figure 3.1C). Previously, similarly prepared human

PrimPol to ours was found to contain Zn2+ in its C-terminal zinc finger domain with an occupancy of ~80% measured by using inductively coupled plasma mass spectrometry

(ICP-MS) 84. This suggests that the presence of 5 mM Mn2+, a transition divalent metal

ion known to bind to zinc finger motifs 140; 141, likely bound to 20% of our prepared

PrimPol, which lacked Zn2+ based on the published ICP-MS assay results 84, and slightly improved the DNA binding affinity of our PrimPol (Figure 3.1B).

3.3.2 Correct nucleotide incorporation efficiency in the presence of Mn2+ or Mg2+.

Besides DNA binding, Mn2+ and Mg2+ may impact the nucleotide incorporation efficiency of PrimPol differently since its AEP domain binds to divalent metal ions. To examine this possibility, single-turnover kinetic assays were performed by mixing a pre- incubated solution of PrimPol (300 nM) and 5′-[32P]-labeled 21/41-mer DNA substrate

D-7 (30 nM, Table 3.1) with increasing concentrations of correct dATP in the presence of 5 mM Mn2+ for various times before being quenched with 0.37 M EDTA (3.2.5

Single-turnover kinetic assays). After the reaction products were separated and quantitated and the time courses were analyzed, we determined the maximum dATP

-1 incorporation rate constant (kp) of 0.066 ± 0.003 s and the apparent equilibrium dissociation constant (Kd) of 11 ± 1 µM for dATP binding (Figure 3.2). The substrate

-3 -1 -1 specificity (kp/Kd) of dATP was then calculated to be 6.0x10 µM s (Table 3.2).

Similar single-turnover kinetic assays were performed for correct dTTP, dCTP, dGTP incorporation onto D-1, D-6, and D-8 (Table 3.1), respectively, and the kinetic parameters are listed in Table 3.2. In the presence of 5 mM Mn2+, correct dNTP

-1 incorporation occurs with a kp in the range of 0.036-0.096 s , a Kd of 11-17 µM, and a

-3 -1 -1 kp/Kd of (2.3-6.0)x10 µM s (Table 3.2). For comparison, we performed similar single- turnover kinetic assays for a single correct dNTP incorporation onto a corresponding

DNA substrate (Table 3.1) by PrimPol in the presence of 5 mM Mg2+, e.g. dGTP onto D-

8. The kinetic data (Table 3.3) show that PrimPol incorporates correct dNTPs in the

2+ -1 presence of 5 mM Mg with a kp of 0.011-0.020 s , a Kd of 262-895 µM, and a kp/Kd of

(2.2-5.0)x10-5 µM-1s-1. Thus, on average, the presence of 5 mM Mg2+ makes PrimPol about 100-fold less efficient during correct dNTP incorporation than in the presence of 5 mM Mn2+ (Tables 3.2 and 3.3).

3.3.3 Substitution fidelity of human PrimPol in the presence of Mn2+ or Mg2+.

To kinetically estimate the incorporation fidelity of PrimPol, we employed similar single-turnover assays to determine the pre-steady-state kinetic parameters (Table 3.2) for the 12 possible incorrect incorporations in the presence of 5 mM Mn2+, e.g. dGTP incorporation opposite dT. Similar approaches have been used by us to determine the fidelities of other DNA polymerases 37; 142; 143; 144; 145. The kinetic data in Table 3.2 demonstrate that PrimPol incorporates incorrect dNTPs in the presence of 5 mM Mn2+

-1 -5 -3 -1 -1 with a kp of 0.0024-0.035 s , a Kd of 3-30 µM, and a kp/Kd of (8.0x10 -2.0x10 ) µM s .

Relative to the kinetic parameters with correct dNTP incorporations (see above), incorrect dNTP incorporations occur with lower kp and kp/Kd values but with comparable

Kd values. The substitution fidelity of PrimPol, defined as (kp/Kd)incorrect/[(kp /Kd)incorrect +

2+ -2 (kp/Kd)correct], in the presence of 5 mM Mn is calculated to be in the range of 3.4x10 to

3.8x10-1 (Table 3.2). This fidelity indicates that PrimPol tends to make one substitution error out of every 4 to 29 nucleotide incorporations in the presence of Mn2+. This substitution fidelity is about 100-fold lower than the fidelity range of 10-2-10-4 previously

estimated by Zafar et. al. through steady-state kinetic methods in the presence of 10 mM

Mn2+ 135. The large discrepancy is likely because the steady-state kinetic parameters are complicated by fast DNA product dissociation146 and are inaccurately determined when nucleotide incorporation, e.g. misincorporation, is inefficient and yields little products.

The low fidelity of PrimPol can be illustrated in the gel images of the time courses of incorrect dNTP incorporations, especially when multiple dCTP misincorporations occurred and even made the products longer than the template 41-mer in the presence of

Mn2+ (Figure 3.3). Similar template-independent dCTP misincorporations by PrimPol have been observed previously 84; 139.

To examine if the switch from Mn2+ to Mg2+ increases the fidelity of PrimPol, we determined the kinetic parameters for individual misincorporations onto D-7 (Table 3.1) in the presence of 5 mM Mg2+. Opposite dT, each incorrect dNTP was inefficiently incorporated and only a small amount of products were formed even after 3 hours of incubation (Figure 3.4C). As a result, we instead determined the kp/Kd value, not individual kp and Kd values, for each dNTP misincorporation and calculated the fidelity

(10-2-10-4) of PrimPol (Table 3.3). Notably, the dCTP:dT misincorporation occurred with a 100-fold lower fidelity value than the other two misincorporations (Table 3.3). This is likely due to two consecutive dCTP incorporations onto D-7 (Table 3.1) with the second being a correct incorporation, while dGTP or dTTP was only misincorporated once onto

D-7 (Figure 3.4), leading to a 100-fold difference in their kp/Kd values (Table 3.3). Thus, the substitution fidelity of PrimPol in the presence of Mg2+ is estimated to be ~10-4, which is 100-1,000 fold higher than its fidelity in the presence of Mn2+ (Table 3.2). Our

fidelity value of ~10-4 is also comparable with the fidelity values estimated from lacZ,

HSV-tk, and M13mp2 forward mutation assays in the presence of Mg2+ 84; 87; 139.

3.3.4 Sugar selectivity of human PrimPol in the presence of Mn2+ and Mg2+.

Eukaryotic primases are responsible for catalyzing de novo primer synthesis on single-stranded DNA for the initiation of leading and lagging strand synthesis147.

Specifically, primases utilize rNTPs to catalyze de novo RNA primer synthesis (7-10 bases), which can then be extended by a specialized DNA polymerase in the presence of dNTPs to generate a hybrid 5-RNA-DNA-3 primer for DNA replication148.

Contrastingly, human PrimPol prefers to utilize dNTPs for both de novo primer synthesis82; 85 and subsequent primer extension139. Replicative polymerases can extend

PrimPol-generated DNA primers during lesion bypass and replication fork restart, as there is no need for further processing of the RNA strands81. To quantitatively determine how much human PrimPol discriminates against rNTPs, we performed single-turnover kinetic assays (3.2.5 Single-turnover kinetic assays) to investigate matched rNTP incorporation onto DNA/DNA substrates (Table 3.1) in the presence of either 5 mM

MnCl2 or 5 mM MgCl2 (data not shown). PrimPol incorporated matched rNTPs with a

-1 range of kp values from 0.0015-0.0092 s and a range of Kd values of 110-744 µM,

-6 -5 -1 -1 yielding calculated kp/Kd values of 3.2x10 to 6.8x10 µM s in the presence of 5 mM

Mn2+ (Table 3.2). The ratio of catalytic efficiencies of correct dNTPs over matched rNTPs [(kp/Kd)dNTP/(kp/Kd)rNTP] (Table 3.2), was calculated to give the sugar selectivity of

PrimPol, which has been done similarly for other DNA polymerases 149; 150; 151; 152; 153.

The sugar selectivity of PrimPol, was found to be 57-1,800 in the presence of 5 mM Mn2+

(Table 3.2). Interestingly, both a decrease in the kp (24-fold on average) and an increase in the Kd (30-fold on average) are responsible for rNTP discrimination by human

PrimPol.

2+ In the presence of Mg , the individual kinetic parameters (kp and Kd) could not be determined due to extremely slow incorporation of matched rNTPs (see above). Instead, we obtained kobs values for each matched rNTP at 1 mM concentrations, and found that

PrimPol incorporated matched rNTPs in the range of (6.8-9.5)x10-5 s-1 (Table 3.4). Since the kobs for correct incorporation of each dNTP at 1 mM was determined above, the ratios of observed nucleotide incorporation rate constants (kobs, dNTP/kobs, rNTP) were calculated to be in the range of 105-260 (Table 3.4). Thus, PrimPol incorporated correct dNTPs 150- fold (on average) more rapidly than matched rNTPs in the presence of Mg2+. Although

2+ we could not determine the Kd, rNTP/Kd, dNTP ratios in the presence of Mg , we assume they are similar to the average 30-fold binding affinity ratio determined in the presence of

Mn2+ (see above). Together, our kinetic data indicate that the sugar selectivity of human

PrimPol in the presence of Mg2+ will be greater than 150 and could be as high as 4,500.

3.3.5 Incorporation efficiencies of the triphosphates of four cytidine analog drugs gemcitabine, cytarabine, emtricitabine, and lamivudine by human PrimPol.

Chain-terminating nucleoside analogs have been developed as chemotherapeutic drugs to treat cancers and antiviral drugs to combat viral infections. For example, gemcitabine (GemC) and cytarabine (AraC) have been used to treat pancreatic

adenocarcinoma154, non-muscle invasive bladder cancer155, non-small cell lung cancer72;

156; 157, and acute myeloid leukemia158 while lamivudine ((-)3TC) and emtricitabine ((-

)FTC) are widely prescribed drugs against human immunodeficiency virus (HIV) and

Hepatitis B (HBV) infections 159; 160; 161. The chain-terminating nucleoside analog drugs are cellularly activated into their triphosphate forms which compete against natural dNTPs for incorporation and terminate cellular and viral genomic replication once incorporated. Besides clinical efficacy, the drugs also cause various clinical toxicities and some of them have not been explained well at a molecular level. Considering human

PrimPol is a recently discovered enzyme, it could incorporate the triphosphates of various nucleoside analog drugs and contribute to their observed clinical toxicities. Here, we investigated the abilities of human PrimPol to incorporate the triphosphates of four cytidine analogs (Figure 3.5) by performing similar single-turnover kinetic assays (3.2.5

Single-turnover kinetic assays) in the presence of 5 mM MnCl2 or MgCl2. The measured

2+ kinetic data are listed in Table 3.5. In the presence of 5 mM Mn , AraCTP (Kd = 21 ± 4

µM) was bound by PrimPol with a similar affinity as dCTP (Kd = 16 ± 4 µM). However,

-1 the incorporation of AraCTP (kp = 0.0057 ± 0.0004 s ) was 10-fold slower than dCTP (kp

= 0.060 ± 0.005 s-1), leading to a discrimination value, calculated as

[(kp/Kd)dCTP/(kp/Kd)analog], of 14 (Table 3.4). In comparison, GemCTP was incorporated with similar kp and Kd values as dCTP, leading to a discrimination value of 2.9 (Table

3.5).

In the presence of 5 mM Mg2+, both AraCTP and GemCTP were incorporated by

PrimPol with 2-fold lower kp values than dCTP. However, the Kd values of AraCTP and

dCTP are comparable while GemCTP was bound with a 5-fold higher Kd. The discrimination factors were calculated to be 2.1 and 9.8 for AraCTP and GemCTP, respectively (Table 3.5).

For (-)3TC-TP and (-)FTC-TP, the two antiviral drugs with L-stereochemistry, we also kinetically investigated their incorporation by human PrimPol. In the presence of 5 mM Mn2+, both (-)3TC-TP and (-)FTC-TP were incorporated inefficiently and only their

-6 -7 -1 -1 kp/Kd values were determined to be 1.2x10 and 9.0x10 µM s , respectively (Table

3.5). In the presence of 5 mM Mg2+, these L-analogs were not incorporated by PrimPol even after 3 hours at 37 °C (data not shown).

3.4. Discussion

In this study, we employed a fluorescence anisotropy assay and determined the

Kd,DNA values for the PrimPol●DNA binary complex in the presence or absence of 5 mM

Mn2+ or Mg2+ (Figure 3.1). Strikingly, in the presence of Mg2+, the affinity of PrimPol to

DNA (Kd,DNA = 979 nM) is significantly lower than other human DNA polymerases

(Table 3.6) and the switch from Mg2+ to Mn2+ increased the DNA binding affinity

(Kd,DNA = 29 nM) of PrimPol by 34-fold. The 29 nM DNA binding affinity is likely caused by the C-terminal zinc finger domain of PrimPol which binds to the single- stranded template region in a primer/template substrate and increases the DNA binding affinity of PrimPol 84. Since Mn2+ can substitute Zn2+ and bind to zinc finger motifs 140;

141 while Mg2+ cannot, the presence of a large molar excess of Mg2+ (5 mM) in our fluorescence anisotropy assay (Figure 3.1) likely disrupted the zinc finger domain of

PrimPol and significantly decreased its DNA binding affinity. Furthermore, the substrate specificity (kp/Kd) values for correct dNTP incorporations determined through pre-steady- state kinetic assays (Figure 3.2) are 100-fold higher with 5 mM Mn2+ ((2.3-6.0)x10-3 µM-

1s-1, Table 3.2) than with 5 mM Mg2+ ((2.2-3.8)x10-5 µM-1s-1, Table 3.3). The 100-fold efficiency difference is mainly contributed by the ~48-fold dNTP binding affinity ratio

2+ 2+ since the kp was only ~2-fold lower with Mg versus Mn (Tables 3.2 and 3.3). After multiplying the kp/Kd difference by the 34-fold DNA binding affinity ratio, PrimPol is

3,400-fold less efficient as a DNA polymerase to bind and elongate a DNA/DNA substrate in the presence of 5 mM Mg2+ over an equal concentration of Mn2+.

Considering the very low DNA and dNTP binding affinities and correct dNTP incorporation rates and efficiencies relative to other kinetically characterized human

DNA polymerases in the presence of Mg2+ (Table 3.6), PrimPol is incapable of competing for access to DNA and is not efficient enough to function as a meaningful

DNA polymerase during DNA replication and lesion bypass in vivo. For example,

PrimPol is 2x106 fold less efficient than human DNA polymerase γ with Mg2+ and may not be able to play a role in mitochondrial DNA replication. However, in the presence of

Mn2+, PrimPol bound both DNA (Figure 3.1) and dNTPs (Table 3.2) with moderately high affinities and incorporated correct dNTPs with comparable efficiencies as human

DNA polymerases µ, κ, and ι with Mg2+ (Table 3.6). In parallel, the primase activity of

PrimPol has been barely observed in the presence of 10 mM Mg2+ but is activated by

Mn2+ with a concentration as low as 50 µM 82. Together, these results suggest that the

AEP domain of PrimPol may utilize Mn2+, not Mg2+, as the divalent metal ion cofactor in

order to carry out both polymerase and primase activities in vivo. Recently, Polymerase

Delta Interacting Protein 2 (PolDIP2) has been shown to interact with the AEP domain and moderately enhance the DNA binding affinity and polymerase processivity of human

PrimPol in the presence of Mg2+ 162. If the enhancement is also true in the presence of

Mn2+, the combined favorable effect of PolDIP2 and Mn2+ will further solidify the role of

PrimPol as a legitimate bifunctional enzyme in replication fork progression and restart, as well as DNA lesion bypass 82; 85; 86; 135. More studies, especially structural investigation, are needed to verify the modulation effect of both PolDIP2 and Mn2+ on the structure and function of PrimPol.

Interestingly, some of the thermodynamic and kinetic parameters reported here are different than those reported by Mislak and Anderson (Antimicrob. Agents

163 163 Chemother. 60, 561-569) in 2016 . Specifically, their reported Kd,DNA values , measured by electrophoretic mobility shift assays (EMSAs), are larger than ours by 11- to

25-fold in the presence of Mn2+ and 8-fold in the presence of Mg2+. The relative

2+ enhancement in the DNA binding affinities by switching from Mg (Kd,DNA = 340-720

2+ 163 nM) to Mn (Kd,DNA = 8 µM) in their studies is comparable to the 34-fold DNA binding affinity enhancement reported here. The large discrepancy between our and their

Kd,DNA values is likely due to the different methods used the two studies. The EMSAs employed by Mislak and Anderson 163 were not performed under true equilibrium conditions. In comparison, our fluorescence anisotropy method measures the protein and

DNA binding and dissociation at equilibrium in solution, and polarization was measured promptly after equilibrium was achieved. Furthermore, the sole variable to affect

equilibrium in an fluorescence anisotropy assay is the concentration of DNA or PrimPol in their binding interactions while the gel matrix and electrical current can perturb the protein-DNA complex formation in an EMSA. Likewise, the slightly lower kp and kp/Kd values measured here may have been caused by differences in the reaction buffers used in our single-turnover assays (3.2.5 Single-turnover kinetic assays) and those used by

Mislak and Anderson (10 mM Bis-Tris propane, pH = 7.0, 1 mM DTT, 10 mM MnCl2)

163. Furthermore, the slow single-turnover rates (Tables 3.2 and 3.3) measured here are comparable to the steady-state rates reported by Zafar et al.135 Therefore, we expect not to observe an initial burst of product formation under pre-steady-state kinetic conditions in which the DNA substrate is only a few fold molar excess over the enzyme.

Additionally, it should be noted that PrimPol is an atypical DNA polymerase and its kinetic mechanism is likely to be different from those of canonical polymerases. Thus, the assignments of the steady-state and pre-steady state rates within the mechanism of polymerization catalyzed by PrimPol remain to be established.

3.4.1 The polymerase activity of human PrimPol is error-prone in the presence of Mn2+.

The substitution fidelity of PrimPol was kinetically determined to be in the range of 3.4x10-2 to 3.8x10-1 in the presence of Mn2+ (Table 3.2). Thus, PrimPol poorly discriminated against incorrect dNTPs during primer elongation. The fidelity of PrimPol is actually comparable to the low fidelities of human Y-family DNA polymerases which bypass DNA lesions in vivo 164. PrimPol incorporated an incorrect dNTP with only 2- to

23-fold slower kp than a correct dNTP and bound to all dNTPs with comparable affinities

(Kd = 3-30 µM), demonstrating a kinetic basis for the low fidelity of PrimPol. These data also indicate that incorrect dNTPs are strong competitive inhibitors against correct dNTPs during polymerization in the presence of Mn2+. Although significantly less efficient, PrimPol has a fidelity of 10-4 in the presence of Mg2+ (Table 3.3) which is 100-

1,000 fold more faithful than in the presence of Mn2+ (Table 3.2). Consistently, the switch from Mg2+ to Mn2+ as the divalent metal ion cofactor is known to increase misincorporation frequency for many DNA polymerases and can lead to increased genomic instability 103; 165; 166; 167; 168.

3.4.2 Human PrimPol displays moderate sugar selectivity in the presence of Mn2+ and

Mg2+.

Human PrimPol has been shown to select dNTPs over rNTPs during primer extension with both DNA and RNA primers 139. Consistently, our kinetic studies quantitatively estimated the sugar selectivity of PrimPol to be 57-1,800 in the presence of

Mn2+ (Table 3.2) and 150-4,500 in the presence of Mg2+ (see 3.3.4 in Results). Thus, human PrimPol displays a modest sugar selectivity (740 on average with Mn2+) 45, which is comparable to the sugar selectivities of human DNA polymerases α (500), γ (1,000) and Rev1 (280), measured using similar kinetic methods72; 169; 170. Interestingly, a very recently published crystal structure of the AEP domain of human PrimPol in complex with DNA and an incoming dATP 171 was modeled with an rATP to replace the dATP in the enzyme active site. Notably, the 2′ hydroxyl of the rATP ribose sterically clashed with the backbone carbonyl of Asn289. Thus, the 67-fold difference between the Kd

values of correct dATP and matched rATP in the presence of Mn2+ (Table 3.2) likely arises from the clash between the rATP and the steric gate residue Asn289 45, similar to what has been observed with other DNA polymerases 30; 31; 32; 44; 45, e.g. rNTP with Tyr12 in Sulfolobus solfataricus DNA polymerase IV (Dpo4) 172; 173. Furthermore, since the

AEP domain of PrimPol contains a single active site that catalyzes both primase and polymerase activities82; 83; 85, the same mode of sugar selection is likely utilized by both the primase and polymerase activities of PrimPol. Our ongoing structural and mutagenic studies of PrimPol will further characterize the steric gate residue in this bifunctional enzyme.

3.4.3 Human PrimPol was able to incorporate the triphosphates of gemcitabine and cytarabine, but not emtricitabine and lamivudine.

Nucleoside Reverse Transcriptase Inhibitors (NRTIs) specifically target the active site of RTs and halt viral genomic replication. Mitochondrial toxicity of NRTIs has been associated with the incorporation of the triphosphates of NRTIs by human DNA polymerase γ, a main off-target issue. Based on the existence of PrimPol in the mitochondria, it is possible that PrimPol could potentially incorporate NRTIs and contribute to their mitochondrial toxicology. Recently, human PrimPol has been shown to incorporate the triphosphates of anti-HIV NRTI drugs zidovudine and abacavir 163 and reprime downstream from NRTI-terminated DNA primers 174. In this paper, we found that human PrimPol incorporated the triphosphates of antiviral drugs emtricitabine ((-)3TC-

TP) and lamivudine ((-)FTC-TP) with extremely low efficiencies (10-6 to 10-7 μM-1 s-1) in

the presence of Mn2+ (Table 4) but did not incorporate them even after 3 hours in the presence of Mg2+ at 37 °C. Since (-)3TC-TP and (-)FTC-TP, unlike natural dNTPs, possess L-stereochemistry (Figure 3.5), our results suggest that PrimPol, like Dpo4175 and human DNA polymerase 161, possesses strong D-stereoselectivity. Our results further suggest that human PrimPol is an unlikely off-target of these two drugs in vivo.

In comparison, human PrimPol incorporated AraCTP, GemCTP, and dCTP, which possess D-stereochemistry (Figure 3.5), with comparable kp/Kd values in the presence of Mn2+ or Mg2+, leading to the discrimination factors in the range of 2-14

(Table 3.4). It suggests that the inhibition of PrimPol by these anticancer chain- terminating nucleotide analogs likely contributes to their clinical toxicities. Interestingly, there is an order for the Kd values: dCTP < AraCTP < GemCTP < rCTP (Tables 3.2-

3.4). This order is closely correlated to the size of the 2-group oriented below the sugar ring, likely due to its clash with an unidentified steric gate residue in PrimPol (see above). As the size of the 2-group increases, the Kd value increases simultaneously.

Specifically, the 2-group is a small hydrogen atom in both dCTP and AraCTP, a medium sized fluorine atom in GemCTP, and a comparatively larger sized hydroxyl group in rCTP (Figure 3.5), which leads to the above Kd order. The slightly larger Kd value of

AraCTP over dCTP may be contributed by the size difference between their 2-groups oriented above the sugar ring.

3.4.4 Conclusion.

In summary, our fluorescence anisotropy and pre-steady-state kinetic studies demonstrate that human PrimPol is a moderately efficient and highly unfaithful DNA polymerase in the presence of Mn2+ but is too inefficient to function as a DNA polymerase in the presence of Mg2+. Human PrimPol possesses modest sugar selectivity and can incorporate the triphosphates of anticancer gemcitabine and cytarabine, but not antiviral emtricitabine and lamivudine.

3.5 Tables

D-1

5′-CGCAGCCGTCCAACCAACTCA-3′

3′-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5′

D-6

5′-CGCAGCCGTCCAACCAACTCA-3′

3′-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5′

D-7

5′-CGCAGCCGTCCAACCAACTCA-3′

3′-GCGTCGGCAGGTTGGTTGAGTTGCAGCTAGGTTACGGCAGG-5′

D-8

5′-CGCAGCCGTCCAACCAACTCA-3′

3′-GCGTCGGCAGGTTGGTTGAGTCGCAGCTAGGTTACGGCAGG-5′

Table 3.1: 21/41-mer DNA substrates

Sugar dNTP k (s-1) K (μM) k /K (μM-1 s-1) Fidelitya p d p d Selectivityb D-1 (Template dA) dTTP 0.096 ± 0.004 17 ± 2 5.8 x 10-3 - dATP 0.0083 ± 0.0001 4.1 ± 0.3 2.0 x 10-3 2.6 x 10-1 dCTP 0.014 ± 0.0009 13 ± 4 1.1 x 10-3 1.7 x 10-1 dGTP 0.0047 ± 0.0007 13 ± 4 3.6 x 10-4 6.0 x 10-2 rUTP 0.0018 ± 0.0002 561 ± 160 3.2 x 10-6 5.5 x 10-4 1800 D-6 (Template dG) dCTP 0.060 ± 0.005 16 ± 4 3.8 x 10-3 - dATP 0.011 ± 0.00005 20 ± 3 5.5 x 10-4 1.3 x 10-1 dGTP 0.035 ± 0.003 27 ± 8 1.3 x 10-3 2.6 x 10-1 dTTP 0.0058 ± 0.0005 14 ± 6 4.2 x 10-4 9.9 x 10-2 rCTP 0.0092 ± 0.0002 136 ± 7 6.7 x 10-5 1.7 x 10-2 57 D-7 (Template dT) dATP 0.066 ± 0.003 11 ± 1 6.0 x 10-3 - dCTP 0.013 ± 0.002 17 ± 9 7.6 x 10-4 1.1 x 10-1 dGTP 0.017 ± 0.001 22 ± 4 7.7 x 10-4 1.1 x 10-1 dTTP 0.0029 ± 0.0002 3 ± 1 1.0 x 10-3 1.4 x 10-1 rATP 0.0047 ± 0.0006 744 ± 234 6.3 x 10-6 1.0 x 10-3 950 D-8 (Template dC) dGTP 0.036 ± 0.002 16 ± 3 2.3 x 10-3 - dATP 0.0049 ± 0.0003 16 ± 3 3.0 x 10-4 1.2 x 10-1 dCTP 0.014 ± 0.001 10 ± 3 1.4 x 10-3 3.8 x 10-1 dTTP 0.0024 ± 0.0004 30 ± 7 8.0 x 10-5 3.4 x 10-2 rGTP 0.0015 ± 0.0001 110 ± 29 1.4 x 10-5 6.1 x 10-3 160 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect] b Calculated as (kp/Kd)dNTP/(kp/Kd)rNTP

Table 3.2: Pre-steady-state kinetic parameters for nucleotide incorporation onto

21/41-mer DNA substrates catalyzed by PrimPol in the presence of 5 mM Mn2+ at 37

C.

-1 -1 -1 a dNTP kp (s ) Kd (μM) kp/Kd (μM s ) Fidelity

D-1 (Template dA) dTTP 0.020 ± 0.002 526 ± 99 3.8 x 10-5

D-6 (Template dG) dCTP 0.013 ± 0.0008 262 ± 55 5.0 x 10-5

D-7 (Template dT) dATP 0.011 ± 0.0007 388 ± 65 2.8 x 10-5 - dCTP Not determined Not determined 3.1 x 10-7 1.1 x 10-2 dGTP Not determined Not determined 1.3 x 10-8 4.6 x 10-4 dTTP Not determined Not determined 1.7 x 10-8 6.1 x 10-4

D-8 (Template dC) dGTP 0.020 ± 0.0006 895 ± 60 2.2 x 10-5 a Calculated as (kp/Kd)incorrect/[(kp/Kd)correct + (kp/Kd)incorrect]

Table 3.3: Pre-steady-state kinetic parameters for dNTP incorporation onto 21/41- mer DNA substrates catalyzed by PrimPol in the presence of 5 mM Mg2+ at 37 C.

-1 Nucleotide kobs (s ) kobs, dNTP/kobs, rNTP D-1 (Template dA) dTTP 0.013 ± 0.001 - rUTP 6.8 x 10-5 260

D-6 (Template dG) dCTP 0.010 ± 0.001 - rCTP 9.5 x 10-5 105

D-7 (Template dT) dATP 0.0086 ± 0.001 - rATP 7.7 x 10-5 110

D-8 (Template dC) dGTP 0.011 ± 0.0005 - rGTP 9.0 x 10-5 120

-1 kp/Kd a NTP kp (s ) Kd (μM) Discrimination (μM-1 s-1) In the presence of 5 mM Mn2+

AraCTP 0.0057 ± 0.0004 21 ± 4 2.7 x 10-4 14

GemCTP 0.058 ± 0.0006 45 ± 2 1.3 x 10-3 2.9

(-)3TC-TP Not determined Not determined 1.2 x 10-6 3,200

(-)FTC-TP Not determined Not determined 9.0 x 10-7 4,200

In the presence of 5 mM Mg2+

AraCTP 0.0076 ± 0.0003 316 ± 44 2.4 x 10-5 2.1

GemCTP 0.0071 ± 0.0006 1,380 ± 255 5.1 x 10-6 9.8

(-)3TC-TP No observed incorporation

(-)FTC-TP No observed incorporation a Calculated as (kp/Kd)dCTP/(kp/Kd)analog with the (kp/Kd)dCTP values from Tables

3.2 and 3.3.

Table 3.5: Pre-steady-state kinetic parameters for incorporation of dCTP analogs onto D-6 (Table 3.1) catalyzed by human PrimPol at 37 C.

Human Kd,DNA Kd,dNTP kp kp/Kd,dNTP Cited DNA (nM) (µM) (s-1) (µM-1s-1) Reference Polymerase PrimPol 979 895 0.02 2.2x10-5 Table 3.3 Pol εa 79 9 219 24 36 Pol γb 39 0.8 37 45 137; 138 Pol β 12 8.7 19 2.2 160; 176 Pol λ 110 2.1 2.5 1.2 160; 177 Pol µ 165 1.8 0.055 0.031 145; 178 Pol η 26 46 39 0.84 58 Pol κ 96 87 1.0 0.011 58 Pol ι 44 117 0.19 1.6x10-3 58; 179 Rev1c 118 2.2 22.4 10 72 aThe exonuclease-deficient mutant of human Pol ε. bLarge (or catalytic) subunit of human Pol γ. cThe listed kinetic parameters were for dCTP incorporation opposite dG.

Table 3.6: Comparison of the kinetic parameters for dGTP incorporation opposite dC by PrimPol and other human DNA polymerases in the presence of Mg2+.

3.6 Figures A 5′-GCCTCGCTGCCGTCGCC-3′ 3′-CGGAGCGACGGCAGCGGTTTTTTTTTTTTT-Cy3-5′ B C

Figure 3.1: Determination of the binding affinity of PrimPol to DNA in the presence or absence of a divalent metal ion. (A) Cy3-labeled DNA 17/30-mer. Increasing amounts of PrimPol were titrated into a fixed concentration of Cy3-labeled 17/30-mer (30 nM). The plot of the concentration of the binary complex PrimPol●DNA versus the concentration of PrimPol was fit to Eq. 3.1 (Materials and Methods) to obtain Kd,DNA. The

Kd,DNA value was 41 ± 5 nM in the absence of any divalent metal ions (B), 29 ± 5 nM in

the presence of 5 mM MnCl2 (B), 979 ± 119 nM in the presence of 5 mM MgCl2 (C).

A B

Figure 3.2: Determination of kinetic parameters for dATP incorporation opposite dT in the presence of Mn2+ at 37 C. (A) A pre-incubated solution of PrimPol (300 nM) and D-7 DNA substrate (30 nM, Table 3.1) was mixed with increasing concentrations of dATP in the presence of 5 mM Mn2+ at 37 C. The product concentrations were plotted against reaction times and each time course was fit to Eq. 3.2 (3.2.5 Single-turnover kinetic assays) to yield kobs. (B) The kobs values were plotted against respective

-1 concentrations of dATP and the plot was fit to Eq. 3.3 to yield a kp of 0.066 ± 0.003 s and a Kd of 11 ± 1 µM.

dCTP Misincorporation D-1 D-7 D-8

Figure 3.3: Gel image of time courses of dNTP misincorporation catalyzed by

PrimPol in the presence of Mn2+ at 37 C. Gel images are from single-turnover kinetic assays where pre-incubated PrimPol (300 nM) and 5′-[32P]-labeled DNA (30 nM) were mixed with dNTP (indicated concentration) in the presence of 5 mM Mn2+ for indicated times before being quenched by 0.37 M EDTA.

Continued on next page

Figure 3.3 Continued dATP Misincorporation D-1 D-6 D-8

dGTP Misincorporation D-1 D-6 D-7

dTTP Misincorporation D-6 D-7 D-8

A — dCTP

B — dGTP

C — dTTP

Figure 3.4: Gel image of time courses of dNTP misincorporation opposite dT catalyzed by PrimPol in the presence of Mg2+ at 37 C. Gel images are from single- turnover kinetic assays where pre-incubated PrimPol (2 µM) and 5′-[32P]-labeled D-7

DNA substrate (Table 3.1, 5 nM) were mixed with dNTP (800 µM) in the presence of 5 mM Mg2+ for indicated times before being quenched by 0.37 M EDTA. (A) dCTP; (B) dGTP; and (C) dTTP.

A B C

dC rC AraC

D E F

GemC (-)3TC (-)FTC

Figure 3.5: Chemical structures of dC, rC, and their analogs used in this study. The triphosphates of these nucleosides were used in single-turnover kinetic assays. (A) dC;

(B) rC; (C) 2′-aracytidine (AraC); (D) 2′,2′-difluorodeoxycytidine (GemC); (E) (-)-β-L-

2′-3-dideoxy-3′-thiacytidine ((-)3TC); and (F) (-)-β -L-2′-3-dideoxy-5-fluoro-3′- thiacytidine ((-)FTC).

Chapter 4. Conclusion

Reproduced in part with permission from Gadkari, V.V., Tokarsky, E.J., Malik,

C.K., Basu, A.K., and Suo, Z. (2014) Mechanistic Investigation of the Bypass of a Bulky

Aromatic DNA Adduct Catalyzed by a Y-Family DNA Polymerase. DNA Repair. 21, 65-

77. The full article is available at https://doi.org/10.1016/j.dnarep.2014.06.003

Varun V. Gadkari and E. John Tokarsky planned and performed the kinetic experiments.

V.V.G., and Zucai Suo analyzed the results. Modified DNA substrates were provided by

Chachal K. Malik and Ashis K. Basu. V.V.G. wrote the initial draft of the manuscript.

Z.S. conceived the research and modified the manuscript. This work was supported by the National Institutes of Health grant ES009127 to both A.K.B. and Z.S.

Reproduced in part with permission from Vyas, R., Efthimiopoulos, G., Tokarsky,

E.J., Malik, C.K., Basu, A.K., and Suo, Z.* (2015) Mechanistic Basis for the Bypass of a

Bulky DNA Adduct Catalyzed by a Y-Family DNA Polymerase. J. Am. Chem. Soc. 137

(37), 12131-12142. The full article is available at https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4582013/

Rajan Vyas was responsible for crystallizing, data collection and solving crystal structures. Georgia Efthimiopoulos and E. John Tokarsky planned and performed the kinetic experiments. R.V., G.E., and Zucai Suo analyzed the results. Modified DNA substrates were provided by Chachal K. Malik and Ashis K. Basu. R.V., and Z.S. wrote the initial draft of the manuscript. Z.S. conceived the research and modified the

manuscript. This work was supported by a National Institutes of Health grant (ES009127) to Z.S. and A.K.B. and National Science Foundation grant (MCB-0960961) to Z.S. The authors are grateful for the usage of the Advanced Photon Source, an Office of Science

User Facility operated for the U.S. Department of Energy (DOE) Office of Science by

Argonne National Laboratory, which was supported by the U.S. DOE under Contract No.

DE-AC02-06CH11357. The authors are also grateful for the usage of the Lilly Research

Laboratories Collaborative Access Team (LRL-CAT) beamline at Sector 31 of the

Advanced Photon Source provided by Eli Lilly Company, which operates the facility.

Reproduced in part with permission from Patra, A., Politica, D.A., Chatterjee, A.,

Tokarsky, E.J., Suo, Z., Basu, A.K., Stone, M.P., and Egli, M. (2016) Mechanism of

Error-Free Bypass of the Environmental Carcinogen N-(2′- Deoxyguanosin-8-yl)-3- aminobenzanthrone Adduct by Human DNA Polymerase. ChemBioChem. 17(21), 2033-

2037. The full article is available at https://doi.org/10.1002/cbic.201600420

Amritraj Patra, and Dustin A. Politica were responsible for crystallization, data collection, and solving structures. E. John Tokarsky planned and performed the kinetic experiments. Martin Egli, Michael P. Stone, and Zucai Suo analyzed the results.

Modified DNA substrates were provided by Chachal K. Malik and Ashis K. Basu. M.E and M.P.S conceived the research and modified the manuscript. This work was supported by National Institutes of Health (NIH) grants P01 CA-160032 and R01 CA-55678 (M.E.,

M .P.S.), R01 ES05509 (M.P.S), R01 ES-009127 and R01 ES-021762(A.K.B.), and P30

CA-68485 (VICC).

4.1 Introduction

In this dissertation, I have reviewed the literature on DNA replication, DNA damage tolerance and DNA repair as well as presented my own research conducted at during my graduate studies at The Ohio State University. All of the processes mentioned in this dissertation have been studied by many groups and the amount of literature that exists is vastly greater than what I have included here. For more in-depth reviews on eukaryotic DNA replication180 see Burgers et. al. 2017, or DNA replication in general see the book titled “DNA Replication, From Old Principles to New Discoveries” by Masai et. al. For a great review of translesion DNA synthesis181 see Vaisman et. al. 2017, and see

Whitaker et. al. 2017 for a review on base excision repair of oxidative damage51. This chapter is dedicated to highlighting other publications involving mechanistic studies of

DNA polymerases that combine techniques of pre-steady-state kinetics and X-ray crystallography. These are publications that I co-authored, and are closely related to the research presented in Chapters 2 and 3. Additionally, future directions related to Y- family DNA polymerases and PrimPol will be discussed.

4.2 Lesion bypass of dGC8-N-ABA and dG1,8 by Sulfolobus solfataricus Dpo4

Sulfolobus solfataricus (Sso) is a species of thermophilic archaeon that inhabit extreme, high-sulfur conditions such as in hot springs and volcanoes, that are characterized by high temperature (~80°C) and acidic pH (2-4). In contrast to eukaryotes that express four Y-family DNA polymerases, Sso expresses a single Y-family DNA polymerase, Dpo4, that is responsible for all lesion bypass in vivo182. Dpo4 is a model Y-

family polymerase that is homologous to Polκ and E. coli DinB. Dpo4 is a commonly studied DNA polymerase based on its structural stability, ease of protein purification, and ability to bypass many types of DNA damage. Our laboratory has studied Dpo4 for over a decade using methods such as pre-steady-state kinetics, thermodynamics, CD- spectroscopy, NMR, X-ray crystallography, computational modelling, and ensemble and single-molecule based fluorescence techniques111; 129; 142; 172; 183; 184; 185; 186; 187.

We have shown in two published studies that Dpo4 is capable of bypassing the lesions, dGC8-N-ABA and dG1,8, that are caused by common environmental pollutants112; 129. dGC8-N-ABA and dG1,8 are both bulky DNA lesions based on their respective ABA and aminopyrene moieties that modify, and are similar in size to a guanine base. These lesions add approximate molecular weights of 244 g/mol for ABA and 231 g/mol for aminopyrene, which are greater than the MW of guanine (151 g/mol). Although these moieties are differently connected to guanine (C8 for dGC8-N-ABA and the primary amine for dG1,8; Figure 4.1 A-B), they similarly cause Dpo4 to stall during lesion bypass and are eventually traversed based on running start assays (Figure 4.2). Dpo4 demonstrates that upon encountering the lesions, that it stalls significantly during the bypass step for both lesions, and during the extension step for dGC8-N-ABA only. Single-turnover assays indicated that Dpo4 demonstrated a 120- and 1188-fold lower polymerase efficiency

C8-N-ABA (kp/Kd,dNTP), compared to control experiments, during direct bypass of dG and dG1,8, respectively. Similarly, during the extension step, Dpo4 showed a 170- and 13.2- fold lower polymerase efficiency, compared to control experiments. These kinetic data coupled with the running-start assays show that dGC8-N-ABA and dG1,8 are tolerated and

bypassed by Dpo4, but the location of covalent bonding bulky moiety on guanine effects the mechanism of polymerization by Dpo4.

Structural studies are necessary to elucidate the observed differences in polymerase stalling based on different bulky lesions i.e. dGC8-N-ABA is a major-groove lesion and dG1,8 is a minor-groove lesion (Figure 4.1). Unfortunately, the crystal structure of Dpo4 bound to DNA containing dGC8-N-ABA (dGC8-N-ABA -DNA) is not currently solved. However, Kirouac et. al. co-crystallized Dpo4 with DNA containing a similar lesion, N-(deoxyguanosin-8-yl)-1-aminopyrene (dGAP; Figure 4.1C) that revealed insights as to how Dpo4 might also bypass dGC8-N-ABA 121. Accordingly, two protein complexes of Dpo4 in complex with DNA containing dGAP (Dpo4•dGAP-DNA) were detected in a single asymmetric unit and these structures were solved at 2.6 Å resolution.

In the binary structures, the aminopyrene moiety adopts two different conformations; one where it occupies the position of the undamaged template DNA backbone that is packed against a hydrophobic pocket in the Little Finger domain, and another where the aminopyrene ring is flipped out and exposed on one side to solvent and packs the other side against the major groove site of the DNA duplex. In similar studies performed by our laboratory, Dpo4 was co-crystallized with DNA containing a dG1,8 lesion (dG1,8-DNA), dCTP, and Ca2+ 129. Similarly, a single crystal contained two Dpo4 complex molecules per asymmetric unit. Interestingly, one of the Dpo4 complexes was determined as a binary structure with DNA (Dpo4•dG1,8-DNA), and the other a ternary structure with

DNA and dCTP (Dpo4•dG1,8-DNA•dCTP). The binary Dpo4•dG1,8-DNA, revealed that the aminopyrene moiety of the dG1,8 lesion occupied the space where a nascent base pair

would form, thus likely blocking the incoming dCTP from binding. Also, the dG base was repositioned into a cleft that exists between the Finger and Little Finger subdomains of Dpo4. This conformation was stabilized by a hydrogen bond that formed between the

N1 atom of the dG1,8 lesion and the backbone carbonyl oxygen atom of Gly58. The ternary Dpo4•dG1,8-DNA•dCTP structure revealed that in addition to the dG base being looped out into the major groove of the DNA, the immediate downstream base dC (+1 position) was also excluded from the DNA duplex. Surprisingly, the ternary structure shows the incoming dCTP forming a base pair with the downstream dG, +2 positions from the dG1,8 lesion, thus incorporation could potentially lead to a frameshift mutation.

Additionally, the aminopyrene moiety remains within the DNA duplex by forming stacking interactions with the nascent base pair and the junction base pair. Thus the aminopyrene moiety would become sandwiched between two base pairs during lesion bypass by Dpo4.

The crystal structures of Dpo4•dGAP-DNA, Dpo4•dG1,8-DNA, and Dpo4•dG1,8-

DNA•dCTP reveal that these bulky lesions often interfere with incorporation by nesting within the DNA duplex and precluding the incoming nucleotide from binding properly at the insertion site. Dpo4 must actively displace and/or stabilize the bulky moiety outside of the DNA duplex to allow for lesion bypass to occur. The presence of dG1,8 forces

Dpo4 to utilize an interesting mechanism where both the lesion containing, and downstream template bases need to be looped out in order to form a base pair with a downstream dG, thus causing a frame-shift in the DNA strand. This mechanism could be the cause of the greater than 1100-fold decrease in polymerase efficiency observed during

single-turnover assays. Lastly, the adoption of two distinct conformations by the templating dGAP and dG1,8, correlate well with biphasic kinetic assays. These assays indicated that incorporation by Dpo4 opposite these bulky lesions results in a fast phase of incorporation that likely matches a conformation where the bulky lesion is looped out of the DNA duplex, and a slow phase of incorporation where the bulky moiety must be evacuated from the insertion site before incorporation can occur123. A representation of this reaction is presented in Scheme 2.2A. This mechanism is likely followed during the bypass of many bulky DNA lesions catalyzed by Dpo4.

Altogether, the studies carried out by our lab and others, show that Dpo4 is capable of tolerating a wide range of distinct DNA lesions into its active site and must utilize structural rearrangements to achieve lesion bypass. Intriguingly, Dpo4 is tasked with all lesion bypass responsibilities, whereas in eukaryotes, five DNA polymerases

(perhaps more) are utilized to catalyze lesion bypass in vivo. Metaphorically, Dpo4 is a jack of all trades and master of none as it is able to bypass many different types of lesions but perhaps not as efficiently as eukaryotic Y-family DNA polymerases. Eukaryotes have likely adapted to utilize multiple specialized DNA polymerases to allow for maximum efficiency during bypass of many types of DNA damage. This ensures that DNA replication for the much larger eukaryotic genome is maintained at peak efficiency.

However, the greater complexity of having multiple Y-family DNA polymerases, requires greater protein coordination and tighter regulation to ensure high fidelity DNA replication.

4.3 Structural basis of dGC8-N-ABA bypass by hPolη

Around the same time that our pre-steady-state kinetic analysis of human Y- family DNA polymerases bypassing dGC8-N-ABA was conducted, our collaborators Patra and Politica et. al. solved the first ternary crystal structure of hPolη in complex with dGC8-N-ABA-DNA, dCTP and Ca2+120. Similar to the solved crystal structures of Dpo4 in complex with dGAP, and dG1,8, hPolη flipped the ABA moiety out of the DNA duplex and into the major groove. Thus, the incoming dCTP was able to form a Watson-Crick base pair with the adducted guanine base. Interestingly, hPolη forms various contacts with the ABA moiety that stabilizes it within a hydrophobic pocket, and into the major groove of the DNA. Ser62 interacts with the ABA moiety by forming an n→π* interaction with the ABA keto group, as well as establishing additional interactions with the aromatic ring. Trp64 and the immediate downstream template thymine stabilize the aromatic ring on the opposite side of the ABA moiety helping to form the hydrophobic pocket. Interestingly, the Watson-Crick base pair is stabilized by Arg61 and Ser62 that helps lock the templating dGC8-N-ABA base and the incoming dCTP into conformations suitable for incorporation (Figure 4.3).

Notably, multiple conformations of the templating dGC8-N-ABA were not identified in the ternary crystal structure, as was observed with Dpo4 and other bulky lesions (see preceding section). However, an NMR solution structure of a DNA duplex containing dGC8-N-ABA was solved that revealed the ABA moiety intercalated in the duplex and the guanine base flipped into the major groove133 (Figure 4.4). Due to the hydrophobic nature of the ABA moiety, resting within the DNA duplex may constitute the most

thermodynamically stable state. Thus, the action of hPolη stabilizes the ABA moiety within a hydrophobic pocket near its active site and allows the adducted guanine base to rotate into the DNA duplex and make the proper contacts to form a Watson-Crick base pair with an incoming dCTP. This hypothesis is supported by our pre-steady-state kinetic biphasic assays that indicated two phases of nucleotide incorporation for the bypass of dGC8-N-ABA catalyzed by hPolη (See section 2.4.2). Altogether these studies show that

Dpo4 and hPolη may share a common mechanism to cope with bulky DNA lesions such as dGC8-N-ABA, dG1,8, or dGAP, by displacing the bulky moiety from the DNA duplex and into the major groove and allowing the adducted base a chance to form proper contacts with an incoming nucleotide.

4.4 Future directions for Y-family DNA polymerases and PrimPol

For many years the study of Y-family DNA polymerases has been carried out by identifying a DNA lesion and attempting to determine which DNA polymerase(s) are capable of bypassing it and determining the mechanism of lesion bypass. This has normally been carried out by utilizing gel-based kinetics techniques, such as those described in this dissertation, as well as X-ray crystallography. Recently, however, single-molecule fluorescence-based techniques have been utilized to reveal previously unknown mechanistic details. Our laboratory and others have studied DNA polymerases such as Sso Dpo4186, Sso PolB188, HIV Reverse Transcriptase189 by single-molecule

Förster Resonance Energy Transfer (FRET). For example, Förster Resonance Energy

Transfer (FRET) has revealed that during 8-oxoG bypass by Dpo4, the identity of the

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incoming nucleotide dictates whether or not Dpo4 adopts a productive nucleotide binding conformation that is suitable for incorporation. These studies could be expanded to include many different types of DNA lesions that have yet to be analyzed. However, this technique suffers based on the ease of protein-labelling via fluorophores with certain proteins, such as Sso Dpo4. To avoid heterogeneity in the protein population upon labelling, Dpo4 encodes a single cysteine that can be knocked out with little structural consequences, thus allowing the researcher to mutate a different residue to cysteine for labelling with various fluorescent dyes. Unfortunately, many DNA polymerases of interest, such as human Y-family DNA polymerases, contain multiple cysteines that are potentially important for structural stability and therefore cannot be mutated and studied by FRET-based techniques.

Genome sequencing of tumor cells has revealed single nucleotide polymorphisms

(SNPs) in genes that encode for DNA polymerases that may indicate a role in tumorigenesis. For example, Polκ mutants F192C and S423R, have been found in various types of cancers and those mutants have been studied by in vitro methods190.

Some mutations in Polκ make it more active, thus making it harder to regulate and potentially allowing it introduce mutations during lesion bypass. Alternatively, some

Polκ variants such as R298H and Y432S, may lower its lesion bypass abilities, thus allowing other, potentially more error-prone DNA polymerases to bypass these lesions, or leading to replication fork stalling, uncoupling and potential cell death. Additionally, exonuclease domain mutants (EDMs) of Polε have been identified in colorectal cancer

(CRC) and endometrial cancer (EEC) tumors191. Whole genome sequencing of these

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hypermutated tumors revealed >100 mutations per 106 bases, which is much lower than the fidelity of Polε calculated from in vitro studies. Likely, these EDMs contain mutations that impair the ability to proof-read by Polε, via loss of 3’-5’ exonuclease activity. A mechanistic basis of Polε EDMs will reveal how they drive mutagenesis in

CRCs and EECs.

One of the major functions of human PrimPol in vivo is its ability to catalyze replication fork restart. However, as indicated by our studies with PrimPol, it displays low efficiency when performing its activities alone. It has been shown that RPA enhances the activity of PrimPol and also regulates it via protein-protein interactions.

Due to the tight binding of single-stranded DNA by RPA, it likely recruits PrimPol to sites downstream of DNA damage, where single-stranded DNA would be found. Thus, performing kinetic assays for the primase and polymerase activities of PrimPol, in the presence of RPA, would reveal mechanistic details that would be relevant in vivo.

Additionally, affinity purification coupled with mass-spectrometry analysis revealed that

PrimPol interacts with 65 bona fide protein partners in vivo. Only a small subset of these identified interacting partners have been further characterized leaving a gap in the knowledge as to how this protein may be regulated. Further characterizing the interaction of PrimPol with the remaining interacting partners will reveal important details for the true functions of this enzyme.

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4.5 Concluding Remarks

In summary, the work completed in this dissertation describes some of the functions of human Y-family DNA polymerases and PrimPol. The importance of understanding the mechanistic details of the catalytic reactions that these enzymes perform allows us infer their relevance in the human cell and to their roles human health.

Further studies of these enzymes will undoubtedly aid in the understanding of the overall process of DNA replication.

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4.6 Figures

Figure 4.1: Bulky DNA lesions.

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Figure 4.2: Running Start Assays. A preincubated solution of 100 nM Dpo4 and 100 nM 5′-32P-labeled (A and C) 17/26-mer, (B) 17/26-mer-dG1,8, (C) 17/26-mer-dGC8-N-ABA was rapidly mixed with all four dNTPs (200 μM each) for various times before being quenched by addition of EDTA to 0.37 M. Sizes of important products are indicated, and the 21st position marks the position of the dGC8-N-ABA lesion from the 3′-terminus of the

DNA template.

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Figure 4.3: Space filling model of the hPolη active site with dGC8-N-ABA. The lesion dGC8-N-ABA (magenta carbon atoms) is accommodated inside a cleft to the side of the active site that allows for extensive hydrophobic interactions between ABA moiety and surrounding residues from the polymerase finger domain. Carbon atoms of the incoming dCTP are colored in yellow.

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Figure 4.4: NMR structure of the 12-mer DNA duplex containing a C8-dG-ABA adduct. C8-dG-ABA moiety is intercalated between base pairs C4:G21 and T6:A19; the complementary base C20 is displaced into the major groove. Figure adapted from

Politica, 2015133.

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References

1. Bell, S. P., Kobayashi, R. & Stillman, B. (1993). Yeast origin recognition complex functions in transcription silencing and DNA replication. Science 262, 1844-9. 2. Bell, S. P. & Stillman, B. (1992). ATP-dependent recognition of eukaryotic origins of DNA replication by a multiprotein complex. Nature 357, 128-34. 3. Bell, S. P., Mitchell, J., Leber, J., Kobayashi, R. & Stillman, B. (1995). The multidomain structure of Orc1p reveals similarity to regulators of DNA replication and transcriptional silencing. Cell 83, 563-8. 4. Cocker, J. H., Piatti, S., Santocanale, C., Nasmyth, K. & Diffley, J. F. (1996). An essential role for the Cdc6 protein in forming the pre-replicative complexes of budding yeast. Nature 379, 180-2. 5. Rowles, A., Chong, J. P., Brown, L., Howell, M., Evan, G. I. & Blow, J. J. (1996). Interaction between the origin recognition complex and the replication licensing system in Xenopus. Cell 87, 287-96. 6. Nishitani, H., Lygerou, Z., Nishimoto, T. & Nurse, P. (2000). The Cdt1 protein is required to license DNA for replication in fission yeast. Nature 404, 625-8. 7. Romanowski, P., Madine, M. A., Rowles, A., Blow, J. J. & Laskey, R. A. (1996). The Xenopus origin recognition complex is essential for DNA replication and MCM binding to chromatin. Curr Biol 6, 1416-25. 8. Tanaka, T., Knapp, D. & Nasmyth, K. (1997). Loading of an Mcm protein onto DNA replication origins is regulated by Cdc6p and CDKs. Cell 90, 649-60. 9. Aparicio, O. M., Stout, A. M. & Bell, S. P. (1999). Differential assembly of Cdc45p and DNA polymerases at early and late origins of DNA replication. Proc Natl Acad Sci U S A 96, 9130-5. 10. Chong, J. P., Thommes, P. & Blow, J. J. (1996). The role of MCM/P1 proteins in the licensing of DNA replication. Trends Biochem Sci 21, 102-6. 11. Evrin, C., Clarke, P., Zech, J., Lurz, R., Sun, J., Uhle, S., Li, H., Stillman, B. & Speck, C. (2009). A double-hexameric MCM2-7 complex is loaded onto origin DNA during licensing of eukaryotic DNA replication. Proc Natl Acad Sci U S A 106, 20240-5. 12. Kawasaki, Y., Kim, H. D., Kojima, A., Seki, T. & Sugino, A. (2006). Reconstitution of Saccharomyces cerevisiae prereplicative complex assembly in vitro. Genes Cells 11, 745-56. 13. Remus, D., Beuron, F., Tolun, G., Griffith, J. D., Morris, E. P. & Diffley, J. F. (2009). Concerted loading of Mcm2-7 double hexamers around DNA during DNA replication origin licensing. Cell 139, 719-30. 14. Hopwood, B. & Dalton, S. (1996). Cdc45p assembles into a complex with Cdc46p/Mcm5p, is required for minichromosome maintenance, and is essential for chromosomal DNA replication. Proc Natl Acad Sci U S A 93, 12309-14.

108

15. Aparicio, O. M., Weinstein, D. M. & Bell, S. P. (1997). Components and dynamics of DNA replication complexes in S. cerevisiae: redistribution of MCM proteins and Cdc45p during S phase. Cell 91, 59-69. 16. Takayama, Y., Kamimura, Y., Okawa, M., Muramatsu, S., Sugino, A. & Araki, H. (2003). GINS, a novel multiprotein complex required for chromosomal DNA replication in budding yeast. Genes Dev 17, 1153-65. 17. Ilves, I., Petojevic, T., Pesavento, J. J. & Botchan, M. R. (2010). Activation of the MCM2-7 helicase by association with Cdc45 and GINS proteins. Mol Cell 37, 247-58. 18. Costa, A., Ilves, I., Tamberg, N., Petojevic, T., Nogales, E., Botchan, M. R. & Berger, J. M. (2011). The structural basis for MCM2-7 helicase activation by GINS and Cdc45. Nat Struct Mol Biol 18, 471-7. 19. Costa, A., Renault, L., Swuec, P., Petojevic, T., Pesavento, J. J., Ilves, I., MacLellan-Gibson, K., Fleck, R. A., Botchan, M. R. & Berger, J. M. (2014). DNA binding polarity, dimerization, and ATPase ring remodeling in the CMG helicase of the eukaryotic replisome. Elife 3, e03273. 20. Randell, J. C., Fan, A., Chan, C., Francis, L. I., Heller, R. C., Galani, K. & Bell, S. P. (2010). Mec1 is one of multiple kinases that prime the Mcm2-7 helicase for phosphorylation by Cdc7. Mol Cell 40, 353-63. 21. Sheu, Y. J. & Stillman, B. (2010). The Dbf4-Cdc7 kinase promotes S phase by alleviating an inhibitory activity in Mcm4. Nature 463, 113-7. 22. Heller, R. C., Kang, S., Lam, W. M., Chen, S., Chan, C. S. & Bell, S. P. (2011). Eukaryotic origin-dependent DNA replication in vitro reveals sequential action of DDK and S-CDK kinases. Cell 146, 80-91. 23. Kamimura, Y., Tak, Y. S., Sugino, A. & Araki, H. (2001). Sld3, which interacts with Cdc45 (Sld4), functions for chromosomal DNA replication in Saccharomyces cerevisiae. EMBO J 20, 2097-107. 24. Masumoto, H., Muramatsu, S., Kamimura, Y. & Araki, H. (2002). S-Cdk- dependent phosphorylation of Sld2 essential for chromosomal DNA replication in budding yeast. Nature 415, 651-5. 25. Tak, Y. S., Tanaka, Y., Endo, S., Kamimura, Y. & Araki, H. (2006). A CDK- catalysed regulatory phosphorylation for formation of the DNA replication complex Sld2-Dpb11. EMBO J 25, 1987-96. 26. Tanaka, S., Umemori, T., Hirai, K., Muramatsu, S., Kamimura, Y. & Araki, H. (2007). CDK-dependent phosphorylation of Sld2 and Sld3 initiates DNA replication in budding yeast. Nature 445, 328-32. 27. Muramatsu, S., Hirai, K., Tak, Y. S., Kamimura, Y. & Araki, H. (2010). CDK- dependent complex formation between replication proteins Dpb11, Sld2, Pol (epsilon}, and GINS in budding yeast. Genes Dev 24, 602-12. 28. Kanke, M., Kodama, Y., Takahashi, T. S., Nakagawa, T. & Masukata, H. (2012). Mcm10 plays an essential role in origin DNA unwinding after loading of the CMG components. EMBO J 31, 2182-94.

109

29. Nick McElhinny, S. A., Gordenin, D. A., Stith, C. M., Burgers, P. M. & Kunkel, T. A. (2008). Division of labor at the eukaryotic replication fork. Mol Cell 30, 137-44. 30. Campbell, J. L. (1986). Eukaryotic DNA replication. Annu Rev Biochem 55, 733- 71. 31. Lehman, I. R. & Kaguni, L. S. (1989). DNA polymerase alpha. J Biol Chem 264, 4265-8. 32. Baranovskiy, A. G., Zhang, Y., Suwa, Y., Babayeva, N. D., Gu, J., Pavlov, Y. I. & Tahirov, T. H. (2015). Crystal structure of the human primase. J Biol Chem 290, 5635-46. 33. Baranovskiy, A. G., Babayeva, N. D., Zhang, Y., Gu, J., Suwa, Y., Pavlov, Y. I. & Tahirov, T. H. (2016). Mechanism of Concerted RNA-DNA Primer Synthesis by the Human Primosome. J Biol Chem 291, 10006-20. 34. Schauer, G. D. & O'Donnell, M. E. (2017). Quality control mechanisms exclude incorrect polymerases from the eukaryotic replication fork. Proc Natl Acad Sci U S A 114, 675-680. 35. Dieckman, L. M., Johnson, R. E., Prakash, S. & Washington, M. T. (2010). Pre- steady state kinetic studies of the fidelity of nucleotide incorporation by yeast DNA polymerase delta. Biochemistry 49, 7344-50. 36. Zahurancik, W. J., Klein, S. J. & Suo, Z. (2013). Kinetic mechanism of DNA polymerization catalyzed by human DNA polymerase epsilon. Biochemistry 52, 7041-9. 37. Zahurancik, W. J., Klein, S. J. & Suo, Z. (2014). Significant contribution of the 3'-->5' exonuclease activity to the high fidelity of nucleotide incorporation catalyzed by human DNA polymerase. Nucleic Acids Res 42, 13853-60. 38. Pursell, Z. F., Isoz, I., Lundstrom, E. B., Johansson, E. & Kunkel, T. A. (2007). Yeast DNA polymerase epsilon participates in leading-strand DNA replication. Science 317, 127-30. 39. Nick McElhinny, S. A., Kissling, G. E. & Kunkel, T. A. (2010). Differential correction of lagging-strand replication errors made by DNA polymerases {alpha} and {delta}. Proc Natl Acad Sci U S A 107, 21070-5. 40. Yurieva, O. & O'Donnell, M. (2016). Reconstitution of a eukaryotic replisome reveals the mechanism of asymmetric distribution of DNA polymerases. Nucleus 7, 360-8. 41. Langston, L. D., Zhang, D., Yurieva, O., Georgescu, R. E., Finkelstein, J., Yao, N. Y., Indiani, C. & O'Donnell, M. E. (2014). CMG helicase and DNA polymerase epsilon form a functional 15-subunit holoenzyme for eukaryotic leading-strand DNA replication. Proc Natl Acad Sci U S A 111, 15390-5. 42. Sun, J., Yuan, Z., Georgescu, R., Li, H. & O'Donnell, M. (2016). The eukaryotic CMG helicase pumpjack and integration into the replisome. Nucleus 7, 146-54. 43. Yuan, Z., Bai, L., Sun, J., Georgescu, R., Liu, J., O'Donnell, M. E. & Li, H. (2016). Structure of the eukaryotic replicative CMG helicase suggests a pumpjack motion for translocation. Nat Struct Mol Biol 23, 217-24.

110

44. Okazaki, R., Okazaki, T., Sakabe, K., Sugimoto, K. & Sugino, A. (1968). Mechanism of DNA chain growth. I. Possible discontinuity and unusual secondary structure of newly synthesized chains. Proc Natl Acad Sci U S A 59, 598-605. 45. Wold, M. S. (1997). Replication protein A: a heterotrimeric, single-stranded DNA-binding protein required for eukaryotic DNA metabolism. Annu Rev Biochem 66, 61-92. 46. Greinert, R., Volkmer, B., Henning, S., Breitbart, E. W., Greulich, K. O., Cardoso, M. C. & Rapp, A. (2012). UVA-induced DNA double-strand breaks result from the repair of clustered oxidative DNA damages. Nucleic Acids Res 40, 10263-73. 47. Stodola, J. L. & Burgers, P. M. (2016). Resolving individual steps of Okazaki- fragment maturation at a millisecond timescale. Nat Struct Mol Biol 23, 402-8. 48. Howes, T. R. & Tomkinson, A. E. (2012). DNA ligase I, the replicative DNA ligase. Subcell Biochem 62, 327-41. 49. Prakash, S., Johnson, R. E. & Prakash, L. (2005). Eukaryotic translesion synthesis DNA polymerases: specificity of structure and function. Annu Rev Biochem 74, 317-53. 50. Petermann, E. & Helleday, T. (2010). Pathways of mammalian replication fork restart. Nat Rev Mol Cell Biol 11, 683-7. 51. Whitaker, A. M., Schaich, M. A., Smith, M. R., Flynn, T. S. & Freudenthal, B. D. (2017). Base excision repair of oxidative DNA damage: from mechanism to disease. Front Biosci (Landmark Ed) 22, 1493-1522. 52. Bebenek, K. & Kunkel, T. A. (2004). Functions of DNA polymerases. Adv Protein Chem 69, 137-65. 53. Cortez, D. (2015). Preventing replication fork collapse to maintain genome integrity. DNA Repair (Amst) 32, 149-57. 54. Lehmann, A. R., Kirk-Bell, S., Arlett, C. F., Paterson, M. C., Lohman, P. H., de Weerd-Kastelein, E. A. & Bootsma, D. (1975). Xeroderma pigmentosum cells with normal levels of excision repair have a defect in DNA synthesis after UV- irradiation. Proc Natl Acad Sci U S A 72, 219-23. 55. Johnson, R. E., Prakash, S. & Prakash, L. (1999). Efficient bypass of a thymine- thymine dimer by yeast DNA polymerase, Poleta. Science 283, 1001-4. 56. Haracska, L., Prakash, S. & Prakash, L. (2000). Replication past O(6)- methylguanine by yeast and human DNA polymerase eta. Mol Cell Biol 20, 8001- 7. 57. Haracska, L., Yu, S. L., Johnson, R. E., Prakash, L. & Prakash, S. (2000). Efficient and accurate replication in the presence of 7,8-dihydro-8-oxoguanine by DNA polymerase eta. Nat Genet 25, 458-61. 58. Sherrer, S. M., Sanman, L. E., Xia, C. X., Bolin, E. R., Malik, C. K., Efthimiopoulos, G., Basu, A. K. & Suo, Z. (2012). Kinetic analysis of the bypass of a bulky DNA lesion catalyzed by human Y-family DNA polymerases. Chem Res Toxicol 25, 730-40.

111

59. Tokarsky, E. J., Gadkari, V. V., Zahurancik, W. J., Malik, C. K., Basu, A. K. & Suo, Z. (2016). Pre-steady-state kinetic investigation of bypass of a bulky guanine lesion by human Y-family DNA polymerases. DNA Repair (Amst) 46, 20-28. 60. Carlson, K. D., Johnson, R. E., Prakash, L., Prakash, S. & Washington, M. T. (2006). Human DNA polymerase kappa forms nonproductive complexes with matched primer termini but not with mismatched primer termini. Proc Natl Acad Sci U S A 103, 15776-81. 61. Washington, M. T., Johnson, R. E., Prakash, L. & Prakash, S. (2002). Human DINB1-encoded DNA polymerase kappa is a promiscuous extender of mispaired primer termini. Proc Natl Acad Sci U S A 99, 1910-4. 62. Lone, S., Townson, S. A., Uljon, S. N., Johnson, R. E., Brahma, A., Nair, D. T., Prakash, S., Prakash, L. & Aggarwal, A. K. (2007). Human DNA polymerase kappa encircles DNA: implications for mismatch extension and lesion bypass. Mol Cell 25, 601-14. 63. Haracska, L., Prakash, L. & Prakash, S. (2002). Role of human DNA polymerase kappa as an extender in translesion synthesis. Proc Natl Acad Sci U S A 99, 16000-5. 64. Jha, V. & Ling, H. (2018). Structural Basis for Human DNA Polymerase Kappa to Bypass Cisplatin Intrastrand Cross-Link (Pt-GG) Lesion as an Efficient and Accurate Extender. J Mol Biol 430, 1577-1589. 65. Song, I., Kim, E. J., Kim, I. H., Park, E. M., Lee, K. E., Shin, J. H., Guengerich, F. P. & Choi, J. Y. (2014). Biochemical characterization of eight genetic variants of human DNA polymerase kappa involved in error-free bypass across bulky N(2)-guanyl DNA adducts. Chem Res Toxicol 27, 919-30. 66. Ogi, T. & Lehmann, A. R. (2006). The Y-family DNA polymerase kappa (pol kappa) functions in mammalian nucleotide-excision repair. Nat Cell Biol 8, 640-2. 67. Haracska, L., Johnson, R. E., Unk, I., Phillips, B. B., Hurwitz, J., Prakash, L. & Prakash, S. (2001). Targeting of human DNA polymerase iota to the replication machinery via interaction with PCNA. Proc Natl Acad Sci U S A 98, 14256-61. 68. Washington, M. T., Johnson, R. E., Prakash, L. & Prakash, S. (2004). Human DNA polymerase iota utilizes different nucleotide incorporation mechanisms dependent upon the template base. Mol Cell Biol 24, 936-43. 69. Nair, D. T., Johnson, R. E., Prakash, S., Prakash, L. & Aggarwal, A. K. (2004). Replication by human DNA polymerase-iota occurs by Hoogsteen base-pairing. Nature 430, 377-80. 70. Washington, M. T., Minko, I. G., Johnson, R. E., Haracska, L., Harris, T. M., Lloyd, R. S., Prakash, S. & Prakash, L. (2004). Efficient and error-free replication past a minor-groove N2-guanine adduct by the sequential action of yeast Rev1 and DNA polymerase zeta. Mol Cell Biol 24, 6900-6. 71. Swan, M. K., Johnson, R. E., Prakash, L., Prakash, S. & Aggarwal, A. K. (2009). Structure of the human Rev1-DNA-dNTP ternary complex. J Mol Biol 390, 699- 709.

112

72. Brown, J. A., Fowler, J. D. & Suo, Z. (2010). Kinetic basis of nucleotide selection employed by a protein template-dependent DNA polymerase. Biochemistry 49, 5504-10. 73. Masuda, Y., Ohmae, M., Masuda, K. & Kamiya, K. (2003). Structure and enzymatic properties of a stable complex of the human REV1 and REV7 proteins. J Biol Chem 278, 12356-60. 74. Guo, C., Fischhaber, P. L., Luk-Paszyc, M. J., Masuda, Y., Zhou, J., Kamiya, K., Kisker, C. & Friedberg, E. C. (2003). Mouse Rev1 protein interacts with multiple DNA polymerases involved in translesion DNA synthesis. EMBO J 22, 6621-30. 75. Mukhopadhyay, S., Clark, D. R., Watson, N. B., Zacharias, W. & McGregor, W. G. (2004). REV1 accumulates in DNA damage-induced nuclear foci in human cells and is implicated in mutagenesis by benzo[a]pyrenediolepoxide. Nucleic Acids Res 32, 5820-6. 76. Shachar, S., Ziv, O., Avkin, S., Adar, S., Wittschieben, J., Reissner, T., Chaney, S., Friedberg, E. C., Wang, Z., Carell, T., Geacintov, N. & Livneh, Z. (2009). Two-polymerase mechanisms dictate error-free and error-prone translesion DNA synthesis in mammals. EMBO J 28, 383-93. 77. Johnson, R. E., Washington, M. T., Haracska, L., Prakash, S. & Prakash, L. (2000). Eukaryotic polymerases iota and zeta act sequentially to bypass DNA lesions. Nature 406, 1015-9. 78. Johnson, R. E., Yu, S. L., Prakash, S. & Prakash, L. (2003). Yeast DNA polymerase zeta (zeta) is essential for error-free replication past thymine glycol. Genes Dev 17, 77-87. 79. Gomez-Llorente, Y., Malik, R., Jain, R., Choudhury, J. R., Johnson, R. E., Prakash, L., Prakash, S., Ubarretxena-Belandia, I. & Aggarwal, A. K. (2013). The architecture of yeast DNA polymerase zeta. Cell Rep 5, 79-86. 80. Iyer, L. M., Koonin, E. V., Leipe, D. D. & Aravind, L. (2005). Origin and evolution of the archaeo-eukaryotic primase superfamily and related palm-domain proteins: structural insights and new members. Nucleic Acids Res 33, 3875-96. 81. Guilliam, T. A., Keen, B. A., Brissett, N. C. & Doherty, A. J. (2015). Primase- polymerases are a functionally diverse superfamily of replication and repair enzymes. Nucleic Acids Res 43, 6651-64. 82. Garcia-Gomez, S., Reyes, A., Martinez-Jimenez, M. I., Chocron, E. S., Mouron, S., Terrados, G., Powell, C., Salido, E., Mendez, J., Holt, I. J. & Blanco, L. (2013). PrimPol, an archaic primase/polymerase operating in human cells. Mol Cell 52, 541-53. 83. Wan, L., Lou, J., Xia, Y., Su, B., Liu, T., Cui, J., Sun, Y., Lou, H. & Huang, J. (2013). hPrimpol1/CCDC111 is a human DNA primase-polymerase required for the maintenance of genome integrity. EMBO Rep 14, 1104-12. 84. Keen, B. A., Jozwiakowski, S. K., Bailey, L. J., Bianchi, J. & Doherty, A. J. (2014). Molecular dissection of the domain architecture and catalytic activities of human PrimPol. Nucleic Acids Res 42, 5830-45. 85. Bianchi, J., Rudd, S. G., Jozwiakowski, S. K., Bailey, L. J., Soura, V., Taylor, E., Stevanovic, I., Green, A. J., Stracker, T. H., Lindsay, H. D. & Doherty, A. J. 113

(2013). PrimPol bypasses UV photoproducts during eukaryotic chromosomal DNA replication. Mol Cell 52, 566-73. 86. Mouron, S., Rodriguez-Acebes, S., Martinez-Jimenez, M. I., Garcia-Gomez, S., Chocron, S., Blanco, L. & Mendez, J. (2013). Repriming of DNA synthesis at stalled replication forks by human PrimPol. Nat Struct Mol Biol 20, 1383-9. 87. Guilliam, T. A., Jozwiakowski, S. K., Ehlinger, A., Barnes, R. P., Rudd, S. G., Bailey, L. J., Skehel, J. M., Eckert, K. A., Chazin, W. J. & Doherty, A. J. (2015). Human PrimPol is a highly error-prone polymerase regulated by single-stranded DNA binding proteins. Nucleic Acids Res 43, 1056-68. 88. Schiavone, D., Jozwiakowski, S. K., Romanello, M., Guilbaud, G., Guilliam, T. A., Bailey, L. J., Sale, J. E. & Doherty, A. J. (2016). PrimPol Is Required for Replicative Tolerance of G Quadruplexes in Vertebrate Cells. Mol Cell 61, 161-9. 89. Kobayashi, K., Guilliam, T. A., Tsuda, M., Yamamoto, J., Bailey, L. J., Iwai, S., Takeda, S., Doherty, A. J. & Hirota, K. (2016). Repriming by PrimPol is critical for DNA replication restart downstream of lesions and chain-terminating nucleosides. Cell Cycle, 0. 90. Lindahl, T. & Nyberg, B. (1972). Rate of depurination of native deoxyribonucleic acid. Biochemistry 11, 3610-8. 91. Cadet, J. & Wagner, J. R. (2013). DNA base damage by reactive oxygen species, oxidizing agents, and UV radiation. Cold Spring Harb Perspect Biol 5. 92. Freudenthal, B. D., Beard, W. A., Perera, L., Shock, D. D., Kim, T., Schlick, T. & Wilson, S. H. (2015). Uncovering the polymerase-induced cytotoxicity of an oxidized nucleotide. Nature 517, 635-9. 93. Vyas, R., Reed, A. J., Tokarsky, E. J. & Suo, Z. (2015). Viewing Human DNA Polymerase beta Faithfully and Unfaithfully Bypass an Oxidative Lesion by Time-Dependent Crystallography. J Am Chem Soc 137, 5225-30. 94. Wilson, S. H. & Kunkel, T. A. (2000). Passing the baton in base excision repair. Nat Struct Biol 7, 176-8. 95. Denver, D. R., Swenson, S. L. & Lynch, M. (2003). An evolutionary analysis of the helix-hairpin-helix superfamily of DNA repair glycosylases. Mol Biol Evol 20, 1603-11. 96. Batebi, H., Dragelj, J. & Imhof, P. (2018). Role of AP-endonuclease (Ape1) active site residues in stabilization of the reactant enzyme-DNA complex. Proteins. 97. Mol, C. D., Izumi, T., Mitra, S. & Tainer, J. A. (2000). DNA-bound structures and mutants reveal abasic DNA binding by APE1 and DNA repair coordination [corrected]. Nature 403, 451-6. 98. Freudenthal, B. D., Beard, W. A., Cuneo, M. J., Dyrkheeva, N. S. & Wilson, S. H. (2015). Capturing snapshots of APE1 processing DNA damage. Nat Struct Mol Biol 22, 924-31. 99. Sawaya, M. R., Prasad, R., Wilson, S. H., Kraut, J. & Pelletier, H. (1997). Crystal structures of human DNA polymerase beta complexed with gapped and nicked DNA: evidence for an induced fit mechanism. Biochemistry 36, 11205-15.

114

100. Taylor, R. M., Wickstead, B., Cronin, S. & Caldecott, K. W. (1998). Role of a BRCT domain in the interaction of DNA ligase III-alpha with the DNA repair protein XRCC1. Curr Biol 8, 877-80. 101. Arlt, V. M. (2005). 3-Nitrobenzanthrone, a potential human cancer hazard in diesel exhaust and urban air pollution: a review of the evidence. Mutagenesis 20, 399-410. 102. Arlt, V. M., Zhan, L., Schmeiser, H. H., Honma, M., Hayashi, M., Phillips, D. H. & Suzuki, T. (2004). DNA adducts and mutagenic specificity of the ubiquitous environmental pollutant 3-nitrobenzanthrone in Muta Mouse. Environ Mol Mutagen 43, 186-95. 103. Beckman, R. A., Mildvan, A. S. & Loeb, L. A. (1985). On the fidelity of DNA replication: manganese mutagenesis in vitro. Biochemistry 24, 5810-7. 104. Brautigam, C. A. & Steitz, T. A. (1998). Structural and functional insights provided by crystal structures of DNA polymerases and their substrate complexes. Curr Opin Struct Biol 8, 54-63. 105. Johnson, R. E., Prakash, L. & Prakash, S. (2005). Biochemical evidence for the requirement of Hoogsteen base pairing for replication by human DNA polymerase iota. Proc Natl Acad Sci U S A 102, 10466-71. 106. Taga, R., Tang, N., Hattori, T., Tamura, K., Sakai, S., Toriba, A., Kizu, R. & Hayakawa, K. (2005). Direct-acting mutagenicity of extracts of coal burning- derived particulates and contribution of nitropolycyclic aromatic hydrocarbons. Mutat Res 581, 91-5. 107. vom Brocke, J., Krais, A., Whibley, C., Hollstein, M. C. & Schmeiser, H. H. (2009). The carcinogenic air pollutant 3-nitrobenzanthrone induces GC to TA transversion mutations in human p53 sequences. Mutagenesis 24, 17-23. 108. Arlt, V. M., Phillips, D. H. & Reynisson, J. (2011). Theoretical investigations on the formation of nitrobenzanthrone-DNA adducts. Org Biomol Chem 9, 6100-10. 109. Kawanishi, M., Fujikawa, Y., Ishii, H., Nishida, H., Higashigaki, Y., Kanno, T., Matsuda, T., Takamura-Enya, T. & Yagi, T. (2013). Adduct formation and repair, and translesion DNA synthesis across the adducts in human cells exposed to 3- nitrobenzanthrone. Mutat Res 753, 93-100. 110. Kunkel, T. A. (2004). DNA replication fidelity. J Biol Chem 279, 16895-8. 111. Maxwell, B. A. & Suo, Z. (2012). Kinetic Basis for the Differing Response to an Oxidative Lesion by a Replicative and a Lesion Bypass DNA Polymerase from Solfolobus solfataricus. Biochemistry 51, 3485-96. 112. Gadkari, V. V., Tokarsky, E. J., Malik, C. K., Basu, A. K. & Suo, Z. (2014). Mechanistic investigation of the bypass of a bulky aromatic DNA adduct catalyzed by a Y-family DNA polymerase. DNA Repair (Amst) 21, 65-77. 113. Sherrer, S. M., Fiala, K. A., Fowler, J. D., Newmister, S. A., Pryor, J. M. & Suo, Z. (2011). Quantitative analysis of the efficiency and mutagenic spectra of abasic lesion bypass catalyzed by human Y-family DNA polymerases. Nucleic Acids Res 39, 609-22.

115

114. Taggart, D. J., Fredrickson, S. W., Gadkari, V. V. & Suo, Z. (2014). Mutagenic potential of 8-oxo-7,8-dihydro-2'-deoxyguanosine bypass catalyzed by human Y- family DNA polymerases. Chem Res Toxicol 27, 931-40. 115. Wolfle, W. T., Washington, M. T., Prakash, L. & Prakash, S. (2003). Human DNA polymerase kappa uses template-primer misalignment as a novel means for extending mispaired termini and for generating single-base deletions. Genes Dev 17, 2191-9. 116. Zhang, Y., Yuan, F., Xin, H., Wu, X., Rajpal, D. K., Yang, D. & Wang, Z. (2000). Human DNA polymerase kappa synthesizes DNA with extraordinarily low fidelity. Nucleic Acids Res 28, 4147-56. 117. Fiala, K. A., Brown, J. A., Ling, H., Kshetry, A. K., Zhang, J., Taylor, J. S., Yang, W. & Suo, Z. (2007). Mechanism of Template-independent Nucleotide Incorporation Catalyzed by a Template-dependent DNA Polymerase. J Mol Biol 365, 590-602. 118. Choi, J. Y., Zang, H., Angel, K. C., Kozekov, I. D., Goodenough, A. K., Rizzo, C. J. & Guengerich, F. P. (2006). Translesion synthesis across 1,N2-ethenoguanine by human DNA polymerases. Chem Res Toxicol 19, 879-86. 119. Pande, P., Malik, C. K., Bose, A., Jasti, V. P. & Basu, A. K. (2014). Mutational analysis of the C8-guanine adduct of the environmental carcinogen 3- nitrobenzanthrone in human cells: critical roles of DNA polymerases eta and kappa and Rev1 in error-prone translesion synthesis. Biochemistry 53, 5323-31. 120. Patra, A., Politica, D. A., Chatterjee, A., Tokarsky, E. J., Suo, Z., Basu, A., Stone, M. P. & Egli, M. (2016). Mechanism of Error-Free Bypass of the Environmental Carcinogen N-(2-Deoxyguanosin-8-yl)-3-aminobenzanthrone Adduct by Human DNA Polymerase. Chembiochem. 121. Kirouac, K. N., Basu, A. K. & Ling, H. (2013). Structural mechanism of replication stalling on a bulky amino-polycyclic aromatic hydrocarbon DNA adduct by a y family DNA polymerase. J Mol Biol 425, 4167-76. 122. Bauer, J., Xing, G., Yagi, H., Sayer, J. M., Jerina, D. M. & Ling, H. (2007). A structural gap in Dpo4 supports mutagenic bypass of a major benzo[a]pyrene dG adduct in DNA through template misalignment. Proc Natl Acad Sci U S A 104, 14905-10. 123. Sherrer, S. M., Brown, J. A., Pack, L. R., Jasti, V. P., Fowler, J. D., Basu, A. K. & Suo, Z. (2009). Mechanistic studies of the bypass of a bulky single-base lesion catalyzed by a Y-family DNA polymerase. J Biol Chem 284, 6379-88. 124. Fiala, K. A., Hypes, C. D. & Suo, Z. (2007). Mechanism of abasic lesion bypass catalyzed by a Y-family DNA polymerase. J Biol Chem 282, 8188-98. 125. Suarez, S. C., Toffton, S. M. & McCulloch, S. D. (2014). Biochemical analysis of DNA polymerase eta fidelity in the presence of replication protein A. PLoS One 9, e97382. 126. McCulloch, S. D., Kokoska, R. J., Masutani, C., Iwai, S., Hanaoka, F. & Kunkel, T. A. (2004). Preferential cis-syn thymine dimer bypass by DNA polymerase eta occurs with biased fidelity. Nature 428, 97-100.

116

127. Kawanishi, M., Enya, T., Suzuki, H., Takebe, H., Matsui, S. & Yagi, T. (1998). Mutagenic specificity of a derivative of 3-nitrobenzanthrone in the supF shuttle vector plasmids. Chem Res Toxicol 11, 1468-73. 128. Brown, J. A., Newmister, S. A., Fiala, K. A. & Suo, Z. (2008). Mechanism of double-base lesion bypass catalyzed by a Y-family DNA polymerase. Nucleic Acids Res 36, 3867-78. 129. Vyas, R., Efthimiopoulos, G., Tokarsky, E. J., Malik, C. K., Basu, A. K. & Suo, Z. (2015). Mechanistic Basis for the Bypass of a Bulky DNA Adduct Catalyzed by a Y-Family DNA Polymerase. J Am Chem Soc 137, 12131-42. 130. Ohashi, E., Murakumo, Y., Kanjo, N., Akagi, J., Masutani, C., Hanaoka, F. & Ohmori, H. (2004). Interaction of hREV1 with three human Y-family DNA polymerases. Genes Cells 9, 523-31. 131. Sherrer, S. M., Taggart, D. J., Pack, L. R., Malik, C. K., Basu, A. K. & Suo, Z. (2012). Quantitative analysis of the mutagenic potential of 1-aminopyrene-DNA adduct bypass catalyzed by Y-family DNA polymerases. Mutat Res 737, 25-33. 132. Tissier, A., Kannouche, P., Reck, M. P., Lehmann, A. R., Fuchs, R. P. & Cordonnier, A. (2004). Co-localization in replication foci and interaction of human Y-family members, DNA polymerase pol eta and REVl protein. DNA Repair (Amst) 3, 1503-14. 133. Politica, D. A., Malik, C. K., Basu, A. K. & Stone, M. P. (2015). Base-Displaced Intercalated Structure of the N-(2'-Deoxyguanosin-8-yl)-3-aminobenzanthrone DNA Adduct. Chem Res Toxicol 28, 2253-66. 134. Schorr, S., Schneider, S., Lammens, K., Hopfner, K. P. & Carell, T. (2010). Mechanism of replication blocking and bypass of Y-family polymerase {eta} by bulky acetylaminofluorene DNA adducts. Proc Natl Acad Sci U S A 107, 20720- 5. 135. Zafar, M. K., Ketkar, A., Lodeiro, M. F., Cameron, C. E. & Eoff, R. L. (2014). Kinetic analysis of human PrimPol DNA polymerase activity reveals a generally error-prone enzyme capable of accurately bypassing 7,8-dihydro-8-oxo-2'- deoxyguanosine. Biochemistry 53, 6584-94. 136. Balajee, A. S. & Geard, C. R. (2004). Replication protein A and gamma-H2AX foci assembly is triggered by cellular response to DNA double-strand breaks. Exp Cell Res 300, 320-34. 137. Lee, H. R. & Johnson, K. A. (2006). Fidelity of the human mitochondrial DNA polymerase. J Biol Chem 281, 36236-40. 138. Graves, S. W., Johnson, A. A. & Johnson, K. A. (1998). Expression, purification, and initial kinetic characterization of the large subunit of the human mitochondrial DNA polymerase. Biochemistry 37, 6050-8. 139. Martinez-Jimenez, M. I., Garcia-Gomez, S., Bebenek, K., Sastre-Moreno, G., Calvo, P. A., Diaz-Talavera, A., Kunkel, T. A. & Blanco, L. (2015). Alternative solutions and new scenarios for translesion DNA synthesis by human PrimPol. DNA Repair (Amst) 29, 127-38. 140. Berg, J. M. (1990). Zinc fingers and other metal-binding domains. Elements for interactions between macromolecules. J Biol Chem 265, 6513-6. 117

141. Thiesen, H. J. & Bach, C. (1991). Transition metals modulate DNA-protein interactions of SP1 zinc finger domains with its cognate target site. Biochem Biophys Res Commun 176, 551-7. 142. Fiala, K. A. & Suo, Z. (2004). Pre-Steady-State Kinetic Studies of the Fidelity of Sulfolobus solfataricus P2 DNA Polymerase IV. Biochemistry 43, 2106-15. 143. Zhang, L., Brown, J. A., Newmister, S. A. & Suo, Z. (2009). Polymerization fidelity of a replicative DNA polymerase from the hyperthermophilic archaeon Sulfolobus solfataricus P2. Biochemistry 48, 7492-501. 144. Fiala, K. A., Abdel-Gawad, W. & Suo, Z. (2004). Pre-Steady-State Kinetic Studies of the Fidelity and Mechanism of Polymerization Catalyzed by Truncated Human DNA Polymerase lambda. Biochemistry 43, 6751-62. 145. Roettger, M. P., Fiala, K. A., Sompalli, S., Dong, Y. & Suo, Z. (2004). Pre- steady-state kinetic studies of the fidelity of human DNA polymerase mu. Biochemistry 43, 13827-38. 146. Johnson, K. A. (1995). Rapid quench kinetic analysis of polymerases, adenosinetriphosphatases, and enzyme intermediates. Methods Enzymol 249, 38- 61. 147. Georgescu, R. E., Schauer, G. D., Yao, N. Y., Langston, L. D., Yurieva, O., Zhang, D., Finkelstein, J. & O'Donnell, M. E. (2015). Reconstitution of a eukaryotic replisome reveals suppression mechanisms that define leading/lagging strand operation. Elife 4, e04988. 148. Zerbe, L. K. & Kuchta, R. D. (2002). The p58 subunit of human DNA primase is important for primer initiation, elongation, and counting. Biochemistry 41, 4891- 900. 149. Brown, J. A., Fiala, K. A., Fowler, J. D., Sherrer, S. M., Newmister, S. A., Duym, W. W. & Suo, Z. (2010). A novel mechanism of sugar selection utilized by a human X-family DNA polymerase. J Mol Biol 395, 282-90. 150. Astatke, M., Ng, K., Grindley, N. D. & Joyce, C. M. (1998). A single side chain prevents Escherichia coli DNA polymerase I (Klenow fragment) from incorporating ribonucleotides. Proc Natl Acad Sci U S A 95, 3402-7. 151. Yang, G., Franklin, M., Li, J., Lin, T. C. & Konigsberg, W. (2002). A conserved Tyr residue is required for sugar selectivity in a Pol alpha DNA polymerase. Biochemistry 41, 10256-61. 152. Nick McElhinny, S. A. & Ramsden, D. A. (2003). Polymerase mu is a DNA- directed DNA/RNA polymerase. Mol Cell Biol 23, 2309-15. 153. Boule, J. B., Rougeon, F. & Papanicolaou, C. (2001). Terminal deoxynucleotidyl transferase indiscriminately incorporates ribonucleotides and deoxyribonucleotides. J Biol Chem 276, 31388-93. 154. Burris, H. A., 3rd, Moore, M. J., Andersen, J., Green, M. R., Rothenberg, M. L., Modiano, M. R., Cripps, M. C., Portenoy, R. K., Storniolo, A. M., Tarassoff, P., Nelson, R., Dorr, F. A., Stephens, C. D. & Von Hoff, D. D. (1997). Improvements in survival and clinical benefit with gemcitabine as first-line therapy for patients with advanced pancreas cancer: a randomized trial. J Clin Oncol 15, 2403-13.

118

155. Shelley, M. D., Jones, G., Cleves, A., Wilt, T. J., Mason, M. D. & Kynaston, H. G. (2012). Intravesical gemcitabine therapy for non-muscle invasive bladder cancer (NMIBC): a systematic review. BJU Int 109, 496-505. 156. Kroep, J. R., Giaccone, G., Tolis, C., Voorn, D. A., Loves, W. J., Groeningen, C. J., Pinedo, H. M. & Peters, G. J. (2000). Sequence dependent effect of paclitaxel on gemcitabine metabolism in relation to cell cycle and cytotoxicity in non-small- cell lung cancer cell lines. Br J Cancer 83, 1069-76. 157. Fowler, J. D., Brown, J. A., Johnson, K. A. & Suo, Z. (2008). Kinetic investigation of the inhibitory effect of gemcitabine on DNA polymerization catalyzed by human mitochondrial DNA polymerase. J Biol Chem 283, 15339-48. 158. Liu Yin, J. A., Johnson, P. R., Davies, J. M., Flanagan, N. G., Gorst, D. W. & Lewis, M. J. (1991). Mitozantrone and cytosine arabinoside as first-line therapy in elderly patients with acute myeloid leukaemia. Br J Haematol 79, 415-20. 159. Brown, J. A., Pack, L. R., Fowler, J. D. & Suo, Z. (2011). Pre-steady-state kinetic analysis of the incorporation of anti-HIV nucleotide analogs catalyzed by human X- and Y-family DNA polymerases. Antimicrob Agents Chemother 55, 276-83. 160. Brown, J. A., Pack, L. R., Fowler, J. D. & Suo, Z. (2012). Presteady state kinetic investigation of the incorporation of anti-hepatitis B nucleotide analogues catalyzed by noncanonical human DNA polymerases. Chem Res Toxicol 25, 225- 33. 161. Vyas, R., Zahurancik, W. J. & Suo, Z. (2014). Structural basis for the binding and incorporation of nucleotide analogs with L-stereochemistry by human DNA polymerase lambda. Proc Natl Acad Sci U S A 111, E3033-42. 162. Guilliam, T. A., Bailey, L. J., Brissett, N. C. & Doherty, A. J. (2016). PolDIP2 interacts with human PrimPol and enhances its DNA polymerase activities. Nucleic Acids Res 44, 3317-29. 163. Mislak, A. C. & Anderson, K. S. (2016). Insights into the Molecular Mechanism of Polymerization and Nucleoside Reverse Transcriptase Inhibitor Incorporation by Human PrimPol. Antimicrob Agents Chemother 60, 561-9. 164. Maxwell, B. A. & Suo, Z. (2014). Recent insight into the kinetic mechanisms and conformational dynamics of Y-Family DNA polymerases. Biochemistry 53, 2804- 14. 165. Pelletier, H., Sawaya, M. R., Wolfle, W., Wilson, S. H. & Kraut, J. (1996). A structural basis for metal ion mutagenicity and nucleotide selectivity in human DNA polymerase beta. Biochemistry 35, 12762-77. 166. El-Deiry, W. S., Downey, K. M. & So, A. G. (1984). Molecular mechanisms of manganese mutagenesis. Proc Natl Acad Sci U S A 81, 7378-82. 167. Goodman, M. F., Keener, S., Guidotti, S. & Branscomb, E. W. (1983). On the enzymatic basis for mutagenesis by manganese. J Biol Chem 258, 3469-75. 168. Loeb, L. A. & Kunkel, T. A. (1982). Fidelity of DNA synthesis. Annu Rev Biochem 51, 429-57. 169. Richardson, F. C., Kuchta, R. D., Mazurkiewicz, A. & Richardson, K. A. (2000). Polymerization of 2'-fluoro- and 2'-O-methyl-dNTPs by human DNA polymerase alpha, polymerase gamma, and primase. Biochem Pharmacol 59, 1045-52. 119

170. Brown, J. A. & Suo, Z. (2011). Unlocking the sugar "steric gate" of DNA polymerases. Biochemistry 50, 1135-42. 171. Rechkoblit, O., Gupta, Y. K., Malik, R., Rajashankar, K. R., Johnson, R. E., Prakash, L., Prakash, S. & Aggarwal, A. K. (2016). Structure and mechanism of human PrimPol, a DNA polymerase with primase activity. Sci Adv 2, e1601317. 172. Sherrer, S. M., Beyer, D. C., Xia, C. X., Fowler, J. D. & Suo, Z. (2010). Kinetic Basis of Sugar Selection by a Y-Family DNA Polymerase from Sulfolobus solfataricus P2. Biochemistry 49, 10179-10186. 173. Kirouac, K. N., Suo, Z. & Ling, H. (2011). Structural mechanism of ribonucleotide discrimination by a Y-family DNA polymerase. J Mol Biol 407, 382-90. 174. Kobayashi, K., Guilliam, T. A., Tsuda, M., Yamamoto, J., Bailey, L. J., Iwai, S., Takeda, S., Doherty, A. J. & Hirota, K. (2016). Repriming by PrimPol is critical for DNA replication restart downstream of lesions and chain-terminating nucleosides. Cell Cycle 15, 1997-2008. 175. Gaur, V., Vyas, R., Fowler, J. D., Efthimiopoulos, G., Feng, J. Y. & Suo, Z. (2014). Structural and kinetic insights into binding and incorporation of L- nucleotide analogs by a Y-family DNA polymerase. Nucleic Acids Res 42, 9984- 95. 176. Tsoi, P. Y. & Yang, M. (2002). Kinetic study of various binding modes between human DNA polymerase beta and different DNA substrates by surface-plasmon- resonance biosensor. Biochem J 361, 317-25. 177. Brown, J. A., Pack, L. R., Sherrer, S. M., Kshetry, A. K., Newmister, S. A., Fowler, J. D., Taylor, J. S. & Suo, Z. (2010). Identification of critical residues for the tight binding of both correct and incorrect nucleotides to human DNA polymerase lambda. J Mol Biol 403, 505-15. 178. Martin, M. J., Juarez, R. & Blanco, L. (2012). DNA-binding determinants promoting NHEJ by human Polmu. Nucleic Acids Res 40, 11389-403. 179. Washington, M. T., Minko, I. G., Johnson, R. E., Wolfle, W. T., Harris, T. M., Lloyd, R. S., Prakash, S. & Prakash, L. (2004). Efficient and error-free replication past a minor-groove DNA adduct by the sequential action of human DNA polymerases iota and kappa. Mol Cell Biol 24, 5687-93. 180. Burgers, P. M. J. & Kunkel, T. A. (2017). Eukaryotic DNA Replication Fork. Annu Rev Biochem 86, 417-438. 181. Vaisman, A. & Woodgate, R. (2017). Translesion DNA polymerases in eukaryotes: what makes them tick? Crit Rev Biochem Mol Biol 52, 274-303. 182. Boudsocq, F., Iwai, S., Hanaoka, F. & Woodgate, R. (2001). Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4): an archaeal DinB-like DNA polymerase with lesion-bypass properties akin to eukaryotic poleta. Nucleic Acids Res 29, 4607-16. 183. Wong, J. H., Brown, J. A., Suo, Z., Blum, P., Nohmi, T. & Ling, H. (2010). Structural insight into dynamic bypass of the major cisplatin-DNA adduct by Y- family polymerase Dpo4. EMBO J 29, 2059-69.

120

184. Lee, E., Fowler, J. D., Suo, Z. & Wu, Z. (2017). Backbone assignment of the binary complex of the full length Sulfolobus solfataricus DNA polymerase IV and DNA. Biomol NMR Assign 11, 39-43. 185. Raper, A. T. & Suo, Z. (2016). Investigation of Intradomain Motions of a Y- Family DNA Polymerase during Substrate Binding and Catalysis. Biochemistry 55, 5832-5844. 186. Raper, A. T., Gadkari, V. V., Maxwell, B. A. & Suo, Z. (2016). Single-Molecule Investigation of Response to Oxidative DNA Damage by a Y-Family DNA Polymerase. Biochemistry 55, 2187-96. 187. Chu, X., Liu, F., Maxwell, B. A., Wang, Y., Suo, Z., Wang, H., Han, W. & Wang, J. (2014). Dynamic conformational change regulates the protein-DNA recognition: an investigation on binding of a Y-family polymerase to its target DNA. PLoS Comput Biol 10, e1003804. 188. Maxwell, B. A. & Suo, Z. (2013). Single-molecule investigation of substrate binding kinetics and protein conformational dynamics of a B-family replicative DNA polymerase. J Biol Chem 288, 11590-600. 189. Liu, S., Harada, B. T., Miller, J. T., Le Grice, S. F. & Zhuang, X. (2010). Initiation complex dynamics direct the transitions between distinct phases of early HIV reverse transcription. Nat Struct Mol Biol 17, 1453-60. 190. Antczak, N. M., Walker, A. R., Stern, H. R., Leddin, E. M., Palad, C., Coulther, T. A., Swett, R. J., Cisneros, G. A. & Beuning, P. J. (2018). Characterization of Nine Cancer-Associated Variants in Human DNA Polymerase kappa. Chem Res Toxicol. 191. Cancer Genome Atlas, N. (2012). Comprehensive molecular characterization of human colon and rectal cancer. Nature 487, 330-7.

121