A Multi-Disciplinary Investigation of Essential DNA Replication Proteins

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Varun V. Gadkari

Graduate Program in Biochemistry

The Ohio State University

2017

Dissertation Committee:

Dr. Zucai Suo, Advisor

Dr. Jane Jackman

Dr. Comert Kural

Dr. Richard Swenson

Copyrighted by

Varun V. Gadkari

2017

Abstract

An organism’s DNA is constantly under attack from various exogenous and endogenous DNA damaging agents. Thus, to assure survival, all living cells have evolved to maintain the genetic integrity of their DNA by various pathways. If left unrepaired,

DNA damage sites, or “lesions” can block DNA replication by stalling DNA polymerases, the enzymes responsible for DNA replication. Ultimately, if a stalled replication fork is not rescued, the cell will undergo apoptosis. To bypass DNA lesions, organisms in all domains of life initiate a process known as Translesion DNA Synthesis

(TLS). During TLS, the stalled replicative DNA polymerase is displaced by specialized

Y-family DNA polymerases that are capable of efficiently bypassing various forms of

DNA damage. While Y-family DNA polymerases are proficient in TLS, the fidelity of the process is a notable cause for concern. TLS mechanisms of the different Y-family

DNA polymerases vary greatly, and often introduce mutations in the DNA which can lead to carcinogenesis. Thus, the activity of Y-family DNA polymerases must be strictly regulated. To this end all living organisms depend on evolutionarily conserved sliding

DNA clamps which bind the DNA in a toroidal fashion, and slide along DNA during replication, serving as a scaffold for the DNA replication and repair machinery. During replication fork progression, the DNA clamp can regulate the various enzymatic activities by binding multiple proteins simultaneously as they process DNA. The overarching goal of my research has been to establish how the mechanisms of DNA replication, and TLS

ii affect DNA clamp mediated polymerase switching (pol switching) to allow for efficient

DNA replication while also preventing DNA mutagenesis.

My working hypothesis is that the DNA clamp is bound to both replicative and

TLS DNA polymerases at the replication fork, and that pol switching is governed by differences in DNA replication efficiency. Upon encountering a DNA lesion, replicative

DNA polymerases are unable to efficiently incorporate dNTPs and continue replication.

Thus, we hypothesize that the stalled replicative polymerases are exchanged in favor of

Y-family DNA polymerases by DNA clamp mediated pol switching to bypass DNA lesions. After TLS, pol switching ensures that the replicative polymerase returns to the replication fork for processive, high fidelity replication. My aim was to understand the

TLS mechanism to determine what parameter signals a pol switching event. I used pre- steady-state kinetics to study the TLS mechanism of Sulfolobus Solfataricus (Sso) Y- family DNA polymerase Dpo4 in response to a large helix-distorting lesion. Additionally,

I used single-molecule Förster Resonance Energy Transfer (smFRET), a fluorescence based technique, to investigate how the binding of Dpo4 to DNA is affected by an oxidative DNA lesion, and further, how a templating lesion can affect nucleotide selectivity. Additionally, I used smFRET to characterize the Sso DNA clamp

Proliferating Cell Nuclear Antigen, to understand the process of DNA clamp opening and closing. Collectively my research has addressed effects of DNA lesions on the DNA binding, nucleotide binding, and nucleotide incorporation by a model TLS DNA polymerase, and the nature of DNA clamp opening. These parameters, combined with previously established data will allow us to infer the mechanism of Pol switching.

iii

Dedicated to my mother, father, and brother. My three constant sources of love,

encouragement, and support.

Acknowledgments

First, and foremost, I would like to acknowledge my doctoral advisor, Dr. Zucai

Suo, for his guidance, patience, and support throughout my graduate career. Over the past few years he has been an outstanding mentor, and instilled in me a strong work ethic, which in combination with his guidance has helped me to achieve my goals as a doctoral student. I would also like to thank my graduate committee members, Dr. Jane Jackman,

Dr. Richard Swenson, and Dr. Comert Kural, for their support, and counsel throughout the course of my graduate career, and for creating an environment conducive to my growth as a scientist. I would also like to thank Dr. Jordi Torrelles, my undergraduate research advisor, for allowing an inexperienced undergraduate student to come into his lab and begin conducting exciting M. tb research. It was there that I discovered my interest in pursuing a career in scientific research.

I would like to acknowledge past and present members of the Suo lab, who have played an integral role in my growth as a scientist, but also as a person: Walter

Zahurancik, Jack Tokarsky, Austin Raper, Andrew Reed, Anthony Stephenson, Petra

Wallenmeyer, Kenny Phi, Dr. David Taggart, Dr. Rajan Vyas, Dr. Vimal Parkash, Dr.

Brian Maxwell, Saul Fredrickson, and Seth Klein. Working with this group of individuals has been a privilege, which I will never take for granted. I would particularly like to thank Walter Zahurancik. We started in the graduate program at the same time, as

v colleagues in the same lab, but quickly became close friends. Over the years, he has become my sounding board and confidant for matters regarding both science, and life in general. Without his friendship, and selfless attitude as a colleague, my graduate school experience would have never been as rewarding. I would also like to thank Dr. David

Taggart for his mentorship during my first two years of graduate school. Simply put, I credit him for teaching me how to be a good scientist. Additionally, I would like to thank

Dr. Brian Maxwell, for training me in the usage of fluorescence techniques, and our single-molecule TIRF microscope. Finally, I would like to thank Dr. Vicki Wysocki, and

Dr. Sophie Harvey for a fruitful collaboration.

I am grateful to The Ohio State University Department of Chemistry &

Biochemistry, namely the biochemistry labs on the 7th floor of the Biological Sciences building who have all at some point aided in my research by letting me borrow reagents, or helped me troubleshoot the roadblocks in my research. I would like to thank administrative staff of the Ohio State Biochemistry Program, especially former director

Dr. Tom Magliery, current director Dr. Jane Jackman, and Franci Brink, for everything that they do to keep OSBP running smoothly, and for all that they have done to help me.

I would like to thank my parents, Vinay and Varsha Gadkari, for their unwavering support throughout my life. They have always taught me to be the best version of myself, and to strive to achieve the highest goals in life. Their constant love and support has made me the person I am today, and for that I am forever grateful. I would also like to thank my little brother Viren, for his love and support. Finally, I would like to thank my friends who have made these last few years some of the best of my life. Their company

vi outside of graduate school has been unmeasurably beneficial. I am blessed to have met so many amazing people who I am certain will be close friends for life.

vii

Vita

2009-2012 ...... B.S. Biochemistry

The Ohio State University, Columbus, Ohio

2012-2017 ...... Ph.D. Biochemistry

The Ohio State University, Columbus, Ohio

2012-2013 ...... Graduate Teaching Associate,

Center for Life Sciences Education, The

Ohio State University, Columbus, Ohio

2013-2016 ...... Graduate Teaching Associate, Department

of Chemistry & Biochemistry, The Ohio

State University, Columbus, Ohio

Publications *Denotes Co-First Authors

1. Gadkari V.V.*, Harvey, S.R.*, Raper, A.T.*, Chu, W., Wang, J., Wysocki, V.,

and Suo, Z.* (2017) Investigation of Conformational Dynamics of a Sliding DNA

Clamp by Single-Molecule Fluorescence, Native Mass Spectrometry, and

Molecular Dynamics. J. Am. Chem. Soc. In Review

2. Raper, A.T.*, Reed, A.J.*, Gadkari, V.V., and Suo, Z.* (2017) Advances in

Structural and Single-Molecule Methods for Investigating DNA Lesion Bypass viii

and Repair Polymerases. Chem. Res. Toxicol. 30(1):260-269. doi:

10.1021/acs.chemrestox.6b00342.

3. Tokarsky, E.J., Gadkari, V.V., Zahurancik, W.J., Malik, C.K., Basu, A.K., and

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.

doi: 10.1016/j.dnarep.2016.08.002.

4. Raper, A.T.*, Gadkari, V.V.*, Maxwell, B.A., and Suo, Z.* (2016) Single-

Molecule Investigation of Response to Oxidative DNA Damage by a Y-Family

DNA Polymerase. Biochemistry. 55(14), 2187-2196.

5. 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. doi:

10.1016/j.dnarep.2014.06.003. doi: 10.1021/acs.biochem.6b00166.

6. Taggart, D.J., Fredrickson S.W., Gadkari, V.V., and 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(5), 931-940. doi:

10.1021/tx500088e.

Fields of Study

Major Field: Biochemistry

Table of Contents

Abstract ...... ii

Acknowledgments...... v

Vita ...... viii

Chapter 1 Introduction to DNA Replication and Repair ...... 1

1.1 DNA Replication and Repair ...... 1

1.2 DNA Polymerases ...... 2

1.3 Y-Family DNA Polymerases ...... 4

1.4 Mechanism of DNA Polymerases ...... 6

1.5 3-Nitrobenznthrone Lesions ...... 8

1.6 Oxidative DNA Damage ...... 9

1.7 Sliding DNA Clamps ...... 10

1.8 Focus of the Dissertation ...... 12

1.9 Schemes ...... 15

1.10 Figures ...... 19

Chapter 2 Mechanistic Investigation of the Bypass of a Bulky Aromatic DNA Adduct

Catalyzed by a Y-Family DNA Polymerase ...... 23 x

2.1 Introduction ...... 23

2.2 Materials and Methods ...... 25

2.2.1 Buffers ...... 25

2.2.2 Enzymes and DNA Substrates ...... 25

2.2.3 Running Start Assays...... 26

2.2.4 Electrophoretic Mobility Shift Assays ...... 27

2.2.5 Substrate Specificity Assays ...... 27

2.2.6 Biphasic Kinetic Assays ...... 28

2.3 Results ...... 29

2.3.1 Bypass of a Site-Specifically Placed dGC8-N-ABA Lesion by Dpo4 ...... 29

2.3.2 Moderate Effect of a dGC8-N-ABA Lesion on DNA Binding by Dpo4 ...... 30

C8-N-ABA 2.3.3 Kinetics of dNTP Incorporation in the presence of a dG lesion ...... 31

2.3.4 Biphasic Kinetics of dNTP Incorporation at Polymerase Pause Sites ...... 33

2.4 Discussion ...... 34

2.4.1 Polymerase Pause Sites ...... 34

2.4.2 Kinetic Basis for Polymerase Pausing as a Result of dGC8-N-ABA ...... 35

2.4.3 Large Effect of dGC8-N-ABA on the Kinetic Parameters of dNTP Incorporation 37

2.4.4 Biological relevance of the kinetic studies of dGC8-N-ABA ...... 42

2.5 Schemes ...... 44

2.6 Tables ...... 46

2.7 Figures ...... 51

Chapter 3 Single-Molecule Investigation of Response to Oxidative DNA Damage by a Y-

Family DNA Polymerase ...... 57

3.1 Introduction ...... 57

3.2 Materials and Methods ...... 59

3.2.1 Preparation of Protein and DNA ...... 59

3.2.2 Steady-state Fluorescence Spectroscopy Assays ...... 60

3.2.3 Single-molecule Measurements ...... 61

3.2.4 Verification of the FRET System ...... 62

3.2.5 Single Molecule Data Analysis ...... 62

3.2.6 Kinetic Assays ...... 65

3.3 Results ...... 65

3.3.1 Design of a FRET System for Monitoring Dpo4 Interaction with DNA ...... 66

3.3.2 Investigation of Dpo4 in a Binary Complex with DNA by smFRET ...... 66

3.3.3 Effect of dNTPs on Dpo4 Binding to Undamaged or Damaged DNA ...... 69

3.4 Discussion ...... 70

3.4.1 Analysis of the Low- and Mid-FRET States ...... 70

3.4.2 Analysis of the high-FRET State ...... 73

xii

3.4.3 Nucleotide Binding ...... 74

3.5 Schemes ...... 78

3.6 Tables ...... 80

3.7 Figures ...... 82

Chapter 4 Investigation of DNA Clamp Opening and Closing by Single-Molecule Forster

Resonance Energy Transfer ...... 96

4.1 Introduction ...... 96

4.2 Materials and Methods ...... 99

4.2.1 Expression of Proteins ...... 99

4.2.2 Purification of Proteins ...... 100

4.2.3 Single Molecule Data Acquisition ...... 101

4.2.4 Single Molecule Measurements ...... 101

4.2.5 Single Molecule Data Analysis ...... 102

4.3 Results ...... 104

4.3.1 Design of the Covalently Linked FRET Construct ...... 104

4.3.2 Single-Molecule FRET Population Histograms ...... 106

4.3.3 Dwell Time Analysis ...... 107

4.3.4 Effect of Replication Factor C on the Equilibrium of PCNA Molecules ...... 109

4.4 Discussion ...... 109

xiii

4.5 Schemes ...... 117

4.6 Tables ...... 118

4.7 Figures ...... 119

References ...... 124

xiv

List of Schemes

Scheme 1.1: Two metal Ion Mechanism of Nucleotidyl Transfer (From Steitz, T.A.,

1998)(37) ...... 15

Scheme 1.2: Kinetic Mechanism of DNA Polymerases ...... 16

Scheme 1.3: Environmental pollutant 3-Nitrobenzanthrone and the resulting bulky adduct lesion ...... 17

Scheme 1.4: Dual coding potential of 8-oxo-dG ...... 18

Scheme 2.1: Bioactivation of 3-Nitrobenzanthrone to form N-(deoxyguanosin-8-yl)-3- aminobenzanthrone (dGC8-N-ABA) ...... 44

Scheme 2.2: Proposed kinetic mechanisms of DNA lesion bypass ...... 45

Scheme 3.1: Proposed Mechanism of Binary and Ternary Complex Formation ...... 78

Scheme 3.2: Conformational sampling of the high-FRET state ...... 79

Scheme 4.1: Proposed Mechanism of PCNA Opening and Closing ...... 117

List of Tables

Table 2.1: DNA Substrates ...... 46

Table 2.2: Binding Affinity of Dpo4 to Damaged and Control DNA Substrates ...... 47

Table 2.3: Kinetic parameters for dNTP incorporation into damaged DNA substrates containing dGC8-N-ABA ...... 48

Table 2.4: Kinetic parameters for dNTP incorporation into a control DNA substrate

18/26-mer...... 49

Table 2.5: Biphasic kinetic parameters for correct dNTP incorporation into DNA

Substrate in the presence of DNA trap...... 50

Table 3.1: Shuttling Rates from Transition Density Plot ...... 80

Table 3.2: Dwell Time Analysis of Dpo4 Binding to DNA ...... 81

Table 4.1 Rates of PCNA opening and closing ...... 118

xvi

List of Figures

Figure 1.1: Conserved structure of DNA polymerases ...... 19

Figure 1.2 DNA polymerase fidelities (From Kunkel, T.A., 2004)(182) ...... 20

Figure 1.3: Conserved structure of Y-family DNA polymerases ...... 21

Figure 1.4: Evolutionarily Conserved Sliding DNA Clamps ...... 22

Figure 2.1: Running start assay ...... 51

bypass extension Figure 2.2: Determination of t50 and t50 ...... 52

Figure 2.3: Determination of Kd,DNA ...... 53

Figure 2.4: Kinetics of dATP incorporation onto 18/26-mer-dGC8-N-ABA ...... 54

Figure 2.5: Biphasic kinetics of correct dNTP incorporation in the presence of a DNA trap ...... 55

Figure 2.6: Quantitative effects of a dGC8-N-ABA lesion on DNA binding and correct dNTP incorporation catalyzed by Dpo4 ...... 56

Figure 3.1: Single-molecule FRET analysis of Dpo4 binding to DNA ...... 82

Figure 3.2: Verification of Observed FRET ...... 83

Figure 3.3: Fluorescence Assay for Multiplicity of Binding ...... 84

Figure 3.4: DNA binding affinity of Dpo4 ...... 85

Figure 3.5: Burst Assay to Confirm Dpo4 Activity ...... 86

Figure 3.6: FRET from Dpo4 binding to single DNA molecules ...... 87

xvii

Figure 3.7: Transition Density Plots ...... 88

Figure 3.8: Dwell Time Analysis of Dpo4 Binding Undamaged DNA ...... 89

Figure 3.9: Dwell Time Analysis of Dpo4 Binding Damaged DNA ...... 90

Figure 3.10: Example Dwell Time Survivor Function ...... 91

Figure 3.11: FRET Histograms of Dpo4 binding DNA with incorrect dNTP ...... 92

Figure 3.12: FRET from Dpo4 binding to single undamaged DNA molecules in the presence of saturating dCTP ...... 93

Figure 3.13: Representative FRET Trajectory for Dpo4 binding DNA in the presence of saturating incorrect nucleotide ...... 94

Figure 3.14: Selectivity of Dpo4 for the Correct Nucleotide...... 95

Figure 4.1: PCNA interconverts between its open and closed conformations...... 120

Figure 4.2: Design of a two state FRET system ...... 119

Figure 4.3: FRET efficiency population histograms in changing concentrations of NaCl

...... 121

Figure 4.4: Dwell Time Analysis ...... 122

Figure 4.5: FRET distribution of PCNA1-2-3 in the presence of RFC ...... 123

xviii

Chapter 1 Introduction to DNA Replication and Repair

1.1 DNA Replication and Repair

The accurate and efficient replication of DNA is essential for the survival of all living organisms. However, DNA is susceptible to various endogenous and exogenous sources of damage such as reactive oxygen species, environmental pollution, and UV radiation. Thus, all living cells have evolved pathways to conserve the genomic integrity central to their very existence. High-fidelity replicative DNA polymerases are responsible for faithful template-dependent DNA replication, and repair pathways such as base excision repair (BER) and nucleotide excision repair (NER) identify and remove damaged DNA base(s). Additionally, if unrepaired DNA damage sites, or “lesions”, are encountered during DNA replication, a specific group of polymerases known as Y-family polymerases are recruited to bypass the lesion in a process referred to as translesion DNA synthesis (TLS). However, due to the low fidelity of the Y-family polymerases, their activity must be regulated by switching back to the high fidelity replicative polymerases after the lesion is bypassed.

To coordinate the various enzymes and proteins involved in DNA maintenance pathways, all living cells depend on an evolutionarily-conserved sliding DNA clamp protein. These DNA clamps encircle double-stranded DNA in toroidal fashion and slide along DNA, acting as both processivity-promoting factors for associated DNA 1 polymerases and as scaffolds to tether other replication and repair machinery to DNA at the replication fork. Sliding DNA clamps are typically multimeric, and can associate with different proteins involved in the replication and repair pathways simultaneously. Thus,

DNA clamps are hypothesized to be essential in orchestrating multi-protein processes such as polymerase switching during TLS. The overarching goals of this dissertation are to utilize an interdisciplinary approach to characterize the kinetic mechanism and conformational dynamics of a model Y-family DNA polymerase involved in TLS, and examine the conformational dynamics of an evolutionarily conserved DNA clamp.

1.2 DNA Polymerases

DNA polymerases (Pols) are the enzymes responsible for DNA replication in all domains of life. Since the discovery of the first DNA polymerase, Escherichia coli (E.

Coli) DNA polymerase I,1-5 extensive research has been dedicated to investigating the structural and mechanistic aspects of the enzymes responsible for one of the most important natural processes in all living organisms. The first crystal structure of a DNA polymerase, E. coli DNA Pol I Klenow Fragment, revealed that Pol I bound DNA in a cleft between three subdomains that resembled a human hand.6 As research on DNA polymerase phylogeny,7,8 structure,9-21 and mechanism22-36 accelerated over the years, it was established that all DNA polymerases possess a conserved core domain oriented in a

“right hand” conformation composed of three canonical subdomains termed finger (F), palm (P), and thumb (T) (Figure 1.1). During DNA replication, the polymerase “grasps”

DNA using the finger and thumb subdomains and orients the terminal 3′-hydroxyl of the

DNA primer into the catalytic active site located in the palm subdomain. The thumb

2 stabilizes the DNA binding interaction of the DNA polymerase core, while the finger assures the proper positioning of the incoming nucleotide (dNTP) for catalysis. In the palm subdomain, three conserved carboxylic acid side chains coordinate two divalent

Mg2+ ions that facilitate the chemical catalysis of nucleotide incorporation (Scheme

1.1).37

Despite such strong structural and mechanistic conservation, there is little sequence similarity between known DNA polymerases aside from the conserved polymerase motifs which include the active site catalytic residues. Thus, it is fascinating that despite lack of strong primary sequence similarity, DNA polymerases in all kingdoms of life have evolved to perform the same fundamental catalytic function. DNA polymerases are phylogenetically classified into six distinct families (A, B, C, D, X, Y) based on their homology, or lack thereof, to DNA polymerases initially discovered in E. coli.7,8,38 DNA polymerases from different families, and even within families, vary significantly in their fidelity (Figure 1.2), processivity, and DNA substrate preference.

These differences are largely due to structural features unique to the different families.

For example, most A- and B-Family DNA polymerases possess N-terminal exonuclease subdomains in addition to the canonical polymerase core domain.39,40 These exonuclease subdomains excise misincorporated nucleotides and allow replication of DNA with significantly higher fidelity (Figure 1.1, 1.2).24,27,32,35,41 In contrast, TLS polymerases of the Y-family have a unique “Little Finger” (LF) subdomain that makes unique contacts with the DNA during DNA binding and replication (Figure 1.1).42,43 The structure and function of the LF has been implicated in the bypass capabilities of the various Y-family

DNA pols by affecting the efficiency and fidelity with which a particular Y-family polymerase responds to specific a DNA lesion.43 Due to the wide variety of roles and activities associated with DNA polymerases, structural and mechanistic studies of this group of enzymes remains an important and fundamental topic in biology.

1.3 Y-Family DNA Polymerases

Although pathways exist to repair damaged DNA bases in double-stranded DNA, a replication fork often encounters damage and must be able to bypass a templating DNA lesion before repair of that lesion can be completed. However, previous studies have shown that high fidelity replicative polymerases can be completely stalled by some DNA lesions.44,45 Thus, the Y-family DNA polymerases have been identified in all three domains of life, e.g. four in humans (DNA polymerases η, κ, ι, and Rev1), four in

Saccharomyces cerevisiae (DNA polymerases η, κ, ι and Rev1), two in Escherichia coli

(DNA polymerases IV and V), and one in Sulfolobus solfataricus (DNA polymerases IV)

(Figure 1.3). The Y-family DNA polymerases are responsible for DNA lesion bypass in vivo during TLS.46-48 Y-family DNA polymerases are capable of DNA lesion bypass due to key structural differences compared to replicative DNA polymerases. While Y-family

DNA polymerases possess a canonical polymerase core domain consisting of palm, thumb, and finger subdomains, the thumb and finger subdomains are unusually small, resulting in larger, more solvent-accessible active sites to accommodate and incorporate an incoming nucleotide opposite DNA lesions of varying size and structure.42,45,49-59

Additionally, Y-family DNA polymerases also contain a C-terminal subdomain known as the little finger (LF),42,43 or polymerase associated domain (PAD),60 which is connected

4 to the thumb domain by a flexible linker. This unique LF subdomain has been extensively studied, and has been shown to impact the DNA binding61 and DNA lesion preferences of

Y-family DNA polymerases.43

Notably, the same structural nuances that allow the Y-family DNA polymerases to catalyze efficient TLS also result in their low fidelity and processivity when replicating undamaged DNA.62 The larger, solvent-exposed active sites are less effective in Watson-

Crick base pair geometry recognition for nucleotide selectivity. Moreover, their lack of an exonuclease domain prevents misincorporation removal. As a result of these shortcomings in fidelity, the Y-family DNA polymerases have also been shown to incorporate antiviral nucleoside analog drugs, leading to speculation that Y-family DNA polymerases might be partially responsible for the clinical toxicity of these drugs.63-65

Due to these adverse features, switching between replicative and Y-family DNA polymerases during TLS is imperative so that introduction of mutations into DNA is minimized while still allowing for efficient TLS.

As the only Y-family DNA polymerase encoded by Sulfolobus solfataricus

(Sso),66 DNA polymerase IV (Dpo4) is hypothesized to be responsible for all lesion bypass activities in vivo, and demonstrates lesion bypass capabilities similar to human

DNA Polymerase η (hPol η) in vitro. It can be recombinantly overexpressed and purified in large quantities from E. coli, and due to its inherent thermostability Dpo4 can retain activity during prolonged assays at 37⁰C. Due to these factors, Dpo4 has long been considered a model Y-family enzyme, and is the most comprehensively studied Y-family

DNA polymerase to date.42,43,45,49-52,54,55,57,61,63,67-110 Our lab has played an essential role in

5 establishing the kinetic mechanisms of polymerization,78,79 fidelity,77 and lesion bypass45,49-52,54,55,57,63,67,68,70-72,74,80,81 of this model Y-family polymerase. Two recent lesion bypass studies of Dpo4 will be a major focus of this dissertation.54,55

1.4 Mechanism of DNA Polymerases

Pre-steady-state kinetic studies31,35,111,112 of various DNA polymerases among the different families have aided in establishing a conserved minimal kinetic mechanism to which all polymerases adhere (Scheme 1.2).31 DNA polymerization begins with the binding of a DNA polymerase (E) to the DNA substrate (DNAn) at the primer/template junction (Step 1) to form the binary complex (E•DNAn). Next, the binary complex

(E•DNAn) binds to a dNTP to form the loose ground-state ternary complex

(E•DNAn•dNTP) (Step 2). The binding of correct dNTP induces a conformational change in the DNA polymerase (E to Eʹ) to form the tight activated ternary complex

(E′•DNAn•dNTP) (Step 3) which includes active site reconfiguration for the catalysis of phosphodiester bond formation (Step 4). After nucleotidyl transfer is complete, the enzyme remains tightly bound to the extended DNA product (DNAn+1) and pyrophosphate (PPi) in the post-catalytic ternary complex (E′•DNAn+1•PPi). The post- catalytic complex undergoes a reverse conformational change (Eʹ to E) to form a loosely bound post-catalytic complex (E•DNAn+1•PPi) (Step 5), followed by the release of pyrophosphate to return to the binary complex extended by one nucleotide (E•DNAn+1)

(Step 6). After Step 6, the DNA polymerase can either dissociate from the extended DNA product, or remain bound and continue subsequent rounds of nucleotide incorporation.

The conformational change in Steps 3 and 5 can be a major subdomain movement

(finger “closing” or “opening”), a global conformational change involving multiple subdomains moving in relation to each other, a slight repositioning of active site residues, or any combination of minor and major structural realignments.55,61,68,69,71,74-76,113-125 The conformational changes in the minimal kinetic mechanism (Steps 3 & 5) have been extensively studied as they are requisite for catalysis of nucleotidyl transfer (Step 4) and pyrophosphate release (Step 6), respectively. For this reason, there has been a long- standing debate regarding the physical nature of the rate-limiting step of nucleotide incorporation. It was long assumed that the closing of the finger subdomain upon nucleotide binding observed in crystallo119,126 was the rate-limiting conformational change. However, investigations of various DNA polymerases by solution-based fluorescence studies revealed that that finger subdomain motion is far too rapid to be rate- limiting.35,114-116,127-132 The current opinion in the field suggests that the rate-limiting step is an active site rearrangement such as the repositioning of catalytic residues, or intradomain motion within the subdomains, which occurs after major domain movements but before chemistry.31,74,76

The domain motions of Y-family DNA polymerases have been extensively studied by our lab55,61,69,71,74,75 by solution-based pre-steady-state kinetics and single- molecule fluorescence techniques to elucidate the contribution of conformational dynamics to their mechanism. Crystallographic evidence indicates that these DNA polymerases do not undergo a large finger subdomain motion as observed with other polymerases, but rather that the finger subdomain is permanently in the “closed” conformation resulting in a “preformed” active site even in the absence of DNA or

7 dNTP.42,60,133,134 Instead, the largest conformational changes occur in the little finger subdomain which rotates relative to the polymerase core to bind the DNA substrate (Step

1, Scheme 1.2).60-62,68,69,74,135-137 Pre-steady-state kinetic studies of various Y-family

DNA polymerases have indicated, however, that these enzymes follow a similar minimal mechanism for replication of undamaged DNA, including a pre-chemistry rate-limiting conformational change step, despite their structural and conformational differences.77,78,112,138-145 Thus, defining these conformational changes is critical to understanding the mechanism of DNA synthesis and lesion bypass catalyzed by Y-family

DNA polymerases.

Lesion bypass studies have demonstrated that Y-family DNA polymerases employ a variety of mechanisms to bypass different lesions. Notably, in recent years our lab has conducted various pre-steady-state investigations to determine how the structure and kinetics of Dpo4, are affected in response to various DNA lesions.45,49-

52,54,55,57,70,71,75,80,81 The mechanisms of bypass of two different DNA lesions by a model

Y-family DNA polymerase, Sso Dpo4, were recently investigated using pre-steady-state kinetics and single-molecule FRET, and will be presented in this dissertation.54,55

1.5 3-Nitrobenznthrone Lesions

One of the common sources of exogenous DNA damage is pollution resulting from fossil fuel combustion. Many of these airborne environmental pollutants are potent mutagens, which are implicated in DNA damage and carcinogenesis upon ingestion.146

One such environmental pollutant is 3-Nitrobenzanthrone (3-NBA), a nitropolyaromatic hydrocarbon released in diesel exhaust and found in ambient air particulate matter

(Scheme 1.3). 3-NBA is especially prevalent in urban areas and areas exposed to high usage of heavy machinery (i.e. construction), and thus presents a significant public and occupational health hazard.147,148 Furthermore, 3-NBA is one of the most mutagenic compounds ever tested on the Ames Salmonella typhimurium (TA98) mutagenicity test,149 and is suspected to be carcinogenic in humans.150-156 Once ingested, 3-NBA is bioactivated by a series of metabolic pathways, to form a reactive intermediate which readily reacts with guanosine and adenosine bases resulting in adducted DNA lesions.153

These adducted lesions, which are often 2- to 3-fold larger than a standard DNA base, are referred to as helix distorting lesions, and can cause a complete replication fork stall, potentially inducing apoptosis.157 The major DNA lesion resulting from 3-NBA metabolism is N-(deoxyguanosin-8-yl)-3-aminobenzanthrone (dGC8-N-ABA) (Scheme 1.3).

It has been found to completely block DNA replication, and requires a TLS polymerase for lesion bypass.54,157 Although Y-family DNA polymerases are uniquely structured to bypass DNA lesions, they still experience great difficulty in bypassing bulky DNA lesions.52-54,56 Consistently, dGC8-N-ABA has been shown to stall multiple Y-family DNA polymerases in running-start DNA polymerization assays.54,56 An investigation of the kinetic mechanism of dGC8-N-ABA bypass by the model Y-family DNA polymerase Dpo4 is a major topic of this dissertation.

1.6 Oxidative DNA Damage

In addition to exogenous sources of damage, the genome is constantly under attack from endogenous sources of DNA damage. For example, aerobic respiration, a process vital for survival, produces reactive oxygen radicals known to be potent DNA

9 damaging agents. One of the most common oxidative lesions found in cells, 7,8-dihydro-

8-oxoguanine (8-oxo-dG), results from oxidation of the C8 atom of guanine (Scheme

1.4). Structural studies have established that 8-oxo-dG is not a helix distorting lesion.158,159 However, the true mutagenic potential of 8-oxo-dG lies in its dual coding potential. Due to free rotation about the N-glycosidic bond the 8-oxo-dG lesion can adopt both an anti-conformation, which forms a canonical Watson-Crick base pair with cytosine, and a syn-conformation, which forms a Hoogsteen base pair with adenine.160

Thus, DNA polymerases vary in their nucleotide incorporation preference opposite 8- oxo-dG, with some favoring the misincorporation of dATP, while others preferentially incorporate correct dCTP.161-163 Structural evidence suggests that differences in response

8-oxo-dG result from how the lesion is accommodated in the active site of a DNA polymerase. The carbonyl group at the C8 position can form unique interactions with residues within the DNA polymerase active site, preferentially stabilizing either the anti or syn conformation of the DNA lesion.92,164 The response of a model Y-family DNA polymerase Dpo4 to 8-oxo-dG by is one of the major topics of this dissertation.

1.7 Sliding DNA Clamps

In all domains of life, DNA replication and maintenance is an essential and complex task requiring multiple proteins and enzymes working in concert. This intricate process, in which there is little tolerance for error, is coordinated by an evolutionarily conserved sliding DNA clamp, which encircles the DNA duplex and regulates the various

DNA processing proteins.165-167 While bound to DNA, the clamp can associate with other proteins acting upon the DNA thereby acting as a “tool belt.” Sliding DNA clamps have

10 been identified and studied in all domains of life, e.g. the β clamp in Escherichia coli, the gene 45 protein in T4 bacteriophage (gp45), and proliferating cell nuclear antigen

(PCNA) in yeast, humans, and the archaeon Sulfolobus solfataricus (Figure 1.4).168,169

Despite low sequence similarity, all sliding DNA clamps have a remarkably similar ring structure with the exception of the gp45 homotrimer, which is more triangular in shape.

All DNA clamps have a conserved toroidal structure with a central hole to accommodate a DNA duplex, and have evolved to perform the same essential role in all living organisms.169

The complexity of the clamp subunit organization varies between organisms. The bacterial β clamp is a homodimer as observed in E. coli, while PCNA clamps in archaeal and eukaryotic organisms are trimeric. Furthermore, while most eukaryotic PCNAs, including human and yeast, are homotrimeric, PCNA from Sso is a heterotrimer consisting of three distinct PCNA monomer subunits: PCNA1, 2, and 3. In all organisms the DNA clamps are known to associate with proteins while they interact with DNA, and in some cases they stimulate the activity of associated enzymes.85,170-179 The heterotrimeric structure of Sso PCNA makes it a particularly good model to study the

DNA replication “tool belt”, as it is hypothesized that the different monomers of the Sso

DNA clamp have specific binding partners. For example the PCNA1 monomer has been shown to interact with Dpo485 and flap endonuclease 1 (FEN1),178 PCNA2 with replicative Sso DNA polymerase I (PolB1),180 and PCNA3 with Sso DNA ligase.179

Furthermore, the three separate interfaces between the subunits of heterotrimeric Sso

PCNA have been studied, and the opening interface has been identified to be at the

PCNA1:PCNA3 interface. Thus, we can expand on this finding to study the dynamics of

PCNA opening and closing, either alone or in the presence of Replication Factor C

(RFC), an evolutionarily conserved clamp loader. RFC and its homologues, which can be found in all domains of life, function to open closed DNA clamp molecules, usually in an

ATP-dependent manner.169 The conformational equilibrium of open and closed PCNA, and the factors that affect this equilibrium, is one of the major topics of this dissertation.

1.8 Focus of the Dissertation

In the Suo lab, our goal is to investigate the mechanism of DNA clamp-mediated polymerase switching, which is necessitated by both the inability of replicative DNA polymerases to bypass DNA lesions and the inherently low fidelity of Y-family DNA polymerases that rescue stalled replication forks. The same attributes that allow Y-family

DNA polymerases to bypass DNA lesions also bestow them with very low fidelity of replication. Therefore, it is imperative that Y-family DNA polymerases are strongly regulated to prevent the introduction of mutations after the DNA lesion has been bypassed. We believe that the polymerase switch is triggered when a replicative polymerase stalls, and when a low processivity Y-family DNA polymerase unbinds after lesion bypass. To this end, my research has primarily focused on the characterization of

DNA damage response by a model Y-family DNA polymerase Sulfolobus Solfataricus

Dpo4. We hypothesize that the Y-family polymerases are less efficient at incorporating nucleotides once extending the lesion bypass product, thus signaling a polymerase switch back to a higher fidelity replicative DNA polymerase. To test this hypothesis, I have applied two techniques to study Dpo4 in response to two different DNA lesions.

Additionally, I have investigated the heterotrimeric PCNA clamp from Sso to determine whether the clamp can exist in an open conformation in the absence of the clamp loader

Replication Factor C. Once I established the equilibria of open vs. closed PCNA, I investigated the role of RFC in opening the DNA clamp.

Our lab was the first to establish the kinetic mechanism of DNA replication by the model Y-family translesion DNA polymerase from Sulfolobus solfataricus Dpo4. Since then we have expanded our understanding of Dpo4 by testing its ability to bypass various lesions. Due to its enzymatic similarities to human DNA polymerases η and κ, Dpo4 is considered an ideal model to predict the potential mutagenic and mechanistic hurdles of newly discovered DNA lesions. The results presented in Chapter 2 are from one such study in which the lesion bypass capabilities of Dpo4 were tested against, N-

(deoxyguanosin-8-yl)-3-aminobenzanthrone, a bulky adduct DNA lesion product of 3-

NBA, an airborne pollutant. Pre-steady-state kinetics were used to determine the mechanism of bypass and extension of the 3-NBA lesion. Additionally, we performed a complete fidelity analysis to determine the mutagenic potential of the 3-NBA-derived lesion. In Chapter 3, single-molecule FRET was used to study the conformational dynamics and substrate binding kinetics of Dpo4 in response to a common oxidative

DNA lesion, 8-oxo-dG. We studied the nucleotide selectivity of Dpo4 opposite the dual- coding lesion, and investigated how the presence of 8-oxo-dG impacted DNA and nucleotide binding by Dpo4.

In Chapter 4, single-molecule FRET was utilized to study the equilibria of PCNA molecules in solution. We first probed whether the PCNA molecules could exist in the

13 open conformation in solution in the absence of the ATP-dependent clamp loader RFC.

Once it was established that the clamp does indeed exist in both open and closed forms in solution, we set out to probe the physical determinants of this equilibrium. A variety of salt conditions were tested and a correlation between PCNA opening and increasing salt concentration in solution was established. Finally, experiments were conducted with RFC to determine how the presence of the clamp loader affected the equilibrium of the PCNA open-closed states.

Overall, the goal of my research has been to characterize one of the polymerases that is involved in DNA replication, while also investigating the DNA clamp responsible for acting as the “tool belt” during DNA replication and lesion bypass. Our goal is to combine our knowledge of the DNA polymerization and DNA clamp loading mechanisms to reconstitute the DNA replication tool belt and study the mechanisms of

DNA polymerase switching between Dpo4 and PolB1, the replicative polymerase from

Sso.

1.9 Schemes

Scheme 1.1: Two metal Ion Mechanism of Nucleotidyl Transfer (From Steitz, T.A., 1998)37

Scheme 1.2: Kinetic Mechanism of DNA Polymerases

Scheme 1.3: Environmental pollutant 3-Nitrobenzanthrone and the resulting bulky adduct lesion

Scheme 1.4: Dual coding potential of 8-oxo-dG

1.10 Figures

Figure 1.1: Conserved structure of DNA polymerases (A) Replicative B-family DNA polymerase from Sulfolobus solfataricus (Sso), DNA

Polymerase B1, is shown in the apo-state. Replicative DNA pols have a canonical polymerase core domain (Finger, Palm, Thumb), and an exonuclease domain. (B) Y- family DNA polymerase from Sso, Dpo4 is shown in a ternary complex. Like all Y- family DNA pols, in addition to the core polymerase domain, Dpo4 has a little finger subdomain. PDB codes: 1S5J181, 1JX442

Figure 1.2 DNA polymerase fidelities (From Kunkel, T.A., 2004)182 The various DNA polymerase families exhibit a wide range of DNA replication fidelities.

This figure shows the likelihood of the different DNA pol families causing single base substitution, or deletion errors.

Figure 1.3: Conserved structure of Y-family DNA polymerases The structure of Y-family DNA polymerases is strongly conserved across all domains of life. All Y-family DNA polymerases have a canonical Polymerase core domain, consisting of the Finger, Palm and Thumb. Additionally, they all have a little finger subdomain, known to make contacts with the DNA during replication. (A) E. coli DNA

Pol IV (DinB), (B) Sso Dpo4, (C) Human DNA Pol η, (D) Yeast DNA Pol η. PDB

Codes: (A) 4IR1183, (B) 1JX442, (C) 3MR2184, (D) 3OHA185

Figure 1.4: Evolutionarily Conserved Sliding DNA Clamps The sliding DNA clamps are conserved in all domains of life. Despite having very little sequence similarity, the overall structure of the clamp is strongly conserved in various organisms. (A) β Clamp from E. coli is a homodimer. (B) Gene protein 45 (gp45) from

T4 bacteriophage is a homotrimer. (C) Proliferating Cell Nuclear Antigen (PCNA) from humans is a homotrimer. (D) Unique PCNA from Sulfolobus solfataricus is a heterotrimer, made up of monomeric subunits PCNA1, PCNA2, and PCNA3. PDB

Codes: (A) 2POL186, (B) 1CZD187, (C) 1AXC188, (D) 2IX2189

Chapter 2 Mechanistic Investigation of the Bypass of a Bulky Aromatic DNA

Adduct Catalyzed by a Y-Family DNA Polymerase

2.1 Introduction

Combustion of every form of fossil fuel produces potent environmental pollutants that are known to affect human health at the molecular level.146 Once metabolized, many of these pollutants become mutagenic and carcinogenic by damaging cellular genomes in a variety of ways, by forming bulky DNA adducts, causing oxidative damage, and producing single and double stranded DNA breaks.146 If the DNA adducts are not recognized and repaired by various cellular DNA repair pathways, they will stall replication machinery and eventually induce apoptosis.157 To rescue stalled DNA replication, cells switch from a replicative polymerase to a DNA lesion bypass polymerase at a lesion site.157 Notably, most of these lesion bypass polymerases belong to the Y-family, one of the six families (A, B, C, D, X, and Y) of DNA polymerases.46

The Y-family enzymes possess relatively flexible and solvent-accessible active sites to accommodate bulky DNA lesions. However, these attributes of the Y-family enzymes also lead to their error-prone manner of DNA synthesis with both undamaged and damaged DNA.182,190 Interestingly, the Y-family DNA polymerases have been identified in all three domains of life, e.g. four in humans (DNA polymerases η, κ, ι, and Rev1), three in Schizosaccharomyces pombe (DNA polymerases η, κ, and Rev1), two in 23

Escherichia coli (DNA polymerases IV and V), and one in Sulfolobus solfataricus (DNA polymerases IV). Being the lone Y-family enzyme in S. solfataricus, DNA polymerases

IV (Dpo4) likely bypasses various lesions in vivo and thus, has been intensely studied as a model enzyme in vitro. With an undamaged DNA template, Dpo4 catalyzes DNA synthesis with a fidelity of one error per 1,000 –10,000 nucleotide incorporations based on pre-steady-state kinetic studies from 37 to 56 °C.77,78 Dpo4 has been found to bypass various DNA lesions, e.g. abasic sites 49,50 and N-(deoxyguanosin-8-yl)-1-aminopyrene

(dGAP).52 The latter lesion is a bulky adduct resulting from a dG base reacting with the metabolites of 1-nitropyrene, a product of incomplete diesel and gasoline combustion.191

Another interesting product of diesel exhaust, albeit present at a lower concentration than 1-nitropyrene, is 3-nitrobenzanthrone (3-NBA, Scheme 2.1).149 3-

NBA, one of the most mutagenic compounds ever tested on the Ames Salmonella typhimurium (TA98) assay, is also found in ambient air particulate matter.149 The mutagenicity of 3-NBA is comparable to 1,8-dinitropyrene, 192 and has been suspected to be carcinogenic to humans.150-152 This possibility is supported by cellular and animal model data.153-156 For example, 3-NBA and its metabolites are found to induce micronuclei and DNA adducts in mouse and human cells,149,155 and form DNA adducts and tumors in rats.148,154-156 Bioactivation of 3-NBA is initiated by an essential nitroreduction step, forming N-hydroxy arylamine (N-OH-ABA) (Scheme 2.1).153 This step is catalyzed by cytochrome P450 and cytochrome P450 reductase in rats,153 and

NAD(P)H:Quinone Oxidoreductase in humans.147 N-OH-ABA is then metabolized to either N-AcO-ABA or N-OSO3H-ABA by N,O-acetyltransferases or sulfotransferases,

24 respectively.153,193 Finally, either of these highly reactive derivatives reacts with DNA to form bulky aromatic DNA adducts including N-(deoxyguanosin-8-yl)-3- aminobenzanthrone (dGC8-N-ABA, Scheme 2.1).

The DNA adducts derived from 3-NBA, if not repaired, are likely bypassed by lesion bypass polymerases in vivo.157 In this paper, we kinetically investigated the potential bypass of a site specifically placed dGC8-N-ABA, one of the three major adducts formed by 3-NBA,194 catalyzed by Dpo4. Our results show that the model Y-family polymerase was indeed capable of bypassing dGC8-N-ABA but its polymerase efficiency and fidelity were significantly affected by the bulky lesion.

2.2 Materials and Methods

2.2.1 Buffers

All pre-steady-state kinetic assays were performed in optimized reaction buffer R

(50 mM HEPES, pH 7.5, 5 mM MgCl2, 50 mM NaCl, 0.1 mM EDTA, 5 mM DTT, 10% glycerol (v/v), and 0.1 mg/ml bovine serum albumin) 5. All electrophoresis mobility shift assays (EMSA) were performed in buffer S (50 mM Tris-Cl, pH 7.5 at 23 °C, 5 mM

MgCl2, 50 mM NaCl, 5 mM DTT, 10% glycerol (v/v), and 0.1 mg/ml bovine serum albumin). The EMSAs were conducted using running buffer A (50 mM Tris acetate, pH

7.5, at 25 °C, 0.5 mM EDTA, and 5.5 mM magnesium acetate). All concentrations are final after mixing. Unless otherwise noted, all reactions were performed at 37 °C.

2.2.2 Enzymes and DNA Substrates

Full length Dpo4 was expressed in E. coli and purified as previously described.77,78 The DNA template containing the dGC8-N-ABA was synthesized as

25 previously described. All primers and templates except 26-mer-dGC8-N-ABA in Table 2.1 were purchased from the Integrated DNA Technologies, and purified by denaturing polyacrylamide gel electrophoresis (PAGE). The DNA primers were 5-32P-labeled by incubation with [-32P] ATP (Perkin Elmer) and OptiKinase (United States Biochemical) for 3 h at 37 °C. The 5-32P-labeled primers were annealed to the unlabeled control 26- mer or 26-mer-dGC8-N-ABA at a molar ratio of 1.00:1.35. The solution was first denatured at 95 °C (for undamaged DNA) or 75 °C (for 26-mer-dGC8-N-ABA) for 5 minutes and then slowly cooled to room temperature.

2.2.3 Running Start Assays

The running start assays were performed as previously described.49-51 Briefly, 5-

32P-labeled DNA (100 nM) and Dpo4 (100 nM) were preincubated in buffer R and subsequently mixed rapidly with a solution containing all four dNTPs (200 μM each) at

37 °C via rapid chemical-quench flow apparatus (KinTek). The reactions were quenched by the addition of EDTA to 0.37 M at specific time points. The reaction products were separated by denaturing PAGE (17% polyacrylamide, 8 M urea) and quantitated using a

Typhoon Trio (GE Healthcare).

Quantitative analysis of the running start assays was performed by determining the relative lesion bypass efficiencies (dGC8-N-ABA bypass %) as a function of reaction time. For each time point t, dGC8-N-ABA bypass% = (B/E) x 100, where the total dGC8-N-ABA bypass events (B) was calculated from the concentration of all intermediate products with sizes greater than or equal to the 21mer, and the total dGC8-N-ABA “encounter” events (E) equaled the summation of the 20-mer concentration and the total dGC8-N-ABA bypass

C8-N-ABA bypass events (B). To quantitatively define the dG bypass efficiency, t50 was defined as the time required to bypass 50% of the total dGC8-N-ABA lesions encountered.

2.2.4 Electrophoretic Mobility Shift Assays

Buffer S containing 5-32P-labeled DNA (5 nM) was titrated with Dpo4 (0 – 220 nM). Binary complexes were then separated from free DNA by native PAGE at a constant voltage of 80 V for 30 min at 25 °C using running buffer A. Complex formation was quantitated by using a Typhoon Trio (GE Healthcare). The plot of the binary complex (Dpo4•DNA) concentrations versus Dpo4 concentrations was fit to Equation 2.1 to yield Kd, DNA, the equilibrium dissociation constant for the binary complex

(Dpo4•DNA) at 25 °C.

2 1/2 [Dpo4•DNA] = 0.5(Kd, DNA + E0 + D0) – 0.5[(Kd, DNA + E0 + D0) – 4E0D0] (Eq. 2.1)

In Equation 2.1, E0 is the active Dpo4 concentration while D0 is the DNA concentration.

2.2.5 Substrate Specificity Assays

A previously described single-turnover dNTP incorporation assay was employed to determine the maximum dNTP incorporation rate constant (kp), and equilibrium

49,51,77,78 dissociation constant (Kd, dNTP) of an incoming dNTP. These two kinetic parameters yield the dNTP incorporation efficiency (kp/Kd, dNTP). Briefly, Dpo4 (120 nM) and 5-32P-labeled DNA substrate (30 nM) were preincubated in Buffer R and then mixed rapidly with a solution containing increasing concentrations of dNTP. The reactions were quenched at specific time points by the addition of EDTA to 0.37 M. Reaction

27 samples were analyzed by denaturing PAGE (17% acrylamide, 8 M urea) and quantitated by using a Typhoon Trio (GE Healthcare). The plot of product formation versus time for each dNTP concentration was fit to a single exponential equation (Equation 2.2),

[Product] = A(1 – exp(- kobst)) (Eq. 2.2)

where kobs is the observed reaction rate constant and A is the reaction amplitude. Next, the kobs values were plotted against the dNTP concentrations and the plot was then fit to a hyperbolic equation (Equation 2.3),

kobs = kp[dNTP]/([dNTP] + Kd, dNTP) (Eq. 2.3)

where kp is the maximum dNTP incorporation rate constant and Kd, dNTP is the equilibrium dissociation constant for the binding of dNTP to the binary complex (Dpo4•DNA) to form the ternary complex (Dpo4•DNA•dNTP).

2.2.6 Biphasic Kinetic Assays

Dpo4 (120 nM) and 5-32P-labeled DNA (30 nM) were first preincubated in buffer

R and the solution was then mixed rapidly with a solution containing both 5 µM DNA trap D-1 (Table 2.1) and 1.2 mM correct dNTP in buffer R by using a rapid chemical- quench flow apparatus. The reactions were quenched at specific times by the addition of

EDTA to 0.37 M. The samples were resolved and quantitated as described above. The product formation was plotted versus time and the plot was fit to a double-exponential

28 equation (Equation 2.4),

[Product] = Af[1 – exp(- kft)] + As[1 – exp(- kst)] (Eq. 2.4)

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

2.3 Results

2.3.1 Bypass of a Site-Specifically Placed dGC8-N-ABA Lesion by Dpo4

Running start assays were employed to examine the response of Dpo4 to a site specifically placed dGC8-N-ABA lesion in the 26-mer-dGC8-N-ABA template relative to the corresponding undamaged 26-mer template (Table 2.1). Dpo4 gradually elongated the

17-mer primer and synthesized the full-length product 26-mer in about 8 s with the control 17/26-mer DNA substrate, whereas it took about 900 s to achieve the same result with the damaged 17/26-mer-dGC8-N-ABA DNA substrate (Figure 2.1). Thus, a single dGC8-N-ABA lesion slowed overall Dpo4-catalyzed DNA polymerization by more than 100-

bypass fold. To quantitatively study the polymerase pause sites, the t50 , defined as the time required to bypass 50% of the total dGC8-N-ABA lesions encountered by Dpo4, was estimated in Figure 2.2 (see Experimental Procedures). Consistent with the observations

bypass control in Figure 2.1, the t50 value was 30 s which is 270-fold greater than the t50 value of 0.11 s for the time to traverse 50% of undamaged dG encountered by Dpo4 at the same template position. Furthermore, the time required to extend 50% of the 21-mer lesion

extension bypass product encountered by Dpo4 (t50 ) was estimated to be 600 s (Figure 2.2),

extension which is 320-fold greater than the t50 value of 1.85 s for the time to traverse 50% of the same position in the undamaged template. As a result, relative to replication of the undamaged DNA substrate, Figure 2.1B shows the strong accumulation of intermediate products of 20-mer and 21-mer for replication of the damaged template, indicating that

Dpo4 paused strongly during the bypass of the lesion and at the subsequent extension

extension bypass step. In summary, the 20-fold greater t50 versus t50 (Figure 2.2) indicates that

Dpo4 pauses significantly more while extending the bypass product than while bypassing the lesion. A similar Dpo4 pausing pattern has been observed previously with another bulky lesion dGAP , as well as smaller lesions such as abasic sites and double base lesions.49,51,52 Interestingly such a pausing pattern is not observed with 8-oxo-dG which is bypassed efficiently by Dpo4.45 Besides the accumulation of the 20-mer and 21-mer products in Figure 2.1B, the 25-mer intermediate product also accumulated in the presence of both control and damaged DNA substrates (Figure 2.1). The accumulation was caused by “polymerase slippage” via primer realignment against the 26-mer template which contains three consecutive dCs at its 5-terminus (Table 2.1).52 Lastly, a small amount of 27-mer was synthesized by Dpo4 with the control DNA substrate 17/26-mer, resulting from a Dpo4-catalyzed blunt-end addition to the full-length 26-mer product.79

2.3.2 Moderate Effect of a dGC8-N-ABA Lesion on DNA Binding by Dpo4

The significant impact of dGC8-N-ABA on the rate of DNA synthesis (Figure 2.1B) may be attributed to an adverse effect on DNA binding by Dpo4. To examine this possibility, EMSAs were employed to measure the binding affinities of Dpo4 to several undamaged and damaged DNA substrates. As an example, Figure 2.3 shows how the

30 binding affinity between Dpo4 and [32P]-labeled 21/26-mer-dGC8-N-ABA was measured.

First, 5-32P-labeled 21/26-mer-dGC8-N-ABA (5 nM) was titrated with varying concentrations of Dpo4 and the resulting binary complex (Dpo4•21/26-mer-dGC8-N-ABA) was separated from free 21/26-mer-dGC8-N-ABA via native PAGE (Figure 2.3A). The titration was repeated four times. After quantitation, the concentration of Dpo4•21/26- mer-dGC8-N-ABA was plotted against the total concentration of Dpo4 and the plot was then fit to Eq. 2.1 (“Experimental Procedures”) to obtain a Kd, DNA of 29 ± 7 nM (Figure

2.3B). Based on measured Kd, DNA values in Table 2.2, Dpo4 bound to the undamaged

DNA substrates with Kd, DNA values ranging from 3.1 to 4.0 nM. In comparison, the single dGC8-N-ABA lesion weakened the binding of the damaged DNA substrates to Dpo4 with affinities in the range of 5.0 to 29 nM. Notably, the affinity ratio (Table 2.2) is greater with 20/26-mer-dGC8-N-ABA (3-fold) and 21/26-mer-dGC8-N-ABA (7.8-fold). However, we concluded that the impact of the lesion on the DNA binding affinity of Dpo4 contributed but was not sufficient to explain the strong accumulation of the 20-mer and 21-mer in

Figure 2.1B.

C8-N-ABA 2.3.3 Kinetics of dNTP Incorporation in the presence of a dG lesion

Other than the DNA binding affinity, the dGC8-N-ABA lesion may affect the kinetics of nucleotide incorporation opposite the lesion and/or at adjacent template positions. To test this scenario, we determined the maximum dNTP incorporation rate constant kp, the equilibrium dissociation constant of dNTP binding Kd, dNTP, and the substrate specificity

(or efficiency) kp/Kd,dNTP for a single dNTP incorporation into undamaged or damaged

DNA substrates with primers 18-mer to 22-mer under single turnover reaction conditions.

As a representative example, Figure 2.4 shows the measurement of the kp and Kd, dNTP values for dATP incorporation into 18/26-mer-dGC8-N-ABA. First, a preincubated solution of Dpo4 (120 nM) and 5ʹ-32P-labeled 18/26-mer-dGC8-N-ABA (30 nM) was rapidly mixed and reacted with varying concentrations of dATP and quenched at various time points.

The plot of product concentrations versus reaction times at each dATP concentration was fit to Eq. 2.2 (“Experimental Procedures”) to yield the observed rate constant, kobs. The kobs values were then plotted against dATP concentrations and the plot was fit to Eq. 2.3

-1 (“Experimental Procedures”) to produce a kp of 2.0 ± 0.4 s and a Kd, dATP of 2,200 ± 600

μM (Table 2.3). With varying sizes of the primer, we similarly measured the kinetic parameters for correct or incorrect nucleotide incorporation opposite dGC8-N-ABA and at the template positions upstream and downstream from the lesion (Table 2.3). The

C8-N-ABA kp/Kd,dNTP values for correct dNTP incorporations into 18/26-mer-dG , 19/26-mer- dGC8-N-ABA , and 22/26-mer-dGC8-N-ABA are either similar to or less than 10-fold lower than the efficiency values with the corresponding control DNA substrates. The efficiency values are 120- and 170-fold lower for correct dNTP incorporation into 20/26-mer-dGC8-

N-ABA and 21/26-mer-dGC8-N-ABA than into the control 20/26-mer and 21/26-mer substrates, respectively (Table 2.3, Table 2.4). Thus, relative to undamaged DNA, Dpo4 had little difficulty elongating damaged DNA at non-pause sites (18/26-mer-dGC8-N-ABA,

19/26-mer-dGC8-N-ABA , and 22/26-mer-dGC8-N-ABA) but struggled at the two strong pause sites (20/26-mer-dGC8-N-ABA and 21/26-mer-dGC8-N-ABA), leading to the accumulation of

20-mer and 21-mer observed in Figure 2.1B. Furthermore, with damaged DNA substrates containing a single site specifically placed dGC8-N-ABA lesion, we found that

Dpo4 incorporated correct over incorrect nucleotides with 10- to 10,000-fold greater efficiency, resulting in polymerase fidelity in the range of 10-1 to 10-6 (Table 2.3). At the non-pause sites, the polymerase fidelity (10-3 to 10-6) is similar to the fidelity measured with control DNA.77 In comparison, the polymerase fidelity (10-1 to 10-3) at the pause sites is 100-fold lower on average. Consequently, the probability of correct dNTP incorporation was above 98% at all sites tested in the damaged template, except with

21/26-mer-dGC8-N-ABA, where the probability of correct nucleotide incorporation fell to

84.3% (Table 2.3).

2.3.4 Biphasic Kinetics of dNTP Incorporation at Polymerase Pause Sites

Our previous kinetic studies have shown that correct dNTP incorporation follows biphasic kinetics at each polymerase pause site induced by various DNA lesions.49,51-53

To examine if the kinetic trend is consistent at the two Dpo4 pause sites caused by dGC8-

N-ABA, we performed a DNA trap experiment. A preincubated solution of Dpo4 (120 nM) and 5-32P-labeled 20/26-mer-dGC8-N-ABA or 21/26-mer-dGC8-N-ABA (30 nM) was rapidly mixed with a solution containing correct dNTP (1.2 mM) and unlabeled D-1 DNA (5

µM, Table 2.1). The reaction was quenched at various times by addition of EDTA to

0.37 M. Previously, we have shown that a 167-fold molar excess of undamaged D-1 over

5-32P-labeled damaged DNA was sufficient to trap any free Dpo4 before its rebinding to the damaged DNA substrate.51,52 As expected, biphasic kinetics of dNTP incorporation was observed with both damaged DNA substrates. Each plot of product concentration versus reaction time (Figure 2.5) was fit to a double exponential equation (Eq. 2.4), yielding the biphasic kinetic parameters listed in Table 2.5. In comparison, the data fit

33 poorly to a single exponential equation (Data not shown). This can be attributed to the fact that with either 20/26-mer-dGC8-N-ABA or 21/26-mer-dGC8-N-ABA, the first phase rate constant (kf) is substantially greater than the second phase rate constant (ks) while the reaction amplitudes in the first phase (Af) and the second phase (As) are comparable. The

C8-N-ABA total reaction amplitudes (Af + As) were 44% with 20/26-mer-dG and 14.3% with

21/26-mer-dGC8-N-ABA, indicating that a substantial amount of damaged DNA was not converted into products during the DNA trap assays. This low overall reaction amplitude was possibly caused by the lesion and/or the reaction conditions. To exclude the latter possibility, similar DNA trap assays were previously performed with the undamaged

20/26-mer and 21/26-mer substrates.52 Only a single fast phase of nucleotide incorporation was observed with the control substrates.52 The reaction amplitudes with both 20/26-mer and 21/26-mer (Table 2.5) are ~ 90%,52 demonstrating that the presence of dGC8-N-ABA significantly lowered overall reaction amplitudes at the polymerase pause sites. Furthermore, to examine if the biphasic kinetics of nucleotide incorporation onto damaged DNA were affected by dNTP concentration, the same DNA trap assay with

20/26-mer-dGC8-N-ABA was performed under 450 or 1,000 µM dCTP (Figure 2.5A) and the kinetic data were listed in Table 2.5. Notably, the biphasic data with 450, 1,000, and

1,200 µM dCTP (Table 2.5) reveal that the reaction amplitudes of the fast and slow phases were unchanged while their rate constants increased with higher dCTP concentrations.

2.4 Discussion

2.4.1 Polymerase Pause Sites

During genomic replication, a bulky lesion like dGC8-N-ABA is expected to act as a road block for high-fidelity replicative DNA polymerases, which are known to have a tight active site and select against DNA base pairs with non-canonical or distorted geometry.195 For low-fidelity, Y-family DNA polymerases like Dpo4, their loose and flexible active site tolerates damaged DNA which is geometrically distorted by various lesions as demonstrated by the ternary crystal structures of Dpo4, an incoming nucleotide, and DNA containing various lesions.82-84,87,89-91,93,196 Our running start assays show that Dpo4 was indeed capable of bypassing the site-specifically placed dGC8-N-ABA lesion and synthesized full-length products albeit at a significantly slower rate than copying a corresponding undamaged DNA template (Figure 2.1). The slower full-length product formation rate is primarily due to two consecutive strong polymerase pause sites evidenced by the accumulation of the 20-mer and 21-mer intermediate products (Figure

2.1B). A similar polymerase pausing pattern was observed in our published kinetic studies of the bypass of dGAP.52 During polymerization, Dpo4 paused when incorporating a nucleotide opposite the dGC8-N-ABA lesion, and even more so while extending the bypass product. These conclusions are consistent with the 270- and 320-

st nd fold longer t50 values for Dpo4 to traverse the 21 and 22 positions, respectively, from the 3-terminus of the damaged template 26-mer-dGC8-N-ABA than the corresponding positions on the control template 26-mer.

2.4.2 Kinetic Basis for Polymerase Pausing as a Result of dGC8-N-ABA

Our DNA-binding assays revealed that the dGC8-N-ABA lesion only reduced the binding affinity of Dpo4 to the damaged DNA substrates by up to 7.8-fold when

35 compared to the corresponding undamaged DNA substrates (Figure 2.6D), and Dpo4 binds to the damaged DNA substrates at non-pause sites with 2-6 fold higher affinities than at pause sites (Table 2.2). Although these affinity differences contribute to the strong pausing of Dpo4 at the two template positions (Figure 2.1B) based on the large t50 differences, a reduction in DNA-binding affinity alone is not sufficient to induce such significant pausing by Dpo4. To further investigate the kinetic basis behind the bypass of dGC8-N-ABA catalyzed by Dpo4, individual nucleotide incorporation efficiencies at five template positions were measured with either damaged or undamaged DNA substrates

(Table 2.1). Our kinetic data (Table 2.3) demonstrated that the dGC8-N-ABA lesion did not alter the kinetic trend that correct nucleotides were incorporated with significantly higher rate constants (kp) and efficiencies (kp/Kd,dNTP) than incorrect nucleotides at all template positions. For the three series reactions during polymerization including 18-mer19- mer20-mer, 19-mer20-mer21-mer, and 20-mer21-mer22-mer, they share the following kinetic pattern based on the kinetic data in Table 2.3: the first conversion step

nd has higher kp and kp/Kd,dNTP values for correct nucleotide incorporation than the 2 conversion step, leading to the accumulation of the intermediates 19-mer, 20-mer, and

21-mer, respectively. The reason why the 19-mer accumulated only at early time points is because the fast and efficient conversion of 19-mer20-mer (0.48 s-1, 9.6 x 10-3 μM-1s-1) was completed at later time points (Figure 2.1B). In contrast, the strong accumulation of

21-mer product is due to large differences (22-28 fold) in both the rate constants and efficiencies in the conversion of 21-mer22-mer (0.0089 s-1, 2.2 x 10-5 μM-1s-1) relative to the conversion of 20-mer21-mer (0.25 s-1, 4.9 x 10-4 μM-1s-1). Similarly, a strong

36 accumulation of 20-mer was contributed by the slower and less efficient conversion of

20-mer21-mer (0.25 s-1, 4.9 x 10-4 μM-1s-1) than the conversion of 19-mer20-mer

(0.48 s-1, 9.6 x 10-3 μM-1s-1).

Notably, the 22-mer intermediate product did not accumulate (Figure 2.1B), despite that the conversion of 22-mer23-mer (3.2x10-3 μM-1s-1) has 3-fold lower efficiency than the conversion of 19-mer20-mer (9.6 x 10-3 μM-1s-1) and the correct nucleotide incorporation efficiency ratio is 11-fold higher at this template position

(Figure 2.6A). The lack of 22-mer accumulation was due to the kinetic pattern in the reaction series 21-mer22-mer23-mer, whereas the 2nd step (4.5 s-1) is much faster than the first step (8.9 x 10-3 s-1). This kinetic pattern is opposite from the pattern deduced from the kinetic series reactions discussed above. Notably, these kinetic patterns for intermediate product formation have been previously revealed in the Dpo4 catalyzed bypass of an abasic site,49 a cisplatin-d(GpG) adduct,51 and a dGAP 52.

2.4.3 Large Effect of dGC8-N-ABA on the Kinetic Parameters of dNTP Incorporation

When comparing the kinetic parameters obtained with control and damaged templates, the correct nucleotide incorporation efficiency ratio,

(kp/Kd,dNTP)control/(kp/Kd,dNTP)damaged, increases dramatically from non-pause to pause sites

(Figure 2.6C). For example, the ratios (Table 2.3) are less than 10-fold at non-pause sites while the ratios are 120 and 170 with primers 20-mer and 21-mer, respectively.

Furthermore, the single dGC8-N-ABA lesion in 20/26-mer-dGC8-N-ABA and 21/26-mer-dGC8-N-

ABA weakened their binding to Dpo4 by 3.0- and 7.8-fold, respectively (Figure 2.6D and

Table 2.2). Thus, the impact of dGC8-N-ABA on Dpo4•DNA binding is significantly 37 smaller than on nucleotide incorporation at and near the lesion site. Since the overall efficiency of a polymerase at a template position is a function of both its binding affinity to DNA and nucleotide incorporation efficiency during processive polymerization, Dpo4 was 360- and 1,330-fold less efficient while elongating primers 20-mer and 21-mer, respectively, with the damaged template 26-mer-dGC8-N-ABA than with the control

C8-N- template 26-mer. Notably, the kp was 46- and 182-fold slower with 20/26-mer-dG

ABA and 21/26-mer-dGC8-N-ABA, respectively, than with the corresponding control substrates (Figure 2.6A), while the equilibrium dissociation constant ratio

(Kd,dNTP)damage/(Kd,dNTP)control was less than 2-fold (Figure 2.6B). Thus, the ratio of

(kp/Kd,dNTP)control/(kp/Kd,dNTP)damaged at each Dpo4 pause site (Figure 2.6C) is dictated largely by the ratio of (kp)control/(kp)damaged. Oddly, correct dCTP was incorporated with a

C8-N-ABA 10-fold greater Kd,dNTP in the presence of 22/26-mer-dG than in the presence of control 22/26-mer while the kp value was not altered by the lesion, leading to a nucleotide incorporation efficiency ratio of 10 at this non-pause site (Figure 2.6). It is unclear how an embedded template nucleotide dGC8-N-ABA weakened the ground-state binding of an incoming nucleotide to Dpo4•22/26-mer-dGC8-N-ABA. It is possible that the bulky dGC8-N-

ABA lesion is even more intrusive when further back in the polymerase active site, leading to the inability of Dpo4 to efficiently bind an incoming nucleotide.

2.4.4 Kinetic Mechanism for dGC8-N-ABA Bypass by Dpo4

Since the presence of dGC8-N-ABA did not significantly affect the binding of Dpo4 to both 20/26-mer-dGC8-N-ABA and 21/26-mer-dGC8-N-ABA, in comparison to the respective control DNA substrates, it is likely that Dpo4 holds onto the damaged DNA substrates

38 when incorporating a correct incoming dNTP with low kp values. This possibility was confirmed by the DNA trap assays which demonstrated that some of the Dpo4•DNA complex did not dissociate even in the slow reaction phase (Figure 2.5). The biphasic kinetics of nucleotide incorporation at each pause site show that a fast phase (Af, kf) preceded a slow phase (As, ks) (Figure 2.5 and Table 2.5). In both cases, the sum of the contribution from the fast (Afkf) and slow (Asks) phases yielded rate constants with 20/26- mer-dGC8-N-ABA (0.18 s-1) and 21/26-mer-dGC8-N-ABA (0.009 s-1) which were relatively

-1 close to the respective observed kobs values of 0.18 and 0.007 s , estimated using

Equation 2.3, 1.2 mM dCTP in Figure 2.5, and corresponding measured Kd,dNTP and kp values (Table 2.3). In comparison, similar DNA trap assays with the control 20/26-mer and 21/26-mer substrates only exhibited single, fast phase kinetics with greater reaction amplitudes of 87% and 90%, respectively (Table 2.5). The less than 100% reaction amplitudes with the control substrates can be attributed to experimental errors, inaccurate measurements of oligomer concentrations, incomplete or imperfect annealing of DNA duplexes, or/and Dpo4 binding at the blunt end rather than the staggered end of DNA.79

Together, the kinetic data in the presence of a DNA trap suggest that the slow phase observed with the damaged DNA substrates was due to the formation of nonproductive

N complexes between Dpo4 and DNA (E•DNAn ) while the fast phase exhibited by each control or damaged DNA substrate indicates the formation of a productive complex

P (E•DNAn ), which was turned over rapidly once a nucleotide was bound. Notably, the kf values are smaller with the damaged DNA substrates than with the control DNA substrates, suggesting that even in the fast phase, Dpo4 did not bind to damaged DNA as

39 productively as it interacted with undamaged DNA. Furthermore, since the slow

N elongation of E•DNAn occurred in the presence of a large molar excess of unlabeled

N P DNA trap, E•DNAn was first transformed into E•DNAn without dissociation with a first order rate of ke, and subsequently elongated (Scheme 2.2A). Interestingly, the mechanism in Scheme 2.2A has been implicated into the bypass of an abasic site,49 a cisplatin-d(GpG) adduct,51 and a dGAP lesion by Dpo4.52 If the mechanism in Scheme

2.2A is correct at each of the two polymerase pause sites in Figure 2.1B, the slow phase rate constant (ks) which is dominated by ke, should not be altered when the dNTP concentration is changed. To test this possibility, we performed the same DNA trap assays with 20/26-mer-dGC8-N-ABA in the presence of different dCTP concentrations. We did not perform such DNA trap experiments with 21/26-mer-dGC8-N-ABA because the overall reaction amplitude (4.3 nM) with this DNA substrate (Table 2.5) is small and the limited accuracy of the kinetic data may not allow us to draw a definite conclusion. With

20/26-mer-dGC8-N-ABA, we performed the DNA trap assays with dCTP concentration of

450, 1,000, and 1,200 M (Figure 2.5A). The biphasic kinetic data in Table 2.5 indicate that the reaction amplitudes (Af and As) do not change with the variation of correct dCTP concentration, which is expected since dNTP concentration should not affect the binding of DNA and Dpo4. In contrast, both the fast and slow phase rate constants are 2- to 3-fold higher in the presence of 1,200 than 450 M of dCTP while the rate constants with 1,000

M dCTP are in the middle. These biphasic kinetic data suggest that the kinetic mechanism in Scheme 2.2A is inaccurate for the bypass of dGC8-N-ABA by Dpo4 and prompted us to propose a new one shown in Scheme 2.2B. In this mechanism, while the

P fast phase kf is still governed by the process of nucleotide incorporation onto E•DNAn as in Scheme 2.2A, the slow phase rate constant (ks) is a function of both the process of

N slow nucleotide incorporation onto E•DNAn (k2) and the interconversion between the non-productive and productive binary complexes (ke). Additionally, our kinetic data in

Table 2.5 also suggest that ke cannot be much larger than k2. Otherwise, Scheme 2.2B will be simplified as Scheme 2.2A.

C8-N-ABA Lastly, the sum of Af and As with either 20/26-mer-dG (44%) or 21/26- mer-dGC8-N-ABA (14.3%) is significantly smaller than the reaction amplitude (~90%) observed with either 20/26-mer or 21/26-mer (Table 2.5). Therefore, significant percentages of 20/26-mer-dGC8-N-ABA (90% - 44% = 46%) and 21/26-mer-dGC8-N-ABA

D (90% - 14% = 76%) bound by Dpo4 were either catalytically incompetent (E•DNAn ) or dissociated during the trap assays if the maximum reaction amplitudes with the two damaged substrates can be as high as their corresponding control substrates (~90%). As

N P the formation of the ternary complexes E•DNAn •dNTP and E•DNAn •dNTP, dNTP

D D binds to E•DNAn and yields E•DNAn •dNTP. Previously, such ternary complexes

D (E•DNAn •dNTP) have been proposed as Dpo4, HIV-1 reverse transcriptase, and T7

DNA polymerase bind to damaged DNA containing dGAP,52 N2-methylguanine,197 O6- benzylguanine,198 and O6-methylguanine.198 Taken together, the above analysis suggests the kinetic mechanism in Scheme 2.2B, rather than in Scheme 2.2A, is accurate for the bypass of dGC8-N-ABA by Dpo4.

Although there are no reported crystal structures of Dpo4 in complex with DNA

41 containing dGC8-N-ABA, several published structures of the Y-family polymerases and

DNA containing similarly bulky lesions support the lesion bypass mechanisms (Scheme

2.2).83,84,199 For example, the binary crystal structures of yeast Polη and DNA containing a damaged templating nucleotide N-2-acetylaminofluorene-dG (AAF-dG) 199 show that the AAF moiety either stacks above the junction base pair, or partially rotates out of the

DNA helix but still blocks incoming dCTP. The first conformation of AAF completely

D blocks an incoming dNTP and thereby represents E•DNAn . The second conformation of

N AAF likely represents E•DNAn , which requires subtle to mild structural changes in order for AAF to be completely rotated out of the DNA helix. If the AAF moiety is completely out of the DNA duplex, the binary complex is considered to be in the form of

P E•DNAn , which productively binds an incoming nucleotide and rapidly incorporates it.

P Interestingly, such a productive conformation (E•DNAn •dNTP) has been displayed by the ternary structures of Dpo4, dNTP, and DNA containing either benzo[a]pyrene diol epoxide (BPDE)-dG or BPDE-dA,83,84 illustrating that the BPDE moiety is flipped out of the DNA helix into a structural gap between the Little Finger and Palm domains of Dpo4 and the distance between the primer 3′-OH and the α–phosphate of the incoming dNTP

(3.9 Å) is close to the optimum catalytic distance (3.4 Å).83,200 Although these binary and ternary structures with AAF-dG199 and BPDE83,84 are instructive, it remains to see how

DNA containing dGC8-N-ABA binds to Dpo4 and forms the three conformations of

D N P E•DNAn , E•DNAn , and E•DNAn .

2.4.4 Biological relevance of the kinetic studies of dGC8-N-ABA

Figure 2.1B shows that a single dGC8-N-ABA considerably stalls DNA replication 42 catalyzed by Dpo4 in vitro. Consistently, dGC8-N-ABA is recently shown to be the strongest replication block of the three major 3-NBA derived DNA adducts in an assay with nucleotide excision repair-deficient human XPA cells.201 Furthermore, the fidelity values in Table 2.3 indicate that Dpo4 is prone to misincorporate nucleotides at the two pause sites (Figure 2.1B). The error-prone manner is especially severe during the extension of the lesion bypass product, where Dpo4 misincoporates dCTP, dATP, and dTTP opposite dC with high frequencies of 10.3, 3.8, and 1.6%, respectively. Notably, dGC8-N-ABA, inducing primarily G-to-A transitions followed by G-to-T transversions, is found to be the most mutagenic 3-NBA adduct with a mutational frequency of 30.6% in the nucleotide excision repair-deficient human XPA cells.201 The cellular results agree with our in vitro fidelity values although it remains to determine the identity of the polymerase which bypasses dGC8-N-ABA in the XPA cells.

Author contributions

Varun V. Gadkari was responsible for planning and executing a majority of the kinetic experiments as well as analyzing the results. V.V.G also wrote the initial draft of the manuscript. E. John Tokarsky helped conduct and analyze some of the kinetic experiments with guidance from V.V.G. Chanchal Malik synthesized the 26-mer containing the 3-NBA lesion. Dr. Zucai Suo conceived the research project and kinetic mechanisms and rewrote the manuscript. The research was supported by the National

Institutes of Health grant ES009127 to Dr. Zucai Suo and Dr. Ashis Basu.

2.5 Schemes

Scheme 2.1: Bioactivation of 3-Nitrobenzanthrone to form N-(deoxyguanosin-8-yl)- 3-aminobenzanthrone (dGC8-N-ABA)

Scheme 2.2: Proposed kinetic mechanisms of DNA lesion bypass

2.6 Tables

Table 2.1: DNA Substrates Primers (positiona) Sequences

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' b26-mer-dGC8-N-ABA 3'-TTGCTGCCGGTCACTTAAGCGCGCCC-5'

DNA trap

5'-CGCAGCCGTCCAACCAACTCA-3'

D-1 (21/41-mer) 3'-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG- 5' aPosition of primer terminus relative to DNA adduct bG designates dGC8-N-ABA.

Table 2.2: Binding Affinity of Dpo4 to Damaged and Control DNA Substrates

DNA Substrate Damaged DNAa Control DNAb,c Affinity Ratiod

nM nM

19/26-mer 7.4 ± 0.8 3.1 ± 0.5 2.4

20/26-mer 12 ± 3 4.0 ± 0.2 3.0

21/26-mer 29 ± 7 3.7 ± 0.2 7.8

22/26-mer 5.0 ± 0.9 3.8 ± 0.6 1.3 aRefer to those DNA substrates with the template 26-mer-dGC8-N-ABA bRefer to those undamaged DNA substrates with the control template 26-mer cValues are from Table 2 of reference52 d Calculated as (Kd, DNA)damaged/(Kd, DNA)control

Table 2.3: Kinetic parameters for dNTP incorporation into damaged DNA substrates containing dGC8-N-ABA

d e dNTP Kd,dNTP kp (kp/Kd, dNTP)damaged Efficiency Fidelity Probability -1 -1 -1 (μM) (s ) (μM s ) Ratioa,b,c

- dCTP 32 ± 5 2.83 ± 0.08 0.088 1.25 - 99.95

C8 dG

-2 -6 -6

- dATP 2200 ± 600 (2.0 ± 0.4) x 10 9.1 x 10 0.85 7.3 x 10 0.01

ABA

mer N - dGTP 350 ± 40 (6.2 ± 0.3) x 10-3 1.8 x 10-5 1.6 1.4 x 10-5 0.02

8/26 -5 -5 1 dTTP - - 1.6 x 10 5.4 1.3 x 10 0.02

- dGTP 50 ± 9 0.48 ± 0.02 9.6 x 10-3 2.6 - 99.82

C8 dG

-3 -6 -4

- dATP 430 ± 60 (2.9 ± 0.2) x 10 6.7 x 10 2.1 7.0 x 10 0.07

ABA

- mer N -3 -6 -4 - dCTP 220 ± 80 (1.4 ± 0.2) x 10 6.4 x 10 22 6.7 x 10 0.06

-6 -4 19/26 dTTP - - 5.5 x 10 1.6 5.7 x 10 0.05

dCTP 510 ± 60 0.25 ± 0.02 4.9 x 10-4 120 - 98.93

-3 -6 -3 ABA

- dATP 640 ± 170 (1.9 ± 0.3) x 10 3.0 x 10 6.3 6.1 x 10 0.61

mer

N -

C8 dGTP 1200 ± 400 (8.1 ± 0.1) x 10-4 6.8 x 10-7 66 1.4 x 10-3 0.14

20/26

dG * dTTP 970 ± 390 (1.6 ± 0.4) x 10-3 1.6 x 10-6 52 3.3 x 10-3 0.32

dGTP 400 ± 80 (8.9 ± 0.7) x 10-3 2.2 x 10-5 170 - 84.26

-4 -6 -2 ABA

- dATP 450 ± 50 (4.5 ± 0.2) x 10 1.0 x 10 2.4 4.3 x 10 3.83

mer

N -

C8 dCTP 160 ± 20 (4.3 ± 0.1) x 10-4 2.7 x 10-6 12 1.1 x 10-1 10.34

21/26

dG * dTTP 420 ± 80 (1.7 ± 0.1) x 10-4 4.0 x 10-7 4.5 1.8 x 10-2 1.57

- dCTP 1400 ± 300 4.5 ± 0.7 3.2 x 10-3 11 - 99.55

C8 dG

-3 -5 -3

- dATP 470 ± 70 (5.4 ± 0.4) x 10 1.1 x 10 0.38 3.4 x 10 0.34

ABA

- mer N -3 -6 -4 - dGTP 1100 ± 200 (1.4 ± 0.2) x 10 1.3 x 10 6.5 4.1 x 10 0.04

-3 -6 -4 22/26 dTTP 770 ± 140 (1.7 ± 0.2) x 10 2.2 x 10 5.0 6.9 x 10 0.07

*Denote pause sites.

a Calculated as (kp/Kd, dNTP)control/(kp/Kd, dNTP)damaged.

b 52 Values for (kp/Kd, dNTP)control 19/26-mer to 22/26-mer are from supplemental Table 1 of reference .

c Values for (kp/Kd, dNTP)control 18/26-mer are listed in Table 2.4.

d Calculated as (kp/Kd, incorrect dNTP)damaged/((kp/Kd, correct dNTP)damaged + (kp/Kd, incorrect dNTP)damaged). e Calculated as ((kp/Kd, dNTP)damaged/(Σ(kp/Kd, dNTP)damaged)) x 100.

Table 2.4: Kinetic parameters for dNTP incorporation into a control DNA substrate 18/26-mer.

a dNTP Kd,dNTP kp (kp/Kd, dNTP)control Fidelity

(μM) (s-1) (μM-1s-1)

dCTP 22 ± 3 2.4 ± 0.1 0.11 -

dATP 1600 ± 500 (1.2 ± 0.3) x 10-2 7.7 x 10-6 7.0 x 10-5

dGTP 180 ± 10 (5.0 ± 0.1) x 10-3 2.8 x10-5 2.5 x 10-4

dTTP 610 ± 40 (5.2 ± 0.2) x 10-3 8.6 x 10-5 7.8 x 10-4

a Calculated as (kp/Kd, incorrect dNTP)damaged/((kp/Kd,correct dNTP)damaged + (kp/Kd, incorrect dNTP)damaged).

Table 2.5: Biphasic kinetic parameters for correct dNTP incorporation into DNA Substrate in the presence of DNA trap.

a a DNA substrate Correct dNTP Af kf As ks

(nM) (s-1) (nM) (s-1)

Nucleotide concentration = 1,200 M b20/26-mer dCTP 26 ± 1 (86.7%) 4.6 ± 0.5 - - b21/26-mer dGTP 27.0 ± 0.4 (90%) 2.3 ± 0.1 - -

20/26-mer-dGC8-N-ABA dCTP 8.6 ± 0.7 (29%) 0.60 ± 0.07 4.5 ± 0.7 (15%) 0.05 ± 0.01

21/26-mer-dGC8-N-ABA dGTP 2.2 ± 0.4 (7.3%) 0.12 ± 0.06 2.1 ± 0.5 (7%) 0.0028 ± 0.0025

Nucleotide concentration = 1,000 M

20/26-mer-dGC8-N-ABA dCTP 8.6 ± 1.7 (29%) 0.32 ± 0.09 4.7 ± 1.7 (16%) 0.04 ± 0.01

Nucleotide concentration = 450 M

20/26-mer-dGC8-N-ABA dCTP 8.7 ± 0.2 (29%) 0.23 ± 0.03 4.7 ± 0.6 (16%) 0.021 ± 0.005 aPercentage after each reaction amplitude is calculated as (reaction amplitude/30 nM) x 100%. bReaction amplitude (A) and rate constant (k) values are from Table 4 of reference52

2.7 Figures

Figure 2.1: Running start assay A preincubated solution of 100 nM Dpo4 and 100 nM 5ʹ-32P-labeled (A) 17/26-mer, or (B) 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 3ʹ-terminus of the DNA template.

bypass extension Figure 2.2: Determination of t50 and t50 The percentages of the extension of the intermediate products 20-mer/26-mer-dGC8-N-ABA (●) and

21-mer/26-mer-dGC8-N-ABA (■) out of all polymerase encounter events were determined by quantitation of the gels in Figure 2.1 and then plotted against reaction time.

A B

Figure 2.3: Determination of Kd,DNA (A) A gel image shown the formation of the binary complex Dpo4•DNA during titration. Dpo4

(0 – 220 nM) was titrated into a solution containing 5ʹ-32P-labeled 21/26-mer-dGC8-N-ABA (5 nM).

The binary complex of Dpo4•DNA was separated from free DNA by native PAGE. The titration was repeated four times. (B) Plot of the binary complex concentration versus the total concentration of Dpo4. Each error bar represents the standard deviation of a complex concentration based on four independent experiments. The data were fit to Equation 2.1 which yielded a Kd, DNA of 29 ± 7 nM.

A B

Figure 2.4: Kinetics of dATP incorporation onto 18/26-mer-dGC8-N-ABA (A) Dpo4 (120 nM) was first preincubated with 5ʹ-32P-labeled 18/26-mer-dGC8-N-ABA (30 nM), and was then rapidly mixed with increasing concentrations of dATP (50 μM, ●; 150 μM, ■; 300 μM,

♦; 500 μM, ▲; 700 μM, ●; 1000 μM, ■; 1600 μM, ♦) for the indicated times before being quenched. Each time course was fitted to Equation 2.2 to yield a kobs. (B) the kobs values were plotted against corresponding dATP concentrations and the plot was fit to Equation 2.3 to

-1 produce a kp of 2.0 ± 0.4 s and a Kd, dATP of 2,200 ± 600 μM.

A B

Figure 2.5: Biphasic kinetics of correct dNTP incorporation in the presence of a DNA trap A preincubated solution of 120 nM Dpo4 and 30 nM 5ʹ-32P-labeled 20/26-mer-dGC8-N-ABA (A) or

21/26-mer- dGC8-N-ABA (B) was mixed rapidly with 5 μM unlabeled 21/41-mer D-1 (DNA trap) and correct dNTP for various times before being quenched by EDTA. The plots of product versus time were fit to Eq. 2.4 and the data were listed in Table 2.5. (A) Dpo4•20/26-mer-dGC8-N-ABA was mixed with the DNA trap and 450 (Blue), 1,000 (Red), or 1,200 μM (Black) dCTP. The inset is a magnification of the time points from 0 to 20 seconds, showing the first phase of product formation. (B) Dpo4•21/26-mer-dGC8-N-ABA was mixed with the DNA trap and 1,200 μM dGTP.

A B

C D

Figure 2.6: Quantitative effects of a dGC8-N-ABA lesion on DNA binding and correct dNTP incorporation catalyzed by Dpo4

The ratios of the kinetic parameters including kp (A), Kd, dNTP (B), kp/Kd, dNTP (C), and Kd, DNA (D) between damaged and corresponding control DNA substrates were plotted against the primer sizes.

Chapter 3 Single-Molecule Investigation of Response to Oxidative DNA Damage by

a Y-Family DNA Polymerase

3.1 Introduction

One of the most common sources of endogenous DNA damage is aerobic respiration. This essential life process generates oxygen radicals, which are known to cause damage to DNA. For example, 8-oxo-7,8-dihydro-2ʹ-deoxyguanine (8-oxo-dG), a major oxidative DNA lesion, is formed by the oxidation of the C8 atom of guanine.

While structural studies have shown that 8-oxo-dG does not significantly distort the local structure of DNA,158-160 the lesion remains particularly mutagenic by often being better accommodated in a polymerase active site following rotation about the N-glycosidic bond to adopt the unusual syn-conformation. While the anti-conformation of 8-oxo-dG forms a correct Watson-Crick base pair with correct dCTP, the syn-conformation readily forms

Hoogsteen interactions with incorrect dATP.45,161,202 Unique interactions with 8-oxo-dG within the polymerase active site can either promote or limit misincorporation events.92,161-164,203 If left unrepaired, 8-oxo-dG:dA mispairs will result in G→T transversion mutations, which are implicated in cancer induction. 204,205

DNA lesions can also interfere with faithful DNA replication by stalling high- fidelity replicative DNA polymerases. While 8-oxo-dG does not completely block DNA synthesis, it does cause polymerase pausing in some instances.206-214 It is proposed that 57 this pausing initiates translesion DNA synthesis (TLS) via a polymerase switching mechanism that allows the cell to recruit Y-family DNA polymerases which are known for bypassing lesions in vitro and in vivo. Notably, the Y-family polymerases have been identified in all three domains of life, e.g. four in humans (DNA polymerases η, κ, ι, and

Rev1), two in Escherichia coli (DNA polymerase IV and V), and one in Sulfolobus solfataricus (DNA polymerase IV (Dpo4)).

Pre-steady-state kinetic and crystallographic data indicate that Dpo4 preferentially incorporates dCTP opposite 8-oxo-dG with high efficiency due to stabilization of the anti-conformation of the lesion in its active site.90,92,163,215 Recently, our real-time stopped-flow FRET studies have revealed how the individual domains of Dpo4 move during substrate binding and catalysis.69,71,75 Dpo4, akin to all canonical polymerases, contains three core polymerase domains denoted as Finger, Palm, and Thumb. Moreover, characteristic of all Y-family members, Dpo4 also contains an auxiliary domain termed the Little Finger (LF) which is connected to the Thumb domain by a highly flexible 14- amino acid residue peptide linker. The LF domain is known to impart unique DNA binding and lesion bypass capabilities to the Y-family polymerases.74 While crystallographic investigation suggests that upon nucleotide binding, no large-scale domain movements exist from the binary to ternary structures,68 our real-time stopped- flow FRET investigations illustrate global conformational changes in Dpo4 during nucleotide binding and incorporation onto undamaged DNA.69,71,75 These investigations permitted the expansion of the minimal kinetic mechanism for polymerase-catalyzed nucleotide incorporation.

Recently, single-molecule FRET (smFRET) methodologies have been employed to investigate the conformational dynamics of DNA polymerases and have illustrated how domain motions impact substrate recognition and selectivity.73,120,122-125,216

Specifically, the role of the Finger domain in nucleotide selectivity has been well characterized.120,124,125 While the Finger domain of Dpo4 has not been exclusively implicated in nucleotide selection, it is part of synchronized, global domain motions which likely play an important role in nucleotide recognition and selectivity.

Understanding the mechanism of nucleotide recognition and selectivity is of even greater importance since the Y-family DNA polymerases lack a proof-reading exonuclease domain. Here, our smFRET study revealed that Dpo4 existed in equilibrium among three

FRET states that likely represent distinct structural conformations of Dpo4 and its positions on the undamaged or 8-oxo-dG-containing DNA substrates. The addition of an incoming nucleotide affected the distribution of the FRET states and the DNA binding stability of the polymerase. Our study further established a mechanism by which Dpo4 can faithfully and efficiently bypass a major oxidative lesion.

3.2 Materials and Methods

3.2.1 Preparation of Protein and DNA

We selected a catalytically active mutant of Dpo4 used in previous work with mutations in the Finger domain (C31S, N70C) for the present study.69,71,75 The engineered Cys mutation allowed for site-specific fluorophore labelling (Figure 3.1B) which was carried out by incubation of Dpo4 with a 15-fold molar excess of Cy5- maleimide (GE Healthcare), overnight at 4˚C in a buffer containing 50 mM Tris (pH 7.2),

150 mM NaCl, 0.5 mM TCEP, and 10% glycerol. Unincorporated free dye was then removed with Micro Bio-Spin columns (Bio-Rad). By measuring the absorbance at 280 nm and 650 nm, the ratio of protein concentration to dye concentration revealed a labelling efficiency of 91% for the reaction.

The 21-mer primer containing a 5ʹ-biotin for surface immobilization as well as a

5-C6-amino-2ʹ-deoxythymidine modification at the 9th base from the 3ʹ terminus for Cy3-

NHS-ester labelling, and control undamaged template were purchased from Integrated

DNA Technologies. This labelling was performed according to the manufacturer’s protocol (GE Healthcare) (Figure 3.1A). The oligonucleotide containing the 8-oxo-dG base was purchased from Midland Certified Reagent Company, Incorporated. The primer and template oligonucleotides were annealed to each other by heating to 80˚C for

5 minutes followed by slow cooling to room temperature.

3.2.2 Steady-state Fluorescence Spectroscopy Assays

Fluorescence spectra were recorded on a Fluoromax-4 (HORIBA Jobin Yvon) at

20˚C. Increasing amounts of Cy5-labeled Dpo4 (0-180 nM) were titrated into a 25 nM solution of the Cy3-labeled DNA substrate (undamaged or damaged) in a buffer containing 25 mM HEPES (pH 7.5), 5 mM CaCl2, 25 mM NaCl and 10 % glycerol. The change in donor quantum yield was plotted against acceptor concentration. The resulting curve was fit to Equation 3.1 to yield the equilibrium dissociation constant for the interaction.

1/2 퐷푁퐴 퐷푁퐴 2 ∆휙 = (Δ휙푇/2퐷0)× {(퐾퐷 + 퐸0 + 퐷0) − 0.5 [(퐾퐷 + 퐸0 + 퐷0) − 4퐸0퐷0] }

(Eq. 3.1) 60 where Δ휙 is the change in the donor quantum yield, Δ휙푇 is the maximum change in donor quantum yield, 퐷0 is the total DNA concentration, 퐸0 is the total Dpo4

퐷푁퐴 concentration, and 퐾퐷 is the equilibrium dissociation constant.

3.2.3 Single-molecule Measurements

Single-molecule fluorescence experiments were conducted on a custom-built prism-type total internal reflection microscope (Eclipse Ts-i) with a 1.2 numerical aperture 60× water emersion objective (Nikon). Cy3 donor dyes were directly excited using a 532 nm laser (CrystaLaser, 100 mW) and the resulting emitted fluorescence was passed through a set of optics, including a high pass filter to reject scattered laser light

(Et5421P, Chroma) and dichroic mirrors (2x T6401p, Chroma) to separate donor signals from acceptor signals. Band pass filters were placed after the dichroic mirrors to minimize signal overlap between donor and acceptor channels (Et575/40m and

Et685/70m, Chroma). The FRET interactions were imaged in real time using an Andor iXon 897 electron-multiplying charge-coupled device camera to record movies at 10 frames per second over several minutes at 20 C. Imaging chambers were assembled from quartz slides and coverslips which were cleaned, passivated, biotinylated and coated with Neutravidin (0.2 mg/mL). The Cy3-labeled DNA was flowed into the imaging chamber at a concentration sufficient to observe single, distinct immobilized molecules

(20-40 pM). In addition to the Dpo4 binding buffer (50 mM HEPES (pH 7.6), 50 mM

NaCl, 0.1 mM EDTA, 5 mM CaCl2, and 0.1 mg/mL BSA), experiments were performed in an imaging solution containing an oxygen scavenging system (0.8% (w/v) d-glucose, 1 mg/mL glucose oxidase, and 0.04 mg/mL catalase) and 2 mM Trolox. Ca(II) was used as

61 the divalent metal ion in place of Mg(II) to prevent the incorporation of dNTPs. Ca(II) in the polymerase active site has been shown to closely mimic Mg(II), but prevents or dramatically reduces dNTP incorporation.87,88 Movies were recorded at 100 ms exposure time resolution, at 20˚C, over several minutes upon introduction of 10-20 nM Cy5- labeled Dpo4 to the imaging chamber. The movies were then processed using IDL (ITT

Visual Information Solutions) and analyzed using custom data acquisition and analysis software which included a MATLAB (MathWorks) script necessary for correcting anti- correlated donor and acceptor time trajectories for background (Center for the Physics of

Living Cells, University of Illinois at Urbana-Champaign).

3.2.4 Verification of the FRET System

To confidently correlate changes in FRET with molecular distance changes rather than unrelated photophysical artifacts, we performed several previously described control experiments.217 We recorded movies of surface immobilized Cy3-DNA following excitation at 532 nm in the presence and absence of WT Dpo4 to show that the donor intensity remains constant until photobleaching between the experiments (Figure 3.2).

Further, we recorded acceptor fluorescence upon direct excitation of 10 nM Cy5-Dpo4 with 638 nm laser light. A histogram composed of acceptor intensities from >100 time trajectories revealed one major distribution and, more notably, the absence of higher intensity distributions eliminating the possibility of multiple Dpo4 molecules binding per

DNA substrate (Figure 3.3).

3.2.5 Single Molecule Data Analysis

From the donor and acceptor time trajectories, apparent FRET (Eapp) was

퐼퐴 = 퐸푎푝푝 (Eq. 3.2) 퐼퐷+퐼퐴 calculated by Equation 3.2, where 퐼퐴 is the acceptor intensity and 퐼퐷 is the donor intensity. Notably, only fluorescence time trajectories with a single photobleaching event were selected for further processing in order to avoid ambiguity in removing background signal. FRET efficiency values from over 200 fluorescence time trajectories with acceptor intensities clearly above background level (as determined by the average background after donor photobleaching) were collected and binned from 0-1 to generate population histograms. The population distributions were fit to a sum of Gaussian - functions using MATLAB, and the percent density of molecules in each respective population was calculated as the total area under each peak. Each histogram was fit with the minimum number of peaks necessary to give a visually acceptable fit. The fit was generally deemed acceptable if the percent error, calculated as the total difference between the experimental bin heights and the computed value of the fit at each bin center, divided by the sum of the bin heights, was less than 10%. However, given the limited signal-to-noise ratio of our single-molecule method and the overlap between FRET distributions, we did not want to bias our analysis to presume a certain number of FRET states. Therefore, to impartially verify our visual assignment of FRET states from the raw trajectory data and the accuracy of the Gaussian fits, we performed a probability based

Hidden Markov modelling (HMM) analysis using HaMMy software.218 To further avoid biasing the results, the HaMMy software was set to converge on the true number of

FRET states in the data from initially guessing the maximum number of FRET states that the program allows (10). We analyzed a subset of single-molecule trajectories (100) for

63 each smFRET experiment by HMM as the HaMMy software was computationally limited in the number of traces that could be processed at one time under our experimental settings (i.e. 10 state model). The HaMMy output files were then additionally processed using accompanying TDP software to generate transition density plots. Kinetic information describing the transitions between states was calculated through the TDP software. Briefly, the transition distributions were fit to Gaussian functions and the calculated peak centers and standard deviations were converted to kinetic rates and rate errors, respectively, by multiplying by the exposure time used during data acquisition (100 ms). HaMMy and TDP were used primarily to complement and verify the results we obtained from the manual analysis of the complete data sets as well as to describe the kinetic rates connecting interconverting FRET states which would be difficult to acquire otherwise.

Thresholds were applied to the total FRET data set, and all FRET events within a

FRET threshold were pooled to conduct Dwell Time Analysis. Due to the overlap between the FRET states observed, the only threshold applied was to separate FRET events representing binding events from photobleaching events or unbinding events. The time associated with each binding event was extracted and plotted to generate a survival function of binding events in Matlab. Survivor functions were subsequently fit to single

(Equation 3.3) or double (Equation 3.4) exponential decay equations.

푓(푡) = 퐴exp(−푘푡) (Eq. 3.3)

푓(푡) = 퐴1 exp(−푘1푡) + 퐴2exp (−푘2푡) (Eq. 3.4)

64 where f(t) is the fraction of molecule remaining bound after time t, 퐴1 and 퐴2 are the amplitudes of the phases, and 푘, 푘1, and 푘2 are the rate constants of Dpo4 unbinding from the DNA substrate. Fits to the data were evaluated based on the coefficient of determination (R2) as well as visual inspection. To avoid over interpreting the results, all data were initially assumed to demonstrate single exponential decay behavior. If the R2 value was less than 0.99 and the fitting curve failed to adequately represent the majority of data, a higher order exponential fit was considered necessary (Equation 3.4).

3.2.6 Kinetic Assays

Polymerase activity was assessed through a burst kinetic assay at 37˚C in a reaction buffer containing 50 mM HEPES, (pH 7.5 at 37˚C), 50 mM NaCl, 0.1 mg/mL

BSA, 0.1 mM EDTA, 5 mM DTT, 10% glycerol, and 5 mM MgCl2. Briefly, 60 nM of 5ʹ-

32P-labeled D-1 DNA substrate was pre-incubated with 15 nM of either WT Dpo4 or

Cy5-labeled mutant (C31S, N70C) Dpo4 and then rapidly mixed with 100 μM dTTP for increasing amounts of time. All fast reactions were performed using a rapid chemical- quench flow apparatus (Kintek). DNA products were then resolved on a denaturing 17% polyacrylamide gel, scanned using a Typhoon Trio (GE Healthcare), and quantitated with

ImageQuant software (Molecular Dynamics). The product formation was then plotted against time (t) and the data were fit to Equation 5.

[Product] = A[1-exp(-푘푏푢푟푠푡t)] + 푘푠푠t (Eq. 5) where A represents the reaction amplitude, 푘푏푢푟푠푡 the single-turnover nucleotide incorporation rate constant, and 푘푠푠 the observed steady-state rate constant.

3.3 Results

3.3.1 Design of a FRET System for Monitoring Dpo4 Interaction with DNA

To investigate the binding of Dpo4 at the single-molecule level, we adapted a previously designed FRET system to attach a Cy5 fluorophore to the Finger domain of

Dpo4 (N70C) and a Cy3 fluorophore to each of the DNA substrates described in Figure

3.1.69 Notably, Dpo4 is known to bind the blunt-end of DNA with a five-fold lower affinity as compared to the primer/template junction of a double-stranded DNA substrate but does not bind to single-stranded DNA.79 Therefore, we added a non-complementary frayed end to each of the DNA substrates (Figure 3.1A) used in this article to eliminate off-target binding by Dpo4 which would have complicated data interpretation.

Fluorescence spectra recorded upon titration of Cy5-labeled Dpo4 into a solution of Cy3- labeled undamaged DNA or 8-oxo-dG containing DNA (damaged DNA) indicate that the

FRET probes are well-positioned to report on the binding of Dpo4 to DNA and that the fluorescent labels and 8-oxo-dG lesion did not affect the DNA binding affinity of Dpo4

(Figure 3.4), as reported previously.45,71 To determine if the mutations or the label affected the polymerase activity of Dpo4, burst kinetic assays of Cy5-labeled Dpo4 were performed to obtain single-turnover nucleotide incorporation and steady-state rate constants. The rate constants for the labelled enzyme are similar to those of wild-type

Dpo4 (Figure 3.5), indicating the mutations and fluorescent label did not alter the polymerase activity of Dpo4.

3.3.2 Investigation of Dpo4 in a Binary Complex with DNA by smFRET

To determine if the binding of Dpo4 to single DNA molecules is perturbed by the presence of a non-helix distorting oxidative lesion, 8-oxo-dG, we first conducted

66 smFRET experiments with DNA substrates containing either a dG, or an 8-oxo-dG lesion at the templating position in the DNA (Figure 3.1). Overall, smFRET time trajectories for Dpo4 binding to either DNA substrate were similar, displaying transitions between three, non-zero FRET efficiency values as determined by visual inspection (Figure

3.1D). FRET efficiencies of >200 individual traces from each single-molecule experiment were collected and compiled into population distribution histograms (Figures

3.6A, B). The Gaussian peak fits of the histograms revealed subpopulations with FRET efficiencies centered at ~0.50 (low-FRET state), ~0.65 (mid-FRET state), and ~0.85

(high-FRET state). Consistently, smFRET investigations of other DNA polymerases, such as E. coli DNA polymerase I (Klenow fragment), and S. solfataricus Polymerase

B1, have also revealed multiple FRET states which suggest various polymerase conformations while bound to DNA.120,124,217 Interestingly, we observed that the oxidative lesion affected the population distributions represented in the histograms

(Figure 3.6B). When bound to the undamaged DNA substrate, most FRET events (51%) were observed to be in the mid-FRET state. However, when an 8-oxo-dG lesion was present, most FRET events (50%) were observed to be in the low-FRET state.

Unexpectedly, our data revealed a small, distinct distribution at a FRET efficiency higher than the two previously described.73 This third state was observed regardless of the DNA substrate used. Given the inherent noise of smFRET data we chose to complement our manual data analysis with a probability-based Hidden Markov modelling analysis218 to unbiasedly confirm the number of non-zero FRET efficiency states. The software idealized a subset of FRET trajectories (100) and converged on three, non-zero FRET

67 states as expected from our visual inspection of the single-molecule trajectories (Figure

3.6A, B). Additional HMM analysis with the program TDP allowed for the construction of two-dimensional transition density plots (TDP) in which initial FRET efficiencies were plotted against final FRET efficiencies for every transition in the data sets (Figure

3.7). Importantly, if the smFRET data contain distinct, reproducible FRET values, then crosspeaks should develop in the TDPs. The corresponding plot in Figure 3.7 clearly shows crosspeaks representing transitions amongst all three states for each of the DNA substrates evaluated. Prominent density in the plots at the highmid, midhigh, highlow, lowhigh, midlow, and lowmid transition areas indicates that Dpo4 was able to freely convert between the three FRET states. Additionally, our TDP analysis gave the shuttling rates for the transitions between the FRET states. Shuttling rates between all states for Dpo4 binding to both DNA substrate were similar (Table 3.1).

To further investigate the response of Dpo4 to undamaged or damaged DNA, a dwell time analysis of the single-molecule binding traces was performed to yield information on the dissociation kinetics of the binary complexes. Dwell time histograms and survivor functions for this analysis are in Figures 3.8A and 3.9A. The survivor functions were best fit to a double exponential decay equation indicating that binding events for the binary complex (Dpo4•DNA) vary in stability (Table 3.2 and Figure

3.10). Notably, a modest decrease in the fast phase rate constant (koff,1) and increase in the slow phase amplitude (A2) for Dpo4 dissociation from the damaged DNA substrate relative to the undamaged one suggest a modest increase in the complex stability of

Dpo4•DNA caused by the presence of the oxidative lesion.

3.3.3 Effect of dNTPs on Dpo4 Binding to Undamaged or Damaged DNA

Single-molecule FRET experiments were performed to probe whether the presence of an incoming nucleotide affected the binding of Dpo4 to DNA. Notably, the addition of correct or incorrect dNTP (6 mM) eliminated the high-FRET state that was observed for the binary complex Dpo4•DNA with either the undamaged or damaged

DNA substrate (Figures 3.6, 3.11, 3.12, 3.13). With correct dCTP, smFRET time trajectories clearly displayed a single, sustained FRET state and the population histograms contain one low-FRET distribution (~0.5) (Figure 3.6C and 3.12). Further,

HMM and TDP analysis (Figure 3.7) indicate negligible transitions between non-zero

FRET states.

Dwell time analysis for Dpo4 binding to either undamaged or damaged DNA in the presence of a correct incoming nucleotide revealed slower (2- to 3-fold) dissociation kinetics compared to the binary complex (Dpo4•DNA) indicating that the correct dNTP stabilizes the ternary complex (Dpo4•DNA•dNTP) (Table 3.2). Dwell time histograms and survivor functions for this analysis are in Figures 3.8B and 3.9B. Remarkably, the correct nucleotide seemed to have a greater stabilizing effect for Dpo4 binding to the damaged DNA substrate as observed through slower dissociation rates (Table 3.2).

Experiments to observe the effect of an incorrect incoming nucleotide on the conformations and kinetics of Dpo4 binding to undamaged or damaged DNA substrates were performed to uncover the mechanism of nucleotide selection by Dpo4. smFRET trajectories of Dpo4 binding in the presence of each of the three incorrect nucleotides display short-lived FRET events with two primary FRET efficiencies (Figure 3.13).

Population histograms further revealed a bimodal distribution (Figure 3.11). Despite the high concentrations of incorrect nucleotide used in each experiment (6 mM), the bimodal distribution may be a result of an inability fully saturate the enzyme.77 Dwell time analysis confirmed that Dpo4 binding events to either DNA substrate were transient with fast dissociation kinetics (Table 3.2, Figures 3.8 and 3.9). However, dwell times for

Dpo4 binding to the damaged versus to the undamaged DNA substrate in the presence of an incorrect nucleotide were marginally longer lived. While the dwell time histograms for the dissociation of Dpo4 from undamaged or damaged DNA in the presence of incorrect nucleotide fit well to the single exponential decay equation (Equation 3), we cannot completely exclude the possibility of longer time scale processes.

3.4 Discussion

3.4.1 Analysis of the Low- and Mid-FRET States

Through smFRET investigation, we demonstrated that the binding of Dpo4 to

DNA is a complex process. The smFRET trajectories and FRET population histograms for this polymerase binding to an undamaged or a damaged DNA substrate (Figure 3.1A) revealed three FRET states (Figure 3.6A, B). Interestingly, our previous stopped-flow

FRET studies69,71,75 have revealed that Dpo4 and DNA forms an initial binary complex

(Complex I), which transitions into a ternary complex (Complex II) after an incoming dNTP stimulates the rapid translocation of Dpo4 along the DNA axis by one base pair in order to empty the space within the active site for nucleotide binding (Scheme 3.1A).

Subsequently, synchronized domain motions of Dpo4 help to tighten its “grip” on both the DNA and nucleotide, leading to the formation of Complex III. Prior to phosphodiester

70 bond formation, Complex IV is formed when Dpo4 undergoes the rate-limiting active site rearrangement to reposition key active site residues and properly align all substrates for catalysis (Scheme 3.1A).67,77 Notably, the binary and ternary crystal structures of

Dpo468,215 represent Complexes I and IV, respectively, but do not inform on Complexes

II and III (Scheme 3.1A). Surprisingly, the primer/template junction base pair in the binary crystal structure occupies the same space as the nascent base pair in the ternary structure,68 suggesting DNA translocation by one base pair during nucleotide binding.

Coupled with the aforementioned stopped-flow FRET69,71,75 and X-ray crystallographic studies68,215, we conclude that the mid-FRET state shown in Figures 3.6A and 3.6B likely represents Complex I while the low-FRET state corresponds to the conformation(s) of Dpo4 in Complex III and/or IV wherein DNA has translocated by one base pair and

Dpo4 has undergone synchronized domain motions. More importantly, the low-FRET state suggests that Dpo4 might have sampled the conformations in Complexes II-IV throughout the DNA binding process (Dpo4•DNA) in our smFRET experiments, even in the absence of an incoming nucleotide. However, as Complex II is a transient species, existing briefly during the nucleotide binding process, the low-FRET state observed during DNA binding by Dpo4 can only represent Complexes III or IV. Interestingly, the addition of correct nucleotide stabilized the polymerase in the low FRET efficiency state representing Complexes III and IV (Figures 3.6C and 3.12). Notably, Complexes III and

IV cannot be distinguished by FRET as they differ by small active site rearrangements which do not affect the distance between the FRET pair in our system (Figure 3.1).

Consistently, a recently published smFRET study of the binding of Dpo4 to

71 undamaged DNA has revealed two similar FRET states which are correlated to a pre- insertion and an insertion site on the DNA substrate.73 The pre-insertion site is consistent with our assignment of Complex I. However, those authors have not considered that the low-FRET state represents Complex III and/or IV. In addition, they concluded that the transition rates (1.1-2.7 s-1) between the two FRET states correspond to the rate of DNA translocation.73 Although our shuttling rates for Dpo4 transitioning between the mid- and low-FRET states with undamaged DNA (2.8-3.3 s-1, Table 3.1) are similar, these rates are too slow to account for the rapid translocation of Dpo4 along DNA. In fact, the translocation was too fast to be accurately measured by our stopped-flow FRET studies at

20 C, implying a rate of >150 s-1.69,71 Recent single-molecule investigations219-221 also suggest that the translocation rate is too fast to be resolved at the frame rates of conventional smFRET cameras (80 ms for the previous study73 and 100 ms for this study). Accurate rate estimation requires high temporal and spatial resolution such as that obtained in a recent study of phi29 polymerase where single-molecule nanopore technology was used to estimate a DNA translocation rate of >250 s-1 at 21 C.219-221

Interestingly, the synchronized domain motions of Dpo4 in Step 3 (Scheme 3.1A) occur at rates of ~10 s-1 which are reasonably comparable to our shuttling rates (Table

3.1).69,71,75 Taken together, the shuttling rates obtained here and in the previous study73 for Dpo4 transitioning between the mid- and low-FRET states are not specifically reporting on the DNA translocation event, but rather on concomitant domain motions as observed through the Cy5-labeled Finger domain. Thus, our single-molecule investigation yielded results which are consistent with those from ensemble studies69,71,75

72 and unveiled a dynamic conformational equilibrium in the domains of Dpo4 that accompanied DNA translocation.

3.4.2 Analysis of the high-FRET State

The high-FRET state (0.87) detected in Figure 3.6A was unexpected but confirmed through repeated experiments and unbiased HMM analysis. Analysis of our single molecule data indicates that Dpo4 can bind directly in this high-FRET state as well as access it through either the low-FRET or mid-FRET state (Scheme 3.1B). Notably, the absence of this high-FRET state in the previous smFRET study of Dpo473 likely stems from the different fluorophore labelling positions selected for the polymerase and DNA substrates in this work. Although more work is required to characterize this high-FRET state, we can speculate on its identity and significance here. Superposition of the apo and the binary (Dpo4•DNA) crystal structures has revealed large structural changes, especially a dramatic 131 rotation of the LF domain relative to the polymerase core, accompanying DNA binding.68 The conformational flexibility of the LF domain is facilitated by a highly flexible, 14 amino acid residue linker connecting the LF and

Thumb domains of Dpo4. This flexibility is further illustrated in the crystal structure of

Dpo4 in complex with a subunit of heterotrimeric proliferating cell nuclear antigen

(PCNA) where the LF domain exists in an extended conformation distinct from that in the apo and binary structures.85 Furthermore, our recent computational analysis of Dpo4 binding to DNA suggests that the LF domain and linker are intimately involved in the complex, multi-step processes the polymerase utilizes to recognize and bind DNA.61

Given the inherent plasticity of the LF domain and linker as well as the conformational

73 equilibria between multiple binding states (Schemes 3.1A, B), we hypothesize that the observed high-FRET state is related to an alternate DNA binding conformation of Dpo4 facilitated by both the dynamic LF domain and the flexible linker (Scheme 3.2A).

Interestingly, this high-FRET state is eliminated in the presence of dNTPs, suggesting that this binding mode may be non-productive and the presence of the nucleotide stimulates its conversion into a catalytically productive conformation at the primer/template junction. Alternatively, Dpo4 may sample other non-productive conformations, such as the polymerase binding to DNA in an inverted conformation which would shorten the distance between the Cy5 in the Finger domain and the Cy3 in the DNA substrate (Scheme 3.2B). Consistently, multiple binding orientations for an enzyme bound to a nucleic acid substrate have been previously observed by smFRET.222,223 Research is currently under way to explicitly characterize this high-FRET state.

3.4.3 Nucleotide Binding

By introducing an incoming nucleotide to the binary complex of Dpo4 with undamaged or damaged DNA, we revealed the stringent nucleotide selectivity of Dpo4 through smFRET experiments. Correct dNTP stabilized the binding of Dpo4•DNA as revealed by the complete shift of its FRET distributions to the low-FRET state

(Complexes III and IV in Scheme 3.1A) (Figures 3.6C and 3.12) and more than 2-fold slower dissociation kinetics of Dpo4 from undamaged or damaged DNA (Table 3.2).

Contrastingly, upon the introduction of an incorrect dNTP, we observed a bimodal population distribution as a result of incomplete population shifts to the low-FRET state

(Figure 3.11). Our dwell time analysis revealed that Dpo4 remained bound to DNA 5- to

10-fold longer in the presence of a correct over an incorrect nucleotide (Table 3.2).

Additionally, observed biphasic decay kinetics in the presence of correct dCTP (Figures

3.8B and 3.9B, Table 3.2) suggest the existence of the loose (Eʹ•DNA•dNTP) and tight

(Eʹʹ•DNA•dNTP) ternary complexes, which is consistent with our previously established kinetic mechanism (Scheme 3.1C).77,78 Interestingly, only fast, single exponential decay kinetics were observed in the presence of an incorrect nucleotide, suggesting the preferred collapse of the loose ternary complex (Figures 3.8 and 3.9, Table 3.2). Thus, our single-molecule analysis indicates that correct nucleotide binding allowed the formation of the catalytically competent, tight complex (Eʹʹ•DNA•dNTP). Interestingly, two competition experiments were conducted wherein the Dpo4•DNA complex was incubated with equal concentrations of either the three incorrect dNTPs or all four dNTPs. In both experiments, the FRET population distribution was bimodal. However, more FRET events were in the low-FRET state in the presence over absence of correct dCTP (83% versus 58%, Figure 3.14). Taken together, these smFRET experiments demonstrated that Dpo4 was able to differentiate between correct versus incorrect dNTPs during nucleotide binding.

Previously, our 32P-based kinetic assays monitoring the 8-oxo-dG bypass have revealed that Dpo4 exhibits surprisingly high fidelity and efficiency. In fact, the efficiency of correct incorporation opposite 8-oxo-dG is higher than that opposite the undamaged templating dG.45 Notably, our smFRET time trajectories are similar between

Dpo4 binding to undamaged versus damaged DNA (Dpo4•DNA) but a smaller low-

FRET state population was observed with the former (35% versus 50%, Figures 3.6A and B). More importantly, the dissociation kinetics of ternary complexes with correct dCTP show a larger population of slower dissociation events from the damaged over undamaged DNA substrates (69.8% versus 53.1%, Table 3.2). In the presence of either correct or incorrect nucleotides, ternary complex dissociation is noticeably slower from the damaged than the undamaged DNA substrate (Table 3.2). Consistently, the ternary crystal structure of Dpo4 with DNA containing a templating 8-oxo-dG and an incoming dCTP shows an auxiliary hydrogen bond and an ion-dipole pair between Arg332 and 8- oxo-dG.215 Overall, our smFRET analysis indicates that Dpo4 preferentially binds DNA containing 8-oxo-dG through additional stabilizing interactions with the lesion.

Interestingly, it has been previously hypothesized that Y-family DNA polymerases derive their lesion bypass specificity through unique active site residues which are optimal for their interactions with the templating lesion(s).133,224 Consistently, our smFRET experiments help to explain the preference of Dpo4 for bypassing 8-oxo-dG which constantly challenges the Sulfolobus solfataricus replication machinery in vivo.225

Notably, comparison of FRET efficiency histograms with damaged DNA (Figure

3.11) shows no preference of Dpo4 for any particular incorrect nucleotide, including dATP despite its capability of forming a Hoogsteen base pair with 8-oxo-dG.

Consistently, ternary crystal structures show that Arg332 stabilizes the anti- over syn- conformation of 8-oxo-dG at the active site of Dpo4 through the hydrogen bond and the ion-dipole pair discussed above.92,215

Author Contributions

Varun V. Gadkari and Austin T. Raper are co-first authors. V.V.G was credited for collecting the single-molecule FRET data of Dpo4 binding to undamaged DNA substrate, as well as the accompanying nucleotide binding studies. A.T.R was credited for collecting the single-molecule FRET data of Dpo4 binding to damaged DNA substrate and the accompanying nucleotide binding studies as well as utilizing Hidden Markov modelling to rationalize the data. V.V.G and A.T.R worked together to collect all supplemental data, and analyze the results of all experiments. Brian A. Maxwell, collected some of the initial single-molecule data for Dpo4, which were not included in the paper, and designed the FRET system to monitor the interaction. Additionally, he was involved in training both co-first authors as they completed this study. A.T.R. and Dr.

Zucai Suo were involved in proposing a kinetic model for the data. The manuscript was written by V.V.G and A.T.R, along with the corresponding author Z.S. The research was funded by the National Institutes of Health [ES009127 to Z.S., T32GM008512 to

A.T.R.]; National Science Foundation [MCB-0960961 to Z.S.].

3.5 Schemes

Scheme 3.1: Proposed Mechanism of Binary and Ternary Complex Formation

(A) Previous proposed mechanism of nucleotide incorporation. E, Eʹ, and Eʹʹ are different forms of the enzyme. Steps after 4 were not shown for clarity of discussion. (B) Binary complex formation. [E•DNA]L,

M H [E•DNA] , and [E•DNA] refer to low-, mid-, and high-FRET binary complexes, respectively. (C)

L L Ternary complex mechanism. [Eʹ•DNA•dNTP] and [Eʹʹ•DNA•dNTP] refer to two different forms of the ternary complex.

Scheme 3.2: Conformational sampling of the high-FRET state

(A) The LF domain of Dpo4 has been shown to adopt multiple structural conformations. The high-FRET state may be an extended conformation of the LF domain. Proximity between fluorophores increases during transition from a compact binary complex to an extended conformation. The LF binds DNA autonomously allowing the core domains to sample conformational states and DNA surface area as facilitated by the flexible linker region (B) Dpo4 may adopt a nonproductive conformation in which the polymerase is incorrectly oriented on the DNA resulting in high-FRET as the proximity between Cy5 in the Finger domain and Cy3 on the substrate increases.

3.6 Tables

Table 3.1: Shuttling Rates from Transition Density Plot

Transition Undamaged Substrate 8-oxo-dG modified Substrate (s-1) (s-1)

LM 2.8 ± 0.7 3.2 ± 0.8 LH 1.5 ± 0.4 1.0 ± 0.4 ML 3.3 ± 0.8 4 ± 2 MH 1.8 ± 0.7 1.8 ± 0.5 HL 1.8 ± 0.4 5 ± 2 HM 1.9 ± 0.5 3 ± 1

L = Low-FRET, M = Mid-FRET, H = High-FRET

Table 3.2: Dwell Time Analysis of Dpo4 Binding to DNA

Average koff,1 koff,2 Binding A1 -1 A2 -1 Time (s) (s ) (s ) Undamaged DNA Substrate DNA 1.3 0.51 ± 0.02 (50.5%) 2.5 ± 0.1 0.50 ± 0.02 (49.5%) 0.50 ± 0.01 DNA + dCTP 2.7 0.46 ± 0.02 (46.9%) 6.3 ± 0.5 0.52 ± 0.01 (53.1%) 0.215 ± 0.005 DNA + dATP 0.3 1.01 ± 0.01 5.0 ± 0.1 - - DNA + dGTP 0.2 1.02 ± 0.02 6.2 ± 0.2 - - DNA + dTTP 0.3 1.02 ± 0.02 5.7 ± 0.2 - - Damaged DNA Substrate DNA 1.6 0.38 ± 0.04 (38.8%) 1.7 ± 0.01 0.60 ± 0.04 (61.2%) 0.46 ± 0.02 DNA + dCTP 4.2 0.29 ± 0.01 (30.2%) 1.16 ± 0.08 0.67 ± 0.01 (69.8%) 0.185 ± 0.003 DNA + dATP 0.8 0.97± 0.02 1.38 ± 0.04 - - DNA + dGTP 0.6 1.04 ± 0.02 2.2 ± 0.07 - - DNA + dTTP 0.9 0.92± 0.02 1.13 ± 0.03 - -

3.7 Figures

Figure 3.1: Single-molecule FRET analysis of Dpo4 binding to DNA

(A) DNA substrate used in single-molecule experiments. The primer is biotinylated at the 5ʹ end (denoted

“B”) to facilitate surface immobilization. “T” denotes Cy3 attached to 5-C6-amino-2ʹ-deoxythymidine.

“G” represents 8-oxo-7,8-dihydro-2ʹ-deoxyguanine (8-oxo-dG) or undamaged dG. (B) The binary complex of Dpo4 and DNA accessed through PDB code 2RDJ. The green star represents a Cy3 donor label on the

DNA substrate. The red star signifies the site of Cy5 acceptor labeling in the Finger domain (N70C) of

Dpo4. (C) Chemical structure of 8-oxo-dG with chemical modifications at the C8 and N7 positions of guanine circled in red. (D) Representative single-molecule fluorescence trajectory for Dpo4 binding to the

DNA substrate containing 8-oxo-dG. Donor and acceptor fluorescence signals are shown in green and red, respectively.

Figure 3.2: Verification of Observed FRET

Potential photophysical phenomenon due to multiple donor molecules immobilized near or protein induced fluorescence enhancement (PIFE), which may introduce ambiguity in data analysis, must be ruled out to verify the legitimacy of the designed FRET system. PIFE is characterized by an increase in the intensity of a fluorophore upon proximal binding of an unlabeled protein. (A) To verify that our DNA coverage was sufficient to observe single, distinct Cy3-labeled DNA molecules, we directly excited 20 pM of surface immobilized Cy3-labeled undamaged DNA substrate (Figure 3.1) in the absence of any proteins or acceptor fluorophores. Donor fluorescence intensity of single molecules was recorded. Constant fluorescence intensity was observed in the donor channel until one-step photobleaching, whereas none was observed in the acceptor channel (data not shown). Notably, multiple photobleaching events would have suggested too dense of DNA surface coverage. (B) Wild-type Dpo4 was introduced into the chamber containing surface immobilized Cy3-labeled DNA. The donor fluorescence intensity did not change appreciably, and once again no fluorescence intensity above background was observed in the acceptor channel. Introduction of 10 nM Cy5-labeled Dpo4 produced decreases in donor fluorescence intensity with corresponding increases in the acceptor fluorescence intensity as shown in Figure 3.1D. 83

Figure 3.3: Fluorescence Assay for Multiplicity of Binding

10 nM Cy5-labled Dpo4 was introduced into a chamber containing surface immobilized Cy3-labeled DNA.

A 638 nm laser was used to directly excite any Cy5-labeled Dpo4 molecules that were immobilized due to

DNA binding. Acceptor channel fluorescence intensity trajectories were recorded, and >100 trajectories were analyzed and used to generate the histogram. A single distribution was observed at an intensity of

~350 (first dashed line) indicating that the Dpo4 molecules bind the DNA substrates in a 1:1 ratio. If multiple Dpo4 molecules were bound to the DNA, a distribution at a higher intensity (~700, second dashed line) due to the additive fluorescence intensities of multiple acceptor fluorophores would have been observed. The large peak at zero intensity is due to background. 84

Figure 3.4: DNA binding affinity of Dpo4

(A) Example fluorescence titration. Cy3-

DNA (25 nM) was excited at 532 nM and

fluorescence emission spectra were

recorded at varying concentrations of

Cy5-Dpo4 N70C ranging from 0 nM

(black) to 180 nM (pink). (B) Change in

donor quantum yield (ΔΦDonor) for the

undamaged (●) or 8-oxo-dG modified (■)

DNA substrates upon titration with Dpo4.

The curves were fit to Eq. 1 to yield a Kd

of 11 ± 1 nM for Dpo4 binding to

undamaged DNA and a Kd of 10 ±1 nM

for Dpo4 binding to the 8-oxo-dG

modified DNA.

Figure 3.5: Burst Assay to Confirm Dpo4 Activity

A preincubated solution of either WT (■), or Cy5-labeled mutant (●) Dpo4 (15 nM) and 21/41mer with a

5ʹ-32P-labeled primer (60 nM) was mixed with dTTP (100 μM) in a rapid chemical quench flow apparatus.

The reactions were quenched at various times with EDTA to 0.37 M, and products were separated by denaturing PAGE and quantitated using ImageQuant. The data were fit by nonlinear regression to the following burst equation: [product] = A[1-exp(-k1t) + k2t] where A is the amplitude of active enzyme, k1 is the observed burst rate constant, and k2 is the observed steady-state rate constant. The rate constants for the wild-type Dpo4 were 1.8 ± 0.6 s-1 and 0.04 ± 0.01 s-1 for the exponential and linear phases, respectively.

The rate constants for the Cy5-labeled mutant Dpo4 were 1.4 ± 0.4 s-1 and 0.03 ± 0.01 s-1 for the exponential and linear phases, respectively. These similar rate constants between wild-type Dpo4 and Cy5- labeled Dpo4 indicate that the fluorescent labeling did not affect the activity of the polymerase.

Figure 3.6: FRET from Dpo4 binding to single DNA molecules

FRET efficiency histograms (left) and representative smFRET trajectories (right) of Dpo4 binding to (A) undamaged DNA (B) 8-oxo-dG modified DNA and (C) 8-oxo-dG modified DNA with 6 mM of correct nucleotide (dCTP). On the left, individual Gaussian peak fits for each state are shown by the dashed black lines with respective population percentage indicated in red. The sum of the individual Gaussians is shown as the solid black line. On the right, representative FRET trajectories are shown in blue and the black lines depict HMM fits to the data.

Figure 3.7: Transition Density Plots

Transition density plots from experiments monitoring the following complexes: (A) Cy5-

Dpo4●Undamaged DNA. (B) Cy5-Dpo4●8-oxoG DNA. (C) Cy5-Dpo4●Undamaged DNA+6 mM dCTP.

(D) Cy5-Dpo4●8-oxoG DNA+6 mM dCTP. The transition density plots (TDP) show the transitions between the low, mid, and high FRET states. FRET efficiencies are plotted based on their initial state, and final state. The recurrence of a particular transition by many molecules results in a crosspeak at that position on the plot. The crosspeaks are represented as colored regions shaded blue to red to yellow to represent an increasing probability of occurrence. Notably, many transitions between low-, mid-, and high-

FRET states are observed when Dpo4 binds DNA, however these transitions are nearly eliminated in the presence of the correct nucleotide (dCTP) at saturating concentrations.

Figure 3.8: Dwell Time Analysis of Dpo4 Binding Undamaged DNA

Survivor functions for the DNA bound state were computed from histograms of binding times from multiple trajectories with 10 nM Cy5-Dpo4 binding to undamaged DNA with or without nucleotide

(Figure 1). Double or single exponential fits are depicted as the black solid lines. (A) Binary Complex (B) with 6 mM dCTP (C) with 6 mM dATP (D) with 6 mM dGTP (E) with 6 mM dTTP

Figure 3.9: Dwell Time Analysis of Dpo4 Binding Damaged DNA

Survivor functions for the DNA bound state were computed from histograms of binding times from multiple trajectories with 10 nM Cy5-Dpo4 binding to damaged DNA with or without nucleotide (Figure

1).Double or single exponential fits are depicted as the black solid lines. (A) Binary Complex (B) with 6 mM dCTP (C) with 6 mM dATP (D) with 6 mM dGTP (E) with 6 mM dTTP

Figure 3.10: Example Dwell Time Survivor Function

Survivor function for the DNA bound state was computed from a histogram of binding times from multiple trajectories with 10 nM Cy5-Dpo4 binding to undamaged DNA. The survivor function was fit to a double exponential decay equation. 50.5% of the population was dissociated in a fast phase with a rate constant of

2.5 ± 0.1 s-1, while 49.5% of the population was dissociated through a slow phase of 0.5 ± 0.01 s-1. Included as a comparsion to the best fit solid line, the dashed line depicts a single exponential fit to the data.

Figure 3.11: FRET Histograms of Dpo4 binding DNA with incorrect dNTP

(A) Population distribution histograms generated from multiple time trajectories of Dpo4 binding to undamaged DNA (top) or 8-oxo-dG modified DNA (bottom) in the presence of 6 mM of either dTTP (A,

D), dGTP (B, E), or dATP (C, F). Individual Gaussian peak fits for each state are shown by the dashed black lines with respective population percentage indicated in red. The sum of the individual Gaussians are shown as the solid black line in each.

Figure 3.12: FRET from Dpo4 binding to single undamaged DNA molecules in the presence of saturating dCTP

(A) FRET efficiency histogram with Gaussian peak fit shown as a solid black line for Dpo4 binding to undamaged DNA with saturating concentration of correct nucleotide (dCTP). (B) Representative single- molecule FRET trajectory of Dpo4 binding to undamaged DNA with saturating concentration of correct nucleotide (dCTP) with HMM data fit depicted as solid black line.

Figure 3.13: Representative FRET Trajectory for Dpo4 binding DNA in the presence of saturating incorrect nucleotide

(A) Representative single-molecule trajectory is shown for a Cy5-labeled Dpo4 molecule bound to

Cy3-labeled undamaged DNA, in the presence of 6 mM dTTP. Donor and acceptor intensities are shown in green and red, respectively. (B) The corresponding FRET signal is shown in blue.

The dotted black lines represent the two main FRET states observed in this experiment: a low

FRET state at ~0.4 and a mid-FRET state at ~0.6. In general, the binding events are very transient, resulting in an average binding time in this experiment of 0.3 seconds.

Undamaged DNA Undamaged DNA dATP, dGTP, dTTP dATP, dGTP, dTTP, dCTP A B

58% 42% 83% 17%

Figure 3.14: Selectivity of Dpo4 for the Correct Nucleotide

Single molecule experiments were conducted using the conditions described in the “Experimental Section”.

Cy5-labeled Dpo4 binding undamaged DNA with either (A) 2 mM dATP, 2 mM dGTP, and 2mM dTTP, or (B) all four nucleotides (1.5 mM each). The total nucleotide concentration in both experiments was 6 mM. (A) In the presence of the three incorrect nucleotides, the population histogram was bimodal as seen previously with the incorrect nucleotide experiments, with 58% of molecules bound in the low FRET state, and 42% bound in the mid FRET state. However, with the addition of all four nucleotides (B), the population of the low FRET state increased 58% to 83%, resembling the trend previously observed with dCTP where almost all molecules are bound in the low FRET state. 17% of molecules still bound in the mid FRET state. This is likely due to the presence of the other three incorrect nucleotides in a 3:1 excess over the correct dCTP.

Chapter 4 Investigation of DNA Clamp Opening and Closing by Single-Molecule

Forster Resonance Energy Transfer

4.1 Introduction

In all three domains of life, DNA replication is an intricate and stringently coordinated process that ensures the faithful and efficient replication of the cell’s genetic material. Logically, such a complicated process is accomplished by a variety of proteins working in concert. Various enzymes, and proteins are involved in DNA related transactions, often working in coordination to ensure complete and faithful replication and maintenance of the DNA.20,31,37,182,224,226-236 In all organisms, this multimolecular model of DNA metabolism is coordinated by an evolutionarily conserved sliding DNA clamp, which encircles the DNA duplex, and serves as a scaffold to tether the replication machinery.165-167 The sliding DNA clamps have been identified and studied in all domains of life, e.g. the β clamp of Escherichia coli DNA Pol III, the gene 45 protein of

T4 bacteriophage, and proliferating cell nuclear antigen (PCNA) in yeast, humans, and the archaeon Sulfolobus solfataricus.168,169 Previous studies have shown that despite low sequence similarity, all sliding DNA clamps share a remarkably similar toroidal structure with a central hole to accommodate a DNA duplex.169

Interestingly, despite the highly conserved ring-shaped structure of sliding clamps, the complexity of their subunit composition varies from domain to domain. As observed in E. coli, the bacterial β clamp is a homodimer, while PCNA clamps in 96 archaeal and eukaryotic organisms are trimeric.169 Furthermore, while most PCNAs, are homotrimeric, PCNA from the archaeon Sulfolobus solfataricus (Sso) is a heterotrimer consisting of three distinct PCNA monomer subunits: PCNA1, 2, and 3.168 The heterotrimeric composition of Sso PCNA presents a unique advantage from an experimental standpoint. The individual monomers and their respective interfaces have been studied to identify specific interactions with partner proteins85,168,178,179 and adjacent

PCNA monomers.179,189,237 Since Sso PCNA has three distinct interfaces, previous studies have established that the formation of the trimer follows a sequential process, which begins with the formation of the PCNA1:PCNA2 dimer, and subsequent recruitment of

PCNA3 to form the stable functional heterotrimer168,179,189,237 which only opens at the

PCNA1:PCNA3 interface to be loaded onto DNA.238 Although biochemical characterization has suggested that the Sso PCNA opens at the PCNA1:PCNA3 interface, no structural evidence is currently available to support this claim. To date, the only existing crystal structures of Sso PCNA clamp are those of the individual monomers,237 the PCNA1:PCNA2 dimer,237 the PCNA1:PCNA2 dimer associated with proteins,85,178 and the closed PCNA123 clamp.179,189,237 In fact, the only crystal structure of an open

DNA clamp currently available in the RCSB Protein Data Bank is that of the open DNA clamp from T4 bacteriophage (gp45 homotrimer) in complex with its clamp loader

(gp62/gp44 complex), a heteropentameric complex which opens DNA clamps in an ATP- dependent manner.239 Currently no structure of a lone open sliding DNA clamp exists.

A previous set of fluorescence studies reports that gp45 DNA clamp from T4 bacteriophage, a homotrimer, exists primarily in an open form in solution,240 despite the

97 fact that the open form of gp45 trimer has never been crystallized without a clamp loader protein present. Thus, we were motivated to investigate the solution state dynamics of

Sulfolobus solfataricus PCNA and to determine whether it can stably exist in an open form in the absence of a clamp loader. We chose to utilize two unique techniques which yield structural information and report on protein conformational dynamics. First, we used Forster resonance energy transfer (FRET), a popular fluorescence technique used to study conformational dynamics of protein-protein and protein-ligand interactions, as well as intramolecular domain motion. To investigate if we could observe conformational behavior of PCNA in solution, we used single molecule FRET (smFRET) so that we could collect FRET data at a molecular level to obtain population data about the preference of PCNA to assume a closed or open conformation in solution. We discovered that the PCNA molecules assumed one of two conformations. Based on FRET efficiency and known structural data, we believe these to be the closed and open forms of PCNA.

Furthermore, we found that the equilibrium between the two forms of PCNA could be controlled by the ionic strength of the environment. In lower salt conditions the DNA clamp is mostly in the closed conformation, and as the salt concentration is increased, the population of DNA clamps tend to favor a more open and extended conformation. It is possible that in vivo where the salt concentration is likely closer to the lower salt concentrations we measured, the sliding DNA clamps exist in both open and closed conformations, but primarily favoring the closed. Thus, DNA clamp loaders are a means of counteracting this equilibrium, by opening the closed DNA clamp by an ATPase driven reaction to counteract the favorable closed conformation.

4.2 Materials and Methods

4.2.1 Expression of Proteins

The covalently linked PCNA heterotrimer (PCNA1-2-3) is expressed as a fusion from a single gene which encodes a 20-peptide linker (ASGAGGSEGGGSEGGTSGAT), one between PCNA1 and PCNA2, and another between PCNA2 and PCNA3. Plasmid to express PCNA1-2-3 was obtained from the lab of Dr. Stephen D. Bell at the University of

Indiana in Bloomington, IN. The PCNA1-2-3 plasmid was mutated to introduce SC mutations at positions PCNA1(S64) and PCNA3(S189) to generate two cysteines for site- specific fluorophore labeling. Additionally, a gene sequence to code for an AviTag peptide (GLNDIFEAQKIEWHE) was cloned into the plasmid to be encoded after the

PCNA1-2-3 protein, and before the 6xHis tag at the C-terminal end (Figure 4.1A). A plasmid containing the gene for Sso Replication Factor C was obtained from the lab of

Dr. Michael Trakselis at Baylor University. All expression plasmids were individually transformed into E. coli strain Rosetta (DE3) and expressed separately using autoinduction in ZYP-5052 medium.241 After growth, cells were harvested (4000 rpm for

20 min) and resuspended in Buffer A (50 mM HEPES [pH 8 at 4⁰C] , 100 mM NaCl, and

1 mM DTT). The resuspended cells were lysed by French pressure cell at 20,000 PSI three times, and the resulting lysate was cleared by ultracentrifugation (40,000 rpm for 40 min). Cleared lysates of all proteins were incubated at 50⁰C for 10 min to precipitate thermostable E. coli proteins which were removed by a second ultracentrifugation step.

The thermostable target proteins were purified from the heat shock supernatant by column chromatography.

4.2.2 Purification of Proteins

The covalently linked PCNA1-2-3 protein has a C-terminal 6x histidine tag, thus the heat shock supernatant was first purified over Ni2+-NTA resin (Qiagen). Bound protein was eluted by linear gradient to 1M Imidazole. Fractions containing PCNA1-2-3 were identified by chromatogram, and SDS-PAGE, pooled, and dialyzed against Buffer

A. The dialyzed sample was then loaded onto a HiTrap Heparin column pre-equilibrated with Buffer A. Protein was eluted by linear gradient to 1 M NaCl, and the fractions containing PCNA1-2-3 were pooled, dialyzed against Buffer A, concentrated to less than

1 mL, and loaded onto HiPrep 26/60 Sephacryl S-100. Purified PCNA1-2-3 was biotinylated following a previously published protocol242,243 and then doubly labeled with fluorophore dyes at the engineered Cys sites in PCNA1, and PCNA3. The protein was incubated with 15-fold molar excess of Cy3-, and Cy5-maleimide (Lumiprobe), overnight at 4⁰C in a buffer containing 50 mM Tris (pH 7.2), 150 mM NaCl, 0.5 mM TCEP, and

10% glycerol. Unincorporated free dye was removed by running the sample through

Superdex 200 10/300 (GE Healthcare). Labeling verified by measuring absorbance at

280, 570, and 650 nm, for protein, Cy3, and Cy5 respectively.

Sso Replication Factor C is a heteropentamer consisting of one large subunit, and four small subunits. The heteropentamer was purified by HiTrapQ, HiTrap Heparin, and size exclusion chromatography. Supernatant containing the RFC heteropentamer was loaded onto a 5 mL HiTrap Q (GE Healthcare), pre-equilibrated with buffer A. Protein was eluted by using a linear gradient to 1 M NaCl. After analysis by SDS-PAGE, the fractions containing clean protein with the correct ratio of small to large subunit were

100 pooled, dialyzed against Buffer A, loaded onto a 5 mL HiTrap Heparin (GE Healthcare), and eluted by linear gradient to 1 M NaCl. After SDS-PAGE, and quantitation of nucleic acid contamination by UV absorbance, pure protein sample free of nucleic acid was dialyzed against Buffer A, concentrated to less than 1 mL, and loaded onto a HiPrep

26/60 Sephacryl S-100 column (GE Healthcare) for purification by size exclusion to select for the correct heteropentameric form of RFC. RFC activity is tested by an ATPase assay to confirm that the RFC turns over ATP at rates comparable to those reported in literature.180

4.2.3 Single Molecule Data Acquisition

Single-molecule fluorescence experiments were conducted on a custom built prism-type total internal reflection microscope described before in Chapter 3 Section 3.2.3. Cy3 donor dyes were directly excited using a 532 nm laser (CrystaLaser, 100 mW) and resulting emitted fluorescence was passed through a set of optics to separate the donor and acceptor channel, as described in Chapter 3 Section 3.2.3. The FRET interactions were imaged in real time using an Andor iXon 897 electron-multiplying charge-coupled device camera to record movies at 2 frames per second, at 20⁰C, over several minutes.

4.2.4 Single Molecule Measurements

Imaging chambers were assembled from quartz slides and coverslips which were cleaned, passivated, and biotinylated. Right before the experiment, the chambers were loaded with neutravidin (0.2 mg/mL) in T50 buffer (10 mM Tris HCl, pH8, 50 mM

NaCl). After a 5 min incubation, the excess neutravidin was washed away with T50. The biotinylated, and doubly labeled PCNA1-2-3 was loaded into the chamber in imaging

101 buffer (50 mM HEPES [pH 7.6], 0.8% w/v D-Glucose, 2 mM Trolox, 0.1 mg/ml BSA, 1 mg/ml glucose oxidase, 0.04 mg/ml catalase), and incubated for 5 minutes to allow for sufficient immobilization. Immobilization was verified by excitation of the sample chamber with the 532 nm laser and viewing different areas of the sample chamber using the EM-CCD camera. Once sufficient immobilization was achieved, the excess molecules were washed out using imaging buffer of the desired salt concentration. Single molecule measurements were conducted in 0, 250, 500, 750, and 1000 mM NaCl. For the experiments with RFC, the chamber was washed after immobilization of PCNA1-2-3, and a reaction mix (50 mM HEPES [pH 7.6], 0.8% w/v D-Glucose, 2 mM Trolox, 0.1 mg/ml BSA, 1 mg/ml glucose oxidase, 0.04 mg/ml catalase, 150 mM NaCl, 2 mM MgCl2

1 μM RFC, and 5 mM ATP) was loaded to the chamber. Movies were recorded for several minutes to capture FRET, as well as donor and acceptor photobleaching events.

4.2.5 Single Molecule Data Analysis

The movies were processed using IDL (ITT Visual Information Solutions) to generate raw donor/acceptor fluorescence, and FRET traces for every molecule. The traces were then filtered in MATLAB based on adherence to a stringent set of guidelines.

Only traces that showed a clear FRET event, as well as a single donor and acceptor photobleaching event were selected. FRET traces that exhibited fluorophore blinking

(multiple photobleaching events) were eliminated from analysis. The selected traces were then corrected for background using a custom software package (Center for the Physics of Living Cells, University of Illinois at Urbana-Champaign). The FRET efficiency values calculated by the MATLAB software were calculated as apparent FRET (Eapp) per

102

Equation 4.1, where ID, and IA are the donor and acceptor fluorescence intensities, respectively.

퐼A 퐸푎푝푝 = (Eq. 4.1) 퐼D+퐼A

For every experiment, FRET efficiency values from >200 FRET trajectories were collected and binned to generate histograms depicting the population distribution of molecules over FRET efficiencies from 0 to 1. The population distributions were fit to sum of Gaussian function using MATLAB, and percent occupancy of FRET states was calculated as the total area under the individual Gaussian fits (red dotted lines). The histograms were fit with two peaks based on the consistent observation of two distinct

FRET states observed during raw data analysis. The summed fit (black line) was plotted to show the integrity of the two fits to the overall data set.

Dwell time analysis was performed to determine the time spent in individual

FRET efficiency states by the PCNA1-2-3 molecules. Thresholds were applied to the total FRET data set, and all FRET events within a FRET threshold were pooled. The thresholds were chosen based on the population histograms and the visually discernable

FRET states in the raw data analysis. A threshold was applied at FRET efficiency of 0.2, to eliminate background after photobleaching. The “Low FRET” state was limited by thresholds at FRET efficiencies of 0.2 and 0.65, and the “High FRET” state was limited by thresholds at FRET efficiencies of 0.65 and 1. The time associated with each FRET event in the respective subsets was extracted and plotted versus time to generate a survival function of FRET events associated with the two FRET states observed. The survival functions were fit to the single exponential decay equation (Equation 4.2),

103 where f(t) is the fraction of molecules in the designated FRET state after time t, A is the amplitude of the function, and k is the decay rate constant associated with the designated

FRET state.

푓(푡) = 퐴푒−푘푡 (Eq. 4.2)

4.3 Results

4.3.1 Design of the Covalently Linked FRET Construct

Previous studies have reported that the PCNA3 monomer alone has no interaction with the lone PCNA1 monomer, and a very weak interaction with the PCNA1:PCNA2 dimer. Despite this, all three monomers ultimately assemble to form a stable, and functional heterotrimer.168 Based on our knowledge of the DNA replication machinery we speculate that the sliding DNA clamps are not always loaded on DNA or bound to other proteins. Thus, they are most likely in solution until such time that they are needed.

We intend to characterize how the Sso PCNA heterotrimer molecules behave in solution in the absence of DNA or other known binding partners using single-molecule FRET

(smFRET). Particularly, we wanted to explore whether all formed heterotrimers exist in the significantly more stable closed form, or in an equilibrium between the open and closed conformations. Our smFRET protein construct was based on a previous study which conclusively proved that only the PCNA1:PCNA3 interface of the heterotrimer participates in clamp opening. Briefly, a series of covalently linked constructs of the Sso

PCNA heterotrimer were used to prove that the clamp is only opened and loaded onto

DNA by the clamp loader, Sso Replication Factor C (RFC), if the PCNA1:3 interface was available for opening.238

104

For our experiments, the PCNA1 and PCNA3 monomers were both labeled with fluorescent dyes, to report on the opening and closing of PCNA by single molecule

FRET. However, the concentration of PCNA3 necessary to overcome the very low affinity of PCNA3 for the PCNA1:PCNA2 dimer would result in extremely high fluorescent background, and a total loss of observable data. To overcome this issue, we chose to conduct our experiments using a previously studied covalently linked trimer.238

A flexible 20 peptide linker (ASGAGGSEGGGSEGGTSGAT) connected PCNA1 (C- terminal) to PCNA2 (N-terminal), and PCNA2 (C-terminal) to PCNA3 (N-terminal), without interfering with the binding interfaces between subunits. Biochemical assays238 showed that this covalently linked construct behaved in a similar manner to the free trimer as a DNA polymerase processivity factor which significantly improved product lengths in a primer extension assay. Briefly, the previous study showed that similarly extended products were observed in a primer extension assay when observing the free

PCNA123 trimer alongside the linked PCNA1-2-3 trimer. Additionally, when the clamp loader Replication Factor C (RFC) was introduced into the extension assays, only the lanes with the PCNA1:PCNA3 interface available for opening exhibited an RFC dependent increase in product length.238

The covalently linked PCNA (PCNA1-2-3) was site-specifically biotinylated, and double labeled with fluorescent dyes for the smFRET experiments. The protein was biotinylated at a Lys residue in a C-terminal, 15 amino acid Avi-Tag sequence right after the PCNA3 subunit, using E. coli enzyme BirA as previously reported (Figure

4.1A).242,243 The biotin was used to immobilize PCNA1-2-3 molecules on the single

105 molecule slides using the near-covalent biotin:neutravidin interaction. PCNA1-2-3 was also doubly labeled with Cy3 and Cy5 fluorescence dyes, at site-specifically engineered cysteines at positions PCNA1(C64) and PCNA3(C189). These sites were chosen due to their ability to report on the closed and open states of PCNA. In the closed form of

PCNA, the fluorophores are approximately 45 angstroms apart within range of the

Förster radius of the Cy3-Cy5 FRET pair (54 Angstroms). We expected that this labeling scheme would yield distinct FRET states that could be assigned to the open, and closed forms of PCNA1-2-3. It should be noted that there will be subpopulations of the protein sample that are either singly labeled, or doubly labeled with the same fluorophore. These molecules will not exhibit a FRET interaction, and thus they will be omitted from the analysis due to lack of FRET events.

4.3.2 Single-Molecule FRET Population Histograms

Previous studies have established that the interfaces between the three monomers are partly stabilized by ionic interactions.179,189,237 Thus, to determine if PCNA1-2-3 was dynamic in solution, we conducted five smFRET experiments at varying concentrations of NaCl (0-1000 mM). Initial visual inspection of the raw fluorescence vs. time traces showed that the PCNA1-2-3 molecules were sampling two distinct non-zero FRET states, and could transition freely between them (Figure 4.2). We observed molecules exhibiting high FRET (FRET eff. ≈ 0.7-0.9) which we attributed to a closed form of

PCNA1-2-3 (Figure 4.1 B, D), and a low FRET (FRET eff. ≈ 0.5-0.6) form corresponding to the open PCNA1-2-3 (Figure 4.1 C, E). While some traces showed transitions between the two FRET states such as the molecules in Figure 4.2, others

106 showed only one long FRET event (>60 s) such as those in Figure 4.1D, and E suggesting that the PCNA1-2-3 molecules are not constantly dynamic.

For each experiment (0 mM NaCl, 250 mM NaCl, 500 mM NaCl, 750 mM NaCl,

1000 mM NaCl), raw FRET data from over 200 traces was collected, processed to remove background, and compiled into a population distribution histogram. All five histograms revealed a distinct bimodal distribution of the molecules into two FRET states, with peaks centered at 0.5 (Low FRET), and 0.8 (High FRET) (Figure 4.3A-E).

Fitting the histograms to a sum of Gaussians function generated occupancy percentages for the two subpopulations. Our results demonstrate that the PCNA1-2-3 molecules exist in an equilibrium between a closed (high FRET), and open (low FRET) form, and that this equilibrium can be perturbed by varying the concentration of NaCl in solution. At 0 mM NaCl, the PCNA1-2-3 molecules were primarily in the high FRET closed conformation, with only about 24% of molecules observed in the open state (Figure

4.3A). Increasing the concentration of NaCl to 250, 500, 750, and 1000 mM resulted in the increase of the low FRET distribution to 42, 52, 65, and 75 % respectively, demonstrating a direct linear correlation between fraction of PCNA1-2-3 molecules in the open state, and concentration of NaCl (Figure 4.3B-F). As the population of open

PCNA1-2-3 molecules increased, we observed a concomitant decrease in the population of closed PCNA1-2-3 molecules suggesting that the increased ionic strength of the buffer was causing the opening of closed PCNA clamps.

4.3.3 Dwell Time Analysis

We performed a dwell time analysis of any single-molecule traces displaying

107 evidence of FRET transitions between the two non-zero FRET states to investigate the kinetics of PCNA opening and closing. PCNA1-2-3 opening is represented by a FRET transition from high FRET to low FRET. Thus, only transitions between the high and low

FRET states were analyzed since transitions to zero were disregarded as photobleaching events. The duration of high FRET states (Closed PCNA lifetimes), and the durations of low FRET states between high FRET events (Open PCNA lifetimes) were recorded and compiled to generate histograms. The integration of these histograms produced survivor functions for the high, and low FRET states, representing the survival of the closed and open form of PCNA respectively (Figure 4.4 A, B). Fitting the survivor functions to a single exponential decay function yields a kinetic parameter representing the decay of the

FRET state being monitored. Fitting the survivor function of the closed PCNA (high

FRET) yields kopen, the rate of PCNA opening (Table 4.1) while fitting the survivor function of open PCNA (low FRET) yields kclose, the rate of PCNA closing (Table 4.1).

Interestingly, the rate of PCNA opening is affected by the ionic strength of the buffer. As the concentration of NaCl in solution increases, we observe a nearly linear increase in

-1 kopen from 0.0258 – 0.130 s (Figure 4.4 C). In contrast, we see no such trend for kclose.

All rates of closing for PCNA1-2-3 are approximately 0.038 – 0.051 s-1 and are not significantly perturbed by the variation in NaCl concentration. These rates correlate well with our histograms, for example at 1000 mM NaCl, we observe a nearly three-fold greater population of open PCNA vs. closed, which aligns well with kopen being nearly three-fold greater than kclose at this salt concentration, pushing the equilibrium towards favoring the open conformation of PCNA1-2-3. Notably, even at the fastest rate of

108 opening (0.130 s-1 at 1000 mM NaCl), PCNA is very slow in opening by itself. This may suggest why clamp loaders are necessary for DNA clamp loading.

4.3.4 Effect of Replication Factor C on the Equilibrium of PCNA Molecules

We established that the equilibrium between closed and open PCNA1-2-3 molecules can be affected by the concentration of NaCl in solution and that the percentage of PCNA1-2-3 molecules in the open state increases in a linear relationship with increasing salt (Figure 4.3F). Next, we investigated whether this equilibrium can be shifted by Replication Factor C, the ATPase driven clamp loader which opens the clamp to load it onto DNA. In a chamber with immobilized PCNA1-2-3 molecules, we introduced a reaction mixture containing 150 mM NaCl, 2 mM MgCl2, 1 μM RFC, and 5 mM ATP, in addition to the imaging buffer components. For this initial experiment, a large excess of RFC was used since lower concentrations of RFC did not appear to have any effect on our experiment. Notably, the Kd of Sso RFC and PCNA has not been measured. More experiments are required to establish the Kd and the optimal reaction conditions. Based on our previous results over a range of salt concentrations, at 150 mM

NaCl, only about 35% of PCNA1-2-3 molecules should be in the open conformation.

However, our results show that nearly 60% of PCNA1-2-3 molecules are in the open conformation, indicating that RFC can counteract the forces that govern the equilibrium between open and closed PCNA (Figure 4.5). These results might suggest why a clamp loader is necessary for DNA clamp loading in all organisms.169

4.4 Discussion

Studies of thermophilic organisms indicate that they have various adaptations

109 which allow them to stabilize their cellular proteins, and survive under extreme conditions. Thermostable proteins like those found in Sso have an increased number of salt bridge interactions, mainly at subunit interfaces, which likely stabilize protein structure and preserve biological activity.244 This is consistent with previous structural results that indicate that in addition to hydrogen bonds, there are many salt bridge interactions at the subunit interfaces of Sso PCNA.237 Crystallographic studies predict that of the three interfaces, the PCNA1:PCNA3 interface has the fewest salt bridges, supporting previous findings that this is the interface involved in closing and opening.

However, since we have covalently linked the PCNA2:3, and PCNA1:2 interface, we can be sure that the change in FRET we are observing is due to the opening of PCNA1:3.

Through smFRET studies we have demonstrated that the PCNA1-2-3 clamp can be both open and closed in solution (Figure 4.1, 4.2). Furthermore, we discovered that the equilibrium between the open and closed conformations can be perturbed by varying the ionic strength of our buffer. By increasing the concentration of NaCl in our smFRET experiments, we could disrupt the electrostatic interactions that are implicated in keeping the PCNA clamp closed thereby pushing the equilibrium to favor a more open form of

PCNA (Figure 4.3). Interestingly, this shift towards the open conformation of PCNA is due to a linear increase in the rate of PCNA opening, kopen, in relation to increasing NaCl concentration (Table 4.1). This supports our theory that by increasing the ionic strength of the buffer the electrostatic interactions at the PCNA1:3 interface are more readily disrupted. In contrast, the rate of PCNA closing, kclose, does not follow any trend in correlation with salt concentrations. One would expect an inverse relationship between

110 kclose and NaCl concentration, however this is not the case. Instead, kclose, remains in the same range over all salt concentrations (Table 4.1). Despite this, we see a large population of PCNA1-2-3 molecules in the closed conformation in low salt conditions.

This is because at the lower salt conditions, PCNA1-2-3 molecules start out in a closed conformation, and very few transition to the open state, since the disruption of electrostatic interactions in a low salt environment is unfavorable. To date, only closed structures of Sso PCNA have been crystallized,168,189,237 suggesting that x-ray crystallography selects for the closed conformation. Based on this we can speculate that the open form is either very flexible and dynamic, or unstable resulting in structural collapse. Furthermore, these previous studies suggest that PCNA prefers to be in a closed conformation, allowing all of the electrostatic interactions at the interfaces to be satisfied, and that the clamp is most stable in the closed form. Interestingly, this is not the first time that a DNA clamp which has only been crystallized closed, was discovered to exist in an open conformation in solution. Previous fluorescence studies of the gp45 trimer from T4 bacteriophage have revealed that the gp45 trimer exists primarily in an open conformation in solution.240

The solution dynamics of DNA clamp opening and closing are believed to be governed by a single reversible step between the closed (PCNAC) and open (PCNAO) form of PCNA (Scheme 4.1A). Here, we propose that the mechanism of clamp opening and closing are both two-step mechanisms (Scheme 4.1B). Based on our smFRET results, we believe that the electrostatic interactions at the PCNA1:3 interface serve to

“lock” the clamp in a closed form. Thus, the first step of PCNA opening, k1, is the

111 disruption of electrostatic interactions at the PCNA1:3 interface, resulting in a closed

PCNA which no longer has interactions between the PCNA1:3 interface holding it

Z together (PCNA ). The second step of opening, k2 is the conformational opening of

PCNA due to a lack of electrostatic interactions at the PCNA1:3 interactions resulting.

We observed that kopen increases with NaCl concentration. Since increase in NaCl results in disruption of electrostatic interactions, this dependence suggests that the first step of clamp opening, the disruption of electrostatic charges, is the rate determining step in

DNA clamp opening, while k2 the conformational opening is rapid. Furthermore, the unaffected rates of PCNA closing, kclose, in response to varying NaCl concentration demonstrate that once it is open, the PCNA clamp does not readily close in response to lowering salt concentration. This is likely because to close, PCNA must undergo a conformational change, to bring the interfaces of PCNA1 and 3 together to interact.

PCNA is not known to exhibit conformational dynamics by itself in vivo in any organism.

Based on this finding, we believe that in the mechanism of clamp closing (Scheme 4.1B) the rate is limited by the conformational change necessary to bring the interfaces

-1 together, k-2. The average kclose of 0.045 s observed for all concentrations of NaCl, is likely equivalent to k-2, the rate of the conformational change wherein the open PCNA closes to bring the PCNA1:3 interface together. As expected, a conformational change step would not be affected by changes in NaCl concentration, thus we observe a relatively unchanged rate of kclose. The second step of closing k-1 represents the electrostatic interactions being formed, however that is not observable by FRET in our experiment.

112

Physiologically it would not be favorable for any organism to undergo vast changes in intracellular salt concentration to complete biological processes. Thus, other mechanisms must exist for the opening of closed DNA clamps in vivo. The clamp must be opened, loaded onto DNA, and closed to allow for DNA replication. For this, all organisms possess evolutionarily conserved clamp loaders. These ATPase driven pentameric complexes bind DNA clamps, and load them onto DNA at the primer template junction and dissociate.169,238,239 However, the nuances of this process are still a matter of debate. One hypothesis suggests that the clamp•clamp loader complex hydrolyzes ATP to open the DNA clamp, overcoming the energy needed to break electrostatic interactions at the clamp opening interface. An alternate hypothesis, is that the clamp loader binds ATP first, stabilizing a conformation of the clamp loader which allows it to preferentially bind an open clamp. This clamp•clamp loader complex then binds DNA, which induces the hydrolysis of ATP to close the clamp, and release the clamp loader from the loaded clamp. In our experiments, closer to physiological salt conditions (250 mM NaCl) nearly 40% of PCNA trimers are in the open conformation.

Based on this, we can speculate that this latter mechanism of clamp loading might be applicable to the Sso system wherein, the ATP bound clamp loader can bind open clamps and load them onto DNA. Interestingly, based on FRET measurements we can estimate that in the open conformation the distance between the PCNA1 and PCNA3 subunits is approximately 10 Å, which is similar to a previous model of open Sso PCNA which predicts a 6-7 Å gap due to a 10⁰opening observed at the PCNA1:PCNA2 interface in the absence of PCNA3.237 Based on this finding, and the knowledge that the diameter of B-

113 form double stranded DNA is approximately 23 Å, we believe that open Sso PCNA is bound by a clamp loader, and loaded onto single stranded DNA at which point the closed clamp slides into place at the primer template junction.

To investigate whether the clamp loader is binding to available open clamps, or binding closed clamps and opening them, we conducted an smFRET experiment by introducing Sso clamp loader Replication Factor C, and ATP onto a slide containing immobilized PCNA1-2-3 molecules. A salt concentration of 150 mM NaCl was selected to simulate a more physiologically relevant cellular event. Based on our linear correlation of PCNA1-2-3 opening vs. salt concentration (Figure 4.3F) we estimate that approximately 35% of PCNA1-2-3 molecules should be in the open conformation at 150 mM NaCl. RFC was used in excess (1 μM) to ensure binding of RFC to the immobilized

PCNA1-2-3 molecules. Excess ATP (5 mM) was used to ensure that sufficient ATP was present since RFC is known to hydrolyze ATP even when it is not bound to a clamp or

DNA.180 Our results indicate that RFC bound the PCNA1-2-3 molecules in our smFRET experiment and shifted the equilibrium towards the open conformation of PCNA1-2-3 molecules. At 150 mM NaCl, we estimated that 35% of PCNA molecules would be in the open state. With RFC and ATP in the smFRET experiment, we observed that nearly 60% of PCNA1-2-3 molecules were in the open state, nearly doubling the population of open

DNA clamps. Based on this finding, we report that RFC was able to bind closed DNA clamps and open them for loading, giving merit to the hypothesis that in Sso, the clamp loader is actually responsible for the opening of the PCNA and not just loading alone.

114

In conclusion, we have demonstrated that the heterotrimeric DNA clamp PCNA in Sulfolobus solfataricus is stabilized in the closed conformation by electrostatic interactions, and that the interactions can be perturbed by varying salt concentrations. We completed a thorough investigation of the range of salt conditions which affect this equilibrium, and established that PCNA opening correlates linearly with increasing NaCl concentration. Based on the FRET data we observed a 10 Å opening at the

PCNA1:PCNA3 interface, indicating that the PCNA molecules are likely loaded onto single stranded DNA before sliding into position at the primer template junction. Dwell time analysis of the smFRET data revealed that PCNA opening and closing are very slow processes, which might suggest why a clamp loader is necessary when the clamps are already open in solution. Furthermore, we showed that Sso Replication Factor C, serves as a biological approach to perturbing the equilibrium between open and closed PCNA.

In our experiments with RFC we observe a shift in population of closed and open PCNA trimers indicating that in Sso RFC is involved in both the clamp opening and clamp loading steps. Finally, in the presence of RFC we did not observe greater opening of

PCNA supporting our hypothesis that PCNA is loaded first onto single stranded DNA.

Going forward, research needs to be conducted to elucidate the role of ATP during the

RFC PCNA opening.

Author Contributions

Varun V. Gadkari generated protein expression plasmids with the help of Anthony A.

Stephenson, purified all PCNA and RFC proteins, and collected most of salt- concentration dependent single molecule data. Austin T. Raper and V.V.G. conceived the

115 idea for surface-immobilizing PCNA through utilization of the Avi-tag technology, and performed fluorophore labeling and protein biotinylation to study PCNA dynamics.

V.V.G obtained and purified the necessary proteins to perform enzymatic biotinylation, and designed the site-specific FRET system to report on clamp opening and closing.

V.V.G. collected, and analyzed most of the data, with some assistance from A.T.R..

V.V.G wrote this chapter. Dr. Zucai Suo conceived the research and kinetic mechanism.

116

4.5 Schemes

Scheme 4.1: Proposed Mechanism of PCNA Opening and Closing

117

4.6 Tables

Table 4.1 Rates of PCNA opening and closing

-1 -1 NaCl (mM) kopen (s ) kclose (s ) Keq (kopen/kclose)

0 0.0258 ± 0.0004 0.044 ± 0.004 0.59

250 0.0374 ± 0.0007 0.050 ± 0.002 0.75

500 0.0476 ± 0.0008 0.0405 ± 0.0009 1.18

750 0.081 ± 0.002 0.051 ± 0.002 1.59

1,000 0.130 ± 0.002 0.038 ± 0.002 3.42

118

4.7 Figures

Figure 4.1: Design of a two state FRET system (A) PCNA1-2-3 is covalently linked at the PCNA1:2, and PCNA2:3 interface. The

PCNA1:3 interface where the clamp is known to open, is doubly labeled with a Cy3 donor and Cy5 acceptor fluorescence dyes. The closed form of PCNA1-2-3 positions the dyes close together allowing for a high FRET efficiency. (B) The opening of PCNA moves the donor and acceptor dyes apart reducing the FRET efficiency. (C) The closed

PCNA1-2-3 molecules produce a high FRET efficiency signal in the 0.6-0.8 range. (D)

The open PCNA1-2-3 molecules produce a low FRET efficiency signal in the 0.4-0.6 range.

119

Figure 4.2: PCNA interconverts between its open and closed conformations. (A) Representative donor (green) and acceptor (red) fluorescence trajectories depicting transition from the PCNA closed (high-FRET) to open (low-FRET) conformation before donor photobleaching. (B) FRET efficiency trajectory (blue) calculated from the donor and acceptor intensities in (A). (C) Representative donor (green) and acceptor (red) fluorescence trajectories depicting transition from the PCNA open (low-FRET) to closed

(high-FRET) conformation before acceptor photobleaching. (D) FRET efficiency trajectory (blue) calculated from the donor and acceptor intensities in (C).

120

Figure 4.1: FRET efficiency population histograms in changing concentrations of NaCl Apparent FRET traces for >200 molecules are compiled, and binned based on FRET efficiency to generate histograms that represent the frequency of occurrence for various

FRET efficiencies. In this investigation, immobilized PCNA1-2-3 molecules were observed in two conformations resulting in two distinct FRET states. The low FRET state

(0.4-0.6) represents open PCNA1-2-3, and a high FRET state (0.6-0.8) represents closed

PCNA1-2-3. Single-molecule FRET experiments were conducted at (A) 0 mM, (B) 250 mM, (C) 500 mM, (D) 750 mM, and (E) 1000 mM NaCl. (F) We observed a salt dependent shift in the equilibrium. The fraction of PCNA1-2-3 molecules in the open state increased in linear correlation with an increase in concentration of NaCl.

121

Figure 4.2: Dwell Time Analysis

Rates of PCNA (A) opening (kopen) and (B) closing (kclose) at each NaCl concentration were extracted by fitting the dwell time survivor functions of the high-FRET and low-

FRET states, respectively, to a single exponential decay equation. (C) Rates of PCNA opening (kopen) were plotted against NaCl concentrations. A linear fit with slope of 1.0 x

-4 -1 -1 10 mM s is shown (black line) to describe the dependence of kopen values on NaCl concentrations.

122

Figure 4.3: FRET distribution of PCNA1-2-3 in the presence of RFC PCNA molecules were immobilized in imaging buffer containing 150 mM NaCl. After sufficient molecules were immobilized, the excess molecules were washed out, and a reaction mix containing imaging buffer, 150 mM NaCl, 2 mM MgCl2, 1 μM RFC, and 5 mM ATP was flowed into the single-molecule slide. Based on the results in Figure 4.3 we expected that ~35% of PCNA1-2-3 molecules would be in the open conformation in a buffer containing 150 mM NaCl. Instead, we observed that nearly 60% of molecules observed were in the low FRET state corresponding to the open conformation, indicating that Replication Factor C was able to affect the population of open PCNA trimer.

123

References

1. Lehman, I.R., Bessman, M.J., Simms, E.S. & Kornberg, A. Enzymatic synthesis of deoxyribonucleic acid. I. Preparation of substrates and partial purification of an enzyme from Escherichia coli. J Biol Chem 233, 163-70 (1958). 2. Bessman, M.J., Lehman, I.R., Simms, E.S. & Kornberg, A. Enzymatic synthesis of deoxyribonucleic acid. II. General properties of the reaction. J Biol Chem 233, 171-7 (1958). 3. Bessman, M.J. et al. Enzymatic Synthesis of Deoxyribonucleic Acid. Iii. The Incorporation of Pyrimidine and Purine Analogues into Deoxyribonucleic Acid. Proc Natl Acad Sci U S A 44, 633-40 (1958). 4. Adler, J., Lehman, I.R., Bessman, M.J., Simms, E.S. & Kornberg, A. Enzymatic Synthesis of Deoxyribonucleic Acid. Iv. Linkage of Single Deoxynucleotides to the Deoxynucleoside Ends of Deoxyribonucleic Acid. Proc Natl Acad Sci U S A 44, 641-7 (1958). 5. Lehman, I.R. et al. Enzymatic Synthesis of Deoxyribonucleic Acid. V. Chemical Composition of Enzymatically Synthesized Deoxyribonucleic Acid. Proc Natl Acad Sci U S A 44, 1191-6 (1958). 6. Ollis, D.L., Brick, P., Hamlin, R., Xuong, N.G. & Steitz, T.A. Structure of large fragment of Escherichia coli DNA polymerase I complexed with dTMP. Nature 313, 762-6 (1985). 7. Ito, J. & Braithwaite, D.K. Compilation and alignment of DNA polymerase sequences. Nucleic Acids Res 19, 4045-57 (1991). 8. Braithwaite, D.K. & Ito, J. Compilation, alignment, and phylogenetic relationships of DNA polymerases. Nucleic Acids Res 21, 787-802 (1993). 9. Kiefer, J.R. et al. Crystal structure of a thermostable Bacillus DNA polymerase I large fragment at 2.1 A resolution. Structure 5, 95-108 (1997). 10. Pelletier, H., Sawaya, M.R., Wolfle, W., Wilson, S.H. & Kraut, J. Crystal structures of human DNA polymerase beta complexed with DNA: implications for catalytic mechanism, processivity, and fidelity. Biochemistry 35, 12742-61 (1996). 11. Eom, S.H., Wang, J. & Steitz, T.A. Structure of Taq polymerase with DNA at the polymerase active site. Nature 382, 278-81 (1996). 12. Kim, Y. et al. Crystal structure of Thermus aquaticus DNA polymerase. Nature 376, 612-6 (1995). 13. Sawaya, M.R., Pelletier, H., Kumar, A., Wilson, S.H. & Kraut, J. Crystal structure of rat DNA polymerase beta: evidence for a common polymerase mechanism. Science 264, 1930-5 (1994).

124

14. Pelletier, H., Sawaya, M.R., Kumar, A., Wilson, S.H. & Kraut, J. Structures of ternary complexes of rat DNA polymerase beta, a DNA template-primer, and ddCTP. Science 264, 1891-903 (1994). 15. Davies, J.F., 2nd, Almassy, R.J., Hostomska, Z., Ferre, R.A. & Hostomsky, Z. 2.3 A crystal structure of the catalytic domain of DNA polymerase beta. Cell 76, 1123-33 (1994). 16. Steitz, T.A. et al. Two DNA polymerases: HIV reverse transcriptase and the Klenow fragment of Escherichia coli DNA polymerase I. Cold Spring Harb Symp Quant Biol 58, 495-504 (1993). 17. Beese, L.S., Friedman, J.M. & Steitz, T.A. Crystal structures of the Klenow fragment of DNA polymerase I complexed with deoxynucleoside triphosphate and pyrophosphate. Biochemistry 32, 14095-101 (1993). 18. Wang, J. et al. Crystal structure of a pol alpha family replication DNA polymerase from bacteriophage RB69. Cell 89, 1087-99 (1997). 19. Doublie, S., Tabor, S., Long, A.M., Richardson, C.C. & Ellenberger, T. Crystal structure of a bacteriophage T7 DNA replication complex at 2.2 A resolution. Nature 391, 251-8 (1998). 20. Beese, L.S., Derbyshire, V. & Steitz, T.A. Structure of DNA polymerase I Klenow fragment bound to duplex DNA. Science 260, 352-5 (1993). 21. Kohlstaedt, L.A., Wang, J., Friedman, J.M., Rice, P.A. & Steitz, T.A. Crystal structure at 3.5 A resolution of HIV-1 reverse transcriptase complexed with an inhibitor. Science 256, 1783-90 (1992). 22. Bryant, F.R., Johnson, K.A. & Benkovic, S.J. Elementary steps in the DNA polymerase I reaction pathway. Biochemistry 22, 3537-46 (1983). 23. Mizrahi, V., Henrie, R.N., Marlier, J.F., Johnson, K.A. & Benkovic, S.J. Rate- limiting steps in the DNA polymerase I reaction pathway. Biochemistry 24, 4010- 8 (1985). 24. Mizrahi, V., Benkovic, P. & Benkovic, S.J. Mechanism of DNA polymerase I: exonuclease/polymerase activity switch and DNA sequence dependence of pyrophosphorolysis and misincorporation reactions. Proc Natl Acad Sci U S A 83, 5769-73 (1986). 25. Mizrahi, V., Benkovic, P.A. & Benkovic, S.J. Mechanism of the idling-turnover reaction of the large (Klenow) fragment of Escherichia coli DNA polymerase I. Proc Natl Acad Sci U S A 83, 231-5 (1986). 26. Kuchta, R.D., Mizrahi, V., Benkovic, P.A., Johnson, K.A. & Benkovic, S.J. Kinetic mechanism of DNA polymerase I (Klenow). Biochemistry 26, 8410-7 (1987). 27. Kuchta, R.D., Benkovic, P. & Benkovic, S.J. Kinetic mechanism whereby DNA polymerase I (Klenow) replicates DNA with high fidelity. Biochemistry 27, 6716- 25 (1988). 28. Mizrahi, V. & Benkovic, S.J. The dynamics of DNA polymerase-catalyzed reactions. Adv Enzymol Relat Areas Mol Biol 61, 437-57 (1988).

125

29. Dahlberg, M.E. & Benkovic, S.J. Kinetic mechanism of DNA polymerase I (Klenow fragment): identification of a second conformational change and evaluation of the internal equilibrium constant. Biochemistry 30, 4835-43 (1991). 30. Eger, B.T. & Benkovic, S.J. Minimal kinetic mechanism for misincorporation by DNA polymerase I (Klenow fragment). Biochemistry 31, 9227-36 (1992). 31. Joyce, C.M. & Benkovic, S.J. DNA polymerase fidelity: kinetics, structure, and checkpoints. Biochemistry 43, 14317-24 (2004). 32. Donlin, M.J., Patel, S.S. & Johnson, K.A. Kinetic partitioning between the exonuclease and polymerase sites in DNA error correction. Biochemistry 30, 538- 46 (1991). 33. Wong, I., Patel, S.S. & Johnson, K.A. An induced-fit kinetic mechanism for DNA replication fidelity: direct measurement by single-turnover kinetics. Biochemistry 30, 526-37 (1991). 34. Kati, W.M., Johnson, K.A., Jerva, L.F. & Anderson, K.S. Mechanism and fidelity of HIV reverse transcriptase. J Biol Chem 267, 25988-97 (1992). 35. Johnson, K.A. The kinetic and chemical mechanism of high-fidelity DNA polymerases. Biochim Biophys Acta 1804, 1041-1048 (2010). 36. Patel, S.S., Wong, I. & Johnson, K.A. Pre-steady-state kinetic analysis of processive DNA replication including complete characterization of an exonuclease-deficient mutant. Biochemistry 30, 511-25 (1991). 37. Steitz, T.A. A mechanism for all polymerases. Nature 391, 231-2 (1998). 38. Burgers, P.M. et al. Eukaryotic DNA polymerases: proposal for a revised nomenclature. J Biol Chem 276, 43487-90 (2001). 39. Freemont, P.S., Friedman, J.M., Beese, L.S., Sanderson, M.R. & Steitz, T.A. Cocrystal structure of an editing complex of Klenow fragment with DNA. Proc Natl Acad Sci U S A 85, 8924-8 (1988). 40. Beese, L.S. & Steitz, T.A. Structural basis for the 3'-5' exonuclease activity of Escherichia coli DNA polymerase I: a two metal ion mechanism. Embo J 10, 25- 33 (1991). 41. Zhang, L., Brown, J.A., Newmister, S.A. & Suo, Z. Polymerization fidelity of a replicative DNA polymerase from the hyperthermophilic archaeon Sulfolobus solfataricus P2. Biochemistry 48, 7492-501 (2009). 42. Ling, H., Boudsocq, F., Woodgate, R. & Yang, W. Crystal structure of a Y-family DNA polymerase in action: a mechanism for error-prone and lesion-bypass replication. Cell 107, 91-102 (2001). 43. Boudsocq, F. et al. Investigating the role of the little finger domain of Y-family DNA polymerases in low fidelity synthesis and translesion replication. J Biol Chem 279, 32932-40 (2004). 44. Friedberg, E.C., Wagner, R. & Radman, M. Specialized DNA polymerases, cellular survival, and the genesis of mutations. Science 296, 1627-30 (2002). 45. Maxwell, B.A. & Suo, Z. Kinetic basis for the differing response to an oxidative lesion by a replicative and a lesion bypass DNA polymerase from Sulfolobus solfataricus. Biochemistry 51, 3485-96 (2012). 46. Ohmori, H. et al. The Y-family of DNA polymerases. Mol Cell 8, 7-8 (2001). 126

47. Friedberg, E.C. & Gerlach, V.L. Novel DNA polymerases offer clues to the molecular basis of mutagenesis. Cell 98, 413-6 (1999). 48. Woodgate, R. A plethora of lesion-replicating DNA polymerases. Genes Dev 13, 2191-5 (1999). 49. Fiala, K.A., Hypes, C.D. & Suo, Z. Mechanism of abasic lesion bypass catalyzed by a Y-family DNA polymerase. J Biol Chem 282, 8188-98 (2007). 50. Fiala, K.A. & Suo, Z. Sloppy bypass of an abasic lesion catalyzed by a Y-family DNA polymerase. J Biol Chem 282, 8199-206 (2007). 51. Brown, J.A., Newmister, S.A., Fiala, K.A. & Suo, Z. Mechanism of double-base lesion bypass catalyzed by a Y-family DNA polymerase. Nucleic Acids Res 36, 3867-78 (2008). 52. Sherrer, S.M. et al. Mechanistic Studies of the Bypass of a Bulky Single-base Lesion Catalyzed by a Y-family DNA Polymerase. J Biol Chem 284, 6379-88 (2009). 53. Sherrer, S.M. et al. Kinetic analysis of the bypass of a bulky DNA lesion catalyzed by human Y-family DNA polymerases. Chem Res Toxicol 25, 730-40 (2012). 54. Gadkari, V.V., Tokarsky, E.J., Malik, C.K., Basu, A.K. & Suo, Z. Mechanistic investigation of the bypass of a bulky aromatic DNA adduct catalyzed by a Y- family DNA polymerase. DNA Repair (Amst) 21, 65-77 (2014). 55. Raper, A.T., Gadkari, V.V., Maxwell, B.A. & Suo, Z. Single-Molecule Investigation of Response to Oxidative DNA Damage by a Y-Family DNA Polymerase. Biochemistry 55, 2187-96 (2016). 56. Tokarsky, E.J. et al. Pre-steady-state kinetic investigation of bypass of a bulky guanine lesion by human Y-family DNA polymerases. DNA Repair (Amst) 46, 20-28 (2016). 57. Vyas, R. et al. Mechanistic Basis for the Bypass of a Bulky DNA Adduct Catalyzed by a Y-Family DNA Polymerase. J Am Chem Soc 137, 12131-42 (2015). 58. Patra, A. et al. Mechanism of Error-Free Bypass of the Environmental Carcinogen N-(2'-Deoxyguanosin-8-yl)-3-aminobenzanthrone Adduct by Human DNA Polymerase eta. Chembiochem 17, 2033-2037 (2016). 59. Jha, V. & Ling, H. Structural basis of accurate replication beyond a bulky major benzo[a]pyrene adduct by human DNA polymerase kappa. DNA Repair (Amst) 49, 43-50 (2017). 60. Trincao, J. et al. Structure of the catalytic core of S. cerevisiae DNA polymerase eta: implications for translesion DNA synthesis. Mol Cell 8, 417-26 (2001). 61. Chu, X. et al. 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 (2014). 62. Yang, W. & Woodgate, R. What a difference a decade makes: insights into translesion DNA synthesis. Proc Natl Acad Sci U S A 104, 15591-8 (2007).

127

63. Gaur, V. et al. Structural and kinetic insights into binding and incorporation of L- nucleotide analogs by a Y-family DNA polymerase. Nucleic Acids Res 42, 9984- 95 (2014). 64. Brown, J.A., Pack, L.R., Fowler, J.D. & Suo, Z. 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 (2011). 65. Brown, J.A., Pack, L.R., Fowler, J.D. & Suo, Z. 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 (2012). 66. She, Q. et al. The complete genome of the crenarchaeon Sulfolobus solfataricus P2. Proc Natl Acad Sci U S A 98, 7835-40 (2001). 67. Fiala, K.A., Sherrer, S.M., Brown, J.A. & Suo, Z. Mechanistic consequences of temperature on DNA polymerization catalyzed by a Y-family DNA polymerase. Nucleic Acids Res 36, 1990-2001 (2008). 68. Wong, J.H., Fiala, K.A., Suo, Z. & Ling, H. Snapshots of a Y-family DNA polymerase in replication: substrate-induced conformational transitions and implications for fidelity of Dpo4. J Mol Biol 379, 317-30 (2008). 69. Xu, C., Maxwell, B.A., Brown, J.A., Zhang, L. & Suo, Z. Global conformational dynamics of a Y-family DNA polymerase during catalysis. PLoS Biol 7, e1000225 (2009). 70. Wong, J.H. et al. Structural insight into dynamic bypass of the major cisplatin- DNA adduct by Y-family polymerase Dpo4. EMBO J 29, 2059-69 (2010). 71. Maxwell, B.A., Xu, C. & Suo, Z. DNA Lesion Alters Global Conformational Dynamics of Y-family DNA Polymerase during Catalysis. J Biol Chem 287, 13040-7 (2012). 72. Sherrer, S.M. et al. Quantitative analysis of the mutagenic potential of 1- aminopyrene-DNA adduct bypass catalyzed by Y-family DNA polymerases. Mutat Res 737, 25-33 (2012). 73. Brenlla, A., Markiewicz, R.P., Rueda, D. & Romano, L.J. Nucleotide selection by the Y-family DNA polymerase Dpo4 involves template translocation and misalignment. Nucleic Acids Res 42, 2555-63 (2014). 74. Maxwell, B.A. & Suo, Z. Recent insight into the kinetic mechanisms and conformational dynamics of Y-Family DNA polymerases. Biochemistry 53, 2804- 14 (2014). 75. Maxwell, B.A., Xu, C. & Suo, Z. Conformational dynamics of a Y-family DNA polymerase during substrate binding and catalysis as revealed by interdomain Forster resonance energy transfer. Biochemistry 53, 1768-78 (2014). 76. Raper, A.T. & Suo, Z. Investigation of Intradomain Motions of a Y-Family DNA Polymerase during Substrate Binding and Catalysis. Biochemistry 55, 5832-5844 (2016). 77. Fiala, K.A. & Suo, Z. Pre-Steady-State Kinetic Studies of the Fidelity of Sulfolobus solfataricus P2 DNA Polymerase IV. Biochemistry 43, 2106-15 (2004). 128

78. Fiala, K.A. & Suo, Z. Mechanism of DNA Polymerization Catalyzed by Sulfolobus solfataricus P2 DNA Polymerase IV. Biochemistry 43, 2116-25 (2004). 79. Fiala, K.A. et al. Mechanism of Template-independent Nucleotide Incorporation Catalyzed by a Template-dependent DNA Polymerase. J Mol Biol 365, 590-602 (2007). 80. Gowda, A.S., Krishnegowda, G., Suo, Z., Amin, S. & Spratt, T.E. Low fidelity bypass of O(2)-(3-pyridyl)-4-oxobutylthymine, the most persistent bulky adduct produced by the tobacco specific nitrosamine 4-(methylnitrosamino)-1-(3- pyridyl)-1-butanone by model DNA polymerases. Chem Res Toxicol 25, 1195- 202 (2012). 81. Sherrer, S.M. et al. Identification of an unfolding intermediate for a DNA lesion bypass polymerase. Chem Res Toxicol 25, 1531-40 (2012). 82. Ling, H., Boudsocq, F., Plosky, B.S., Woodgate, R. & Yang, W. Replication of a cis-syn thymine dimer at atomic resolution. Nature 424, 1083-7 (2003). 83. Ling, H. et al. Crystal structure of a benzo[a]pyrene diol epoxide adduct in a ternary complex with a DNA polymerase. Proc Natl Acad Sci U S A 101, 2265-9 (2004). 84. Bauer, J. et al. 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 (2007). 85. Xing, G., Kirouac, K., Shin, Y.J., Bell, S.D. & Ling, H. Structural insight into recruitment of translesion DNA polymerase Dpo4 to sliding clamp PCNA. Mol Microbiol 71, 678-91 (2009). 86. Boudsocq, F., Iwai, S., Hanaoka, F. & Woodgate, R. 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 (2001). 87. Zang, H. et al. DNA adduct bypass polymerization by Sulfolobus solfataricus DNA polymerase Dpo4: analysis and crystal structures of multiple base pair substitution and frameshift products with the adduct 1,N2-ethenoguanine. J Biol Chem 280, 29750-64 (2005). 88. Irimia, A. et al. Calcium is a cofactor of polymerization but inhibits pyrophosphorolysis by the Sulfolobus solfataricus DNA polymerase Dpo4. Biochemistry 45, 5949-56 (2006). 89. Zang, H., Chowdhury, G., Angel, K.C., Harris, T.M. & Guengerich, F.P. Translesion synthesis across polycyclic aromatic hydrocarbon diol epoxide adducts of deoxyadenosine by Sulfolobus solfataricus DNA polymerase Dpo4. Chem Res Toxicol 19, 859-67 (2006). 90. Zang, H. et al. Efficient and High Fidelity Incorporation of dCTP Opposite 7,8- Dihydro-8-oxodeoxyguanosine by Sulfolobus solfataricus DNA Polymerase Dpo4. J Biol Chem 281, 2358-72 (2006). 91. Eoff, R.L., Angel, K.C., Egli, M. & Guengerich, F.P. Molecular basis of selectivity of nucleoside triphosphate incorporation opposite O6-benzylguanine 129

by sulfolobus solfataricus DNA polymerase Dpo4: steady-state and pre-steady- state kinetics and x-ray crystallography of correct and incorrect pairing. J Biol Chem 282, 13573-84 (2007). 92. Eoff, R.L., Irimia, A., Angel, K.C., Egli, M. & Guengerich, F.P. Hydrogen bonding of 7,8-dihydro-8-oxodeoxyguanosine with a charged residue in the little finger domain determines miscoding events in Sulfolobus solfataricus DNA polymerase Dpo4. J Biol Chem 282, 19831-43 (2007). 93. Eoff, R.L., Irimia, A., Egli, M. & Guengerich, F.P. Sulfolobus solfataricus DNA polymerase Dpo4 is partially inhibited by "wobble" pairing between O6- methylguanine and cytosine, but accurate bypass is preferred. J Biol Chem 282, 1456-67 (2007). 94. Choi, J.Y. et al. Roles of the four DNA polymerases of the crenarchaeon Sulfolobus solfataricus and accessory proteins in DNA replication. J Biol Chem 286, 31180-93 (2011). 95. Goodenough, A.K. et al. Site specific synthesis and polymerase bypass of oligonucleotides containing a 6-hydroxy-3,5,6,7-tetrahydro-9H-imidazo[1,2- a]purin-9-one base, an intermediate in the formation of 1,N2-etheno-2'- deoxyguanosine. Chem Res Toxicol 18, 1701-14 (2005). 96. Irimia, A., Eoff, R.L., Pallan, P.S., Guengerich, F.P. & Egli, M. Structure and activity of Y-class DNA polymerase DPO4 from Sulfolobus solfataricus with templates containing the hydrophobic thymine analog 2,4-difluorotoluene. J Biol Chem 282, 36421-33 (2007). 97. Beckman, J.W., Wang, Q. & Guengerich, F.P. Kinetic analysis of correct nucleotide insertion by a Y-family DNA polymerase reveals conformational changes both prior to and following phosphodiester bond formation as detected by tryptophan fluorescence. J Biol Chem 283, 36711-23 (2008). 98. Zhang, H. et al. Versatility of Y-family Sulfolobus solfataricus DNA polymerase Dpo4 in translesion synthesis past bulky N2-alkylguanine adducts. J Biol Chem 284, 3563-76 (2009). 99. Christov, P.P., Angel, K.C., Guengerich, F.P. & Rizzo, C.J. Replication past the N5-methyl-formamidopyrimidine lesion of deoxyguanosine by DNA polymerases and an improved procedure for sequence analysis of in vitro bypass products by mass spectrometry. Chem Res Toxicol 22, 1086-95 (2009). 100. Eoff, R.L. et al. Structural and functional analysis of Sulfolobus solfataricus Y- family DNA polymerase Dpo4-catalyzed bypass of the malondialdehyde- deoxyguanosine adduct. Biochemistry 48, 7079-88 (2009). 101. Eoff, R.L., Sanchez-Ponce, R. & Guengerich, F.P. Conformational changes during nucleotide selection by Sulfolobus solfataricus DNA polymerase Dpo4. J Biol Chem 284, 21090-9 (2009). 102. Zhang, H. et al. Structure-function relationships in miscoding by Sulfolobus solfataricus DNA polymerase Dpo4: guanine N2,N2-dimethyl substitution produces inactive and miscoding polymerase complexes. J Biol Chem 284, 17687-99 (2009).

130

103. Zhang, H., Beckman, J.W. & Guengerich, F.P. Frameshift deletion by Sulfolobus solfataricus P2 DNA polymerase Dpo4 T239W is selective for purines and involves normal conformational change followed by slow phosphodiester bond formation. J Biol Chem 284, 35144-53 (2009). 104. Zhang, H. & Guengerich, F.P. Effect of N2-guanyl modifications on early steps in catalysis of polymerization by Sulfolobus solfataricus P2 DNA polymerase Dpo4 T239W. J Mol Biol 395, 1007-18 (2010). 105. Christov, P.P. et al. The C8-2'-deoxyguanosine adduct of 2-amino-3- methylimidazo[1,2-d]naphthalene, a carbocyclic analogue of the potent mutagen 2-amino-3-methylimidazo[4,5-f]quinoline, is a block to replication in vitro. Chem Res Toxicol 23, 1076-88 (2010). 106. Christov, P.P. et al. Comparison of the in vitro replication of the 7-(2-oxoheptyl)- 1,N2-etheno-2'-deoxyguanosine and 1,N2-etheno-2'-deoxyguanosine lesions by Sulfolobus solfataricus P2 DNA polymerase IV (Dpo4). Chem Res Toxicol 23, 1330-41 (2010). 107. Irimia, A., Loukachevitch, L.V., Eoff, R.L., Guengerich, F.P. & Egli, M. Metal- ion dependence of the active-site conformation of the translesion DNA polymerase Dpo4 from Sulfolobus solfataricus. Acta Crystallogr Sect F Struct Biol Cryst Commun 66, 1013-8 (2010). 108. Kotapati, S. et al. Translesion synthesis across 1,N6-(2-hydroxy-3- hydroxymethylpropan-1,3-diyl)-2'-deoxyadenosine (1,N6-gamma-HMHP-dA) adducts by human and archebacterial DNA polymerases. J Biol Chem 287, 38800-11 (2012). 109. Cho, S.H. & Guengerich, F.P. Replication past the butadiene diepoxide-derived DNA adduct S-[4-(N(6)-deoxyadenosinyl)-2,3-dihydroxybutyl]glutathione by DNA polymerases. Chem Res Toxicol 26, 1005-13 (2013). 110. Stornetta, A., Angelov, T., Guengerich, F.P. & Sturla, S.J. Incorporation of nucleoside probes opposite O(6)-methylguanine by Sulfolobus solfataricus DNA polymerase Dpo4: importance of hydrogen bonding. Chembiochem 14, 1634-9 (2013). 111. Sweasy, J.B. Fidelity mechanisms of DNA polymerase beta. Prog Nucleic Acid Res Mol Biol 73, 137-69 (2003). 112. Washington, M.T., Carlson, K.D., Freudenthal, B.D. & Pryor, J.M. Variations on a theme: Eukaryotic Y-family DNA polymerases. Biochim Biophys Acta 1804, 1113-1123 (2010). 113. Garcia-Diaz, M., Bebenek, K., Krahn, J.M., Kunkel, T.A. & Pedersen, L.C. A closed conformation for the Pol lambda catalytic cycle. Nat Struct Mol Biol 12, 97-98 (2005). 114. Rothwell, P.J., Mitaksov, V. & Waksman, G. Motions of the fingers subdomain of klentaq1 are fast and not rate limiting: implications for the molecular basis of fidelity in DNA polymerases. Mol Cell 19, 345-55 (2005). 115. Rothwell, P.J. & Waksman, G. A pre-equilibrium before nucleotide binding limits fingers subdomain closure by Klentaq1. J Biol Chem 282, 28884-92 (2007).

131

116. Allen, W.J., Rothwell, P.J. & Waksman, G. An intramolecular FRET system monitors fingers subdomain opening in Klentaq1. Protein Sci 17, 401-8 (2008). 117. Joyce, C.M. et al. Fingers-closing and other rapid conformational changes in DNA polymerase I (Klenow fragment) and their role in nucleotide selectivity. Biochemistry 47, 6103-16 (2008). 118. Rothwell, P.J. et al. dNTP-dependent conformational transitions in the fingers subdomain of Klentaq1 DNA polymerase: insights into the role of the "nucleotide-binding" state. J Biol Chem 288, 13575-91 (2013). 119. Doublie, S., Sawaya, M.R. & Ellenberger, T. An open and closed case for all polymerases. Structure 7, R31-5 (1999). 120. Maxwell, B.A. & Suo, Z. Single-molecule investigation of substrate binding kinetics and protein conformational dynamics of a B-family replicative DNA polymerase. J Biol Chem 288, 11590-11600 (2013). 121. Xu, C., Maxwell, B.A. & Suo, Z. Conformational dynamics of Thermus aquaticus DNA polymerase I during catalysis. J Mol Biol 426, 2901-17 (2014). 122. Christian, T.D., Romano, L.J. & Rueda, D. Single-molecule measurements of synthesis by DNA polymerase with base-pair resolution. Proc Natl Acad Sci U S A 106, 21109-14 (2009). 123. Santoso, Y. et al. Conformational transitions in DNA polymerase I revealed by single-molecule FRET. Proc Natl Acad Sci U S A 107, 715-20 (2010). 124. Berezhna, S.Y., Gill, J.P., Lamichhane, R. & Millar, D.P. Single-molecule Forster resonance energy transfer reveals an innate fidelity checkpoint in DNA polymerase I. J Am Chem Soc 134, 11261-8 (2012). 125. Markiewicz, R.P., Vrtis, K.B., Rueda, D. & Romano, L.J. Single-molecule microscopy reveals new insights into nucleotide selection by DNA polymerase I. Nucleic Acids Res 40, 7975-84 (2012). 126. Li, Y., Korolev, S. & Waksman, G. Crystal structures of open and closed forms of binary and ternary complexes of the large fragment of Thermus aquaticus DNA polymerase I: structural basis for nucleotide incorporation. Embo J 17, 7514-25 (1998). 127. Kellinger, M.W. & Johnson, K.A. Nucleotide-dependent conformational change governs specificity and analog discrimination by HIV reverse transcriptase. Proc Natl Acad Sci U S A 107, 7734-9 (2010). 128. Tsai, Y.C. & Johnson, K.A. A new paradigm for DNA polymerase specificity. Biochemistry 45, 9675-87 (2006). 129. Zhong, X., Patel, S.S., Werneburg, B.G. & Tsai, M.D. DNA polymerase beta: multiple conformational changes in the mechanism of catalysis. Biochemistry 36, 11891-900 (1997). 130. Purohit, V., Grindley, N.D. & Joyce, C.M. Use of 2-aminopurine fluorescence to examine conformational changes during nucleotide incorporation by DNA polymerase I (Klenow fragment). Biochemistry 42, 10200-11 (2003). 131. Dunlap, C.A. & Tsai, M.D. Use of 2-aminopurine and tryptophan fluorescence as probes in kinetic analyses of DNA polymerase beta. Biochemistry 41, 11226-35 (2002). 132

132. Showalter, A.K. & Tsai, M.D. A reexamination of the nucleotide incorporation fidelity of DNA polymerases. Biochemistry 41, 10571-6 (2002). 133. Zhou, B.L., Pata, J.D. & Steitz, T.A. Crystal structure of a DinB lesion bypass DNA polymerase catalytic fragment reveals a classic polymerase catalytic domain. Mol Cell 8, 427-37 (2001). 134. Yang, W. Damage repair DNA polymerases Y. Curr Opin Struct Biol 13, 23-30 (2003). 135. DeLucia, A.M., Grindley, N.D. & Joyce, C.M. Conformational changes during normal and error-prone incorporation of nucleotides by a Y-family DNA polymerase detected by 2-aminopurine fluorescence. Biochemistry 46, 10790-803 (2007). 136. Alt, A. et al. Bypass of DNA lesions generated during anticancer treatment with cisplatin by DNA polymerase eta. Science 318, 967-70 (2007). 137. Lone, S. et al. Human DNA polymerase kappa encircles DNA: implications for mismatch extension and lesion bypass. Mol Cell 25, 601-14 (2007). 138. Kondratick, C.M., Washington, M.T., Prakash, S. & Prakash, L. Acidic residues critical for the activity and biological function of yeast DNA polymerase eta. Mol Cell Biol 21, 2018-25 (2001). 139. Minko, I.G., Washington, M.T., Prakash, L., Prakash, S. & Lloyd, R.S. Translesion DNA synthesis by yeast DNA polymerase eta on templates containing N2-guanine adducts of 1,3-butadiene metabolites. J Biol Chem 276, 2517-22 (2001). 140. Washington, M.T., Johnson, R.E., Prakash, L. & Prakash, S. Accuracy of lesion bypass by yeast and human DNA polymerase eta. Proc Natl Acad Sci U S A 98, 8355-60 (2001). 141. Washington, M.T., Johnson, R.E., Prakash, S. & Prakash, L. Mismatch extension ability of yeast and human DNA polymerase eta. J Biol Chem 276, 2263-6 (2001). 142. Washington, M.T., Prakash, L. & Prakash, S. Yeast DNA polymerase eta utilizes an induced-fit mechanism of nucleotide incorporation. Cell 107, 917-27 (2001). 143. Washington, M.T., Johnson, R.E., Prakash, L. & Prakash, S. The mechanism of nucleotide incorporation by human DNA polymerase eta differs from that of the yeast enzyme. Mol Cell Biol 23, 8316-22 (2003). 144. Washington, M.T., Prakash, L. & Prakash, S. Mechanism of nucleotide incorporation opposite a thymine-thymine dimer by yeast DNA polymerase eta. Proc Natl Acad Sci U S A 100, 12093-8 (2003). 145. Brown, J.A. et al. Pre-Steady-State Kinetic Analysis of Truncated and Full- Length Saccharomyces cerevisiae DNA Polymerase Eta. J Nucleic Acids 2010(2010). 146. Oh, S.M., Kim, H.R., Park, Y.J., Lee, S.Y. & Chung, K.H. Organic extracts of urban air pollution particulate matter (PM2.5)-induced genotoxicity and oxidative stress in human lung bronchial epithelial cells (BEAS-2B cells). Mutation Research 723, 142-51 (2011). 147. Arlt, V.M. et al. Environmental pollutant and potent mutagen 3-nitrobenzanthrone forms DNA adducts after reduction by NAD(P)H:quinone oxidoreductase and 133

conjugation by acetyltransferases and sulfotransferases in human hepatic cytosols. Cancer Res 65, 2644-52 (2005). 148. Bieler, C.A. et al. DNA adduct formation by the environmental contaminant 3- nitrobenzanthrone after intratracheal instillation in rats. Int J Cancer 116, 833-8 (2005). 149. Enya T., S.H., Watanabe T., Hirayama T., Hisamatsu Y., . 3-Nitrobenzanthrone a powerful bacterial mutagen and suspected human carcinogen found in diesel exhausts and airborne particulates. Environ. Sci. Technol. 31, 2772-2776 (1997). 150. IARC monographs on the evaluation of carcinogenic risks to humans. Diesel and gasoline engine exhausts and some nitroarenes. International Agency for Research on Cancer. IARC Monogr Eval Carcinog Risks Hum 46, 1-458 (1989). 151. Farmer, P.B. Diesel fuel and exhaust emissions: is there a human carcinogenic risk? Lancet 350, 1118 (1997). 152. EPA, U.S. Health Assessment Document for Diesel Engine Exhaust. (2002). 153. Arlt, V.M. et al. Activation of 3-nitrobenzanthrone and its metabolites by human acetyltransferases, sulfotransferases and cytochrome P450 expressed in Chinese hamster V79 cells. Int J Cancer 105, 583-92 (2003). 154. Arlt, V.M. et al. DNA adduct formation by the ubiquitous environmental pollutant 3-nitrobenzanthrone and its metabolites in rats. Biochem Biophys Res Commun 300, 107-14 (2003). 155. Bieler, C.A., Arlt, V.M., Wiessler, M. & Schmeiser, H.H. DNA adduct formation by the environmental contaminant 3-nitrobenzanthrone in V79 cells expressing human cytochrome P450 enzymes. Cancer Lett 200, 9-18 (2003). 156. Nagy, E. et al. DNA adduct and tumor formations in rats after intratracheal administration of the urban air pollutant 3-nitrobenzanthrone. Carcinogenesis 26, 1821-8 (2005). 157. Kawanishi, M., Kanno, T., Nishida, H., Takamura-Enya, T. & Yagi, T. Translesion DNA synthesis across various DNA adducts produced by 3- nitrobenzanthrone in Escherichia coli. Mutation Research 754, 32-8 (2013). 158. Oda, Y. et al. NMR studies of a DNA containing 8-hydroxydeoxyguanosine. Nucleic Acids Res 19, 1407-12 (1991). 159. Lipscomb, L.A. et al. X-ray structure of a DNA decamer containing 7,8-dihydro- 8-oxoguanine. Proc Natl Acad Sci U S A 92, 719-23 (1995). 160. McAuley-Hecht, K.E. et al. Crystal structure of a DNA duplex containing 8- hydroxydeoxyguanine-adenine base pairs. Biochemistry 33, 10266-70 (1994). 161. Beard, W.A., Batra, V.K. & Wilson, S.H. DNA polymerase structure-based insight on the mutagenic properties of 8-oxoguanine. Mutat Res 703, 18-23 (2010). 162. Brieba, L.G. et al. Structural basis for the dual coding potential of 8-oxoguanosine by a high-fidelity DNA polymerase. Embo J 23, 3452-61 (2004). 163. Rechkoblit, O. et al. Impact of conformational heterogeneity of OxoG lesions and their pairing partners on bypass fidelity by Y family polymerases. Structure 17, 725-36 (2009).

134

164. Brieba, L.G., Kokoska, R.J., Bebenek, K., Kunkel, T.A. & Ellenberger, T. A lysine residue in the fingers subdomain of T7 DNA polymerase modulates the miscoding potential of 8-oxo-7,8-dihydroguanosine. Structure 13, 1653-9 (2005). 165. Noble, E., Spiering, M.M. & Benkovic, S.J. Coordinated DNA Replication by the Bacteriophage T4 Replisome. Viruses 7, 3186-200 (2015). 166. Lewis, J.S., Jergic, S. & Dixon, N.E. The E. coli DNA Replication Fork. Enzymes 39, 31-88 (2016). 167. Zhang, D. & O'Donnell, M. The Eukaryotic Replication Machine. Enzymes 39, 191-229 (2016). 168. Dionne, I., Nookala, R.K., Jackson, S.P., Doherty, A.J. & Bell, S.D. A heterotrimeric PCNA in the hyperthermophilic archaeon Sulfolobus solfataricus. Mol Cell 11, 275-82 (2003). 169. Indiani, C. & O'Donnell, M. The replication clamp-loading machine at work in the three domains of life. Nat Rev Mol Cell Biol 7, 751-61 (2006). 170. Stukenberg, P.T., Studwell-Vaughan, P.S. & O'Donnell, M. Mechanism of the sliding beta-clamp of DNA polymerase III holoenzyme. J Biol Chem 266, 11328- 34 (1991). 171. Haracska, L. et al. Stimulation of DNA synthesis activity of human DNA polymerase kappa by PCNA. Mol Cell Biol 22, 784-91 (2002). 172. Garg, P., Stith, C.M., Majka, J. & Burgers, P.M. Proliferating cell nuclear antigen promotes translesion synthesis by DNA polymerase zeta. J Biol Chem 280, 23446-50 (2005). 173. Garg, P. & Burgers, P.M. Ubiquitinated proliferating cell nuclear antigen activates translesion DNA polymerases eta and REV1. Proc Natl Acad Sci U S A 102, 18361-6 (2005). 174. Pillon, M.C., Miller, J.H. & Guarne, A. The endonuclease domain of MutL interacts with the beta sliding clamp. DNA Repair (Amst) 10, 87-93 (2011). 175. Strzalka, W. & Ziemienowicz, A. Proliferating cell nuclear antigen (PCNA): a key factor in DNA replication and cell cycle regulation. Ann Bot 107, 1127-40 (2011). 176. Becherel, O.J., Fuchs, R.P. & Wagner, J. Pivotal role of the beta-clamp in translesion DNA synthesis and mutagenesis in E. coli cells. DNA Repair (Amst) 1, 703-8 (2002). 177. Mueser, T.C., Hinerman, J.M., Devos, J.M., Boyer, R.A. & Williams, K.J. Structural analysis of bacteriophage T4 DNA replication: a review in the Virology Journal series on bacteriophage T4 and its relatives. Virol J 7, 359 (2010). 178. Dore, A.S. et al. Structure of an archaeal PCNA1-PCNA2-FEN1 complex: elucidating PCNA subunit and client enzyme specificity. Nucleic Acids Res 34, 4515-26 (2006). 179. Pascal, J.M. et al. A flexible interface between DNA ligase and PCNA supports conformational switching and efficient ligation of DNA. Mol Cell 24, 279-91 (2006).

135

180. Bauer, R.J., Wolff, I.D., Zuo, X., Lin, H.K. & Trakselis, M.A. Assembly and distributive action of an archaeal DNA polymerase holoenzyme. J Mol Biol 425, 4820-36 (2013). 181. Savino, C. et al. Insights into DNA replication: the crystal structure of DNA polymerase B1 from the archaeon Sulfolobus solfataricus. Structure 12, 2001-8 (2004). 182. Kunkel, T.A. DNA replication fidelity. J Biol Chem 279, 16895-8 (2004). 183. Sharma, A., Kottur, J., Narayanan, N. & Nair, D.T. A strategically located serine residue is critical for the mutator activity of DNA polymerase IV from Escherichia coli. Nucleic Acids Res 41, 5104-14 (2013). 184. Biertumpfel, C. et al. Structure and mechanism of human DNA polymerase eta. Nature 465, 1044-8 (2010). 185. Silverstein, T.D. et al. Structural basis for error-free replication of oxidatively damaged DNA by yeast DNA polymerase eta. Structure 18, 1463-70 (2010). 186. Kong, X.P., Onrust, R., O'Donnell, M. & Kuriyan, J. Three-dimensional structure of the beta subunit of E. coli DNA polymerase III holoenzyme: a sliding DNA clamp. Cell 69, 425-37 (1992). 187. Moarefi, I., Jeruzalmi, D., Turner, J., O'Donnell, M. & Kuriyan, J. Crystal structure of the DNA polymerase processivity factor of T4 bacteriophage. J Mol Biol 296, 1215-23 (2000). 188. Gulbis, J.M., Kelman, Z., Hurwitz, J., O'Donnell, M. & Kuriyan, J. Structure of the C-terminal region of p21(WAF1/CIP1) complexed with human PCNA. Cell 87, 297-306 (1996). 189. Williams, G.J. et al. Structure of the heterotrimeric PCNA from Sulfolobus solfataricus. Acta Crystallogr Sect F Struct Biol Cryst Commun 62, 944-8 (2006). 190. Fowler, J.D. & Suo, Z. Biochemical, structural, and physiological characterization of terminal deoxynucleotidyl transferase. Chem Rev 106, 2092-110 (2006). 191. Gallager J., H.U., George M., Hendee L., Phillips D.H., Lewtas J. Formation of DNA adducts in rat lung following chronic inhalation of diesel emissions, carbon black and titanium dioxide particles. Carcinogenesis 15, 1291-1299 (1994). 192. Enya T., S.H., Watanabe T., Hirayama T., Hisamatsu Y., . 3-Nitrobenzanthrone a powerful bacterial mutagen and suspected human carcinogen found in diesel exhausts and airborne particulates. Environmental Science & Technology 31, 2772-2776 (1997). 193. Oda, Y., Zhang, Y., Buchinger, S., Reifferscheid, G. & Yang, M. Roles of human sulfotransferases in genotoxicity of carcinogens using genetically engineered umu test strains. Environ Mol Mutagen 53, 152-64 (2012). 194. Arlt, V.M. et al. Identification of three major DNA adducts formed by the carcinogenic air pollutant 3-nitrobenzanthrone in rat lung at the C8 and N2 position of guanine and at the N6 position of adenine. Int J Cancer 118, 2139-46 (2006). 195. Kool, E.T. Active site tightness and substrate fit in DNA replication. Annu Rev Biochem 71, 191-219 (2002).

136

196. Ling, H., Boudsocq, F., Woodgate, R. & Yang, W. Snapshots of replication through an abasic lesion; structural basis for base substitutions and frameshifts. Mol Cell 13, 751-62 (2004). 197. Choi, J.Y. & Guengerich, F.P. Analysis of the effect of bulk at N2-alkylguanine DNA adducts on catalytic efficiency and fidelity of the processive DNA polymerases bacteriophage T7 exonuclease- and HIV-1 reverse transcriptase. J Biol Chem 279, 19217-29 (2004). 198. Woodside, A.M. & Guengerich, F.P. Misincorporation and stalling at O(6)- methylguanine and O(6)-benzylguanine: evidence for inactive polymerase complexes. Biochemistry 41, 1039-50 (2002). 199. Schorr, S., Schneider, S., Lammens, K., Hopfner, K.P. & Carell, T. 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 (2010). 200. Batra, V.K. et al. Magnesium-induced assembly of a complete DNA polymerase catalytic complex. Structure 14, 757-66 (2006). 201. Kawanishi, M. et al. Adduct formation and repair, and translesion DNA synthesis across the adducts in human cells exposed to 3-nitrobenzanthrone. Mutat Res 753, 93-100 (2013). 202. Marnett, L.J. Oxyradicals and DNA damage. Carcinogenesis 21, 361-70 (2000). 203. Krahn, J.M., Beard, W.A., Miller, H., Grollman, A.P. & Wilson, S.H. Structure of DNA polymerase beta with the mutagenic DNA lesion 8-oxodeoxyguanine reveals structural insights into its coding potential. Structure 11, 121-7 (2003). 204. Al-Tassan, N. et al. Inherited variants of MYH associated with somatic G:C-- >T:A mutations in colorectal tumors. Nat Genet 30, 227-32 (2002). 205. Petitjean, A. et al. Impact of mutant p53 functional properties on TP53 mutation patterns and tumor phenotype: lessons from recent developments in the IARC TP53 database. Hum Mutat 28, 622-9 (2007). 206. Furge, L.L. & Guengerich, F.P. Analysis of nucleotide insertion and extension at 8-oxo-7,8-dihydroguanine by replicative T7 polymerase exo- and human immunodeficiency virus-1 reverse transcriptase using steady-state and pre-steady- state kinetics. Biochemistry 36, 6475-87 (1997). 207. Einolf, H.J., Schnetz-Boutaud, N. & Guengerich, F.P. Steady-state and pre- steady-state kinetic analysis of 8-oxo-7,8-dihydroguanosine triphosphate incorporation and extension by replicative and repair DNA polymerases. Biochemistry 37, 13300-12 (1998). 208. Einolf, H.J. & Guengerich, F.P. Fidelity of nucleotide insertion at 8-oxo-7,8- dihydroguanine by mammalian DNA polymerase delta. Steady-state and pre- steady-state kinetic analysis. J Biol Chem 276, 3764-71 (2001). 209. McCulloch, S.D., Kokoska, R.J., Garg, P., Burgers, P.M. & Kunkel, T.A. The efficiency and fidelity of 8-oxo-guanine bypass by DNA polymerases delta and eta. Nucleic Acids Res 37, 2830-40 (2009). 210. Maga, G. et al. 8-oxo-guanine bypass by human DNA polymerases in the presence of auxiliary proteins. Nature 447, 606-8 (2007). 137

211. McCulloch, S.D. & Kunkel, T.A. The fidelity of DNA synthesis by eukaryotic replicative and translesion synthesis polymerases. Cell Res 18, 148-61 (2008). 212. Sabouri, N., Viberg, J., Goyal, D.K., Johansson, E. & Chabes, A. Evidence for lesion bypass by yeast replicative DNA polymerases during DNA damage. Nucleic Acids Res 36, 5660-7 (2008). 213. Hogg, M. et al. Kinetics of mismatch formation opposite lesions by the replicative DNA polymerase from bacteriophage RB69. Biochemistry 49, 2317-25 (2010). 214. de Vega, M. & Salas, M. A highly conserved Tyrosine residue of family B DNA polymerases contributes to dictate translesion synthesis past 8-oxo-7,8-dihydro-2'- deoxyguanosine. Nucleic Acids Res 35, 5096-107 (2007). 215. Rechkoblit, O. et al. Stepwise translocation of Dpo4 polymerase during error-free bypass of an oxoG lesion. PLoS Biol 4, e11 (2006). 216. Luo, G., Wang, M., Konigsberg, W.H. & Xie, X.S. Single-molecule and ensemble fluorescence assays for a functionally important conformational change in T7 DNA polymerase. Proc Natl Acad Sci U S A 104, 12610-5 (2007). 217. Lamichhane, R., Berezhna, S.Y., Gill, J.P., Van der Schans, E. & Millar, D.P. Dynamics of site switching in DNA polymerase. J Am Chem Soc 135, 4735-42 (2013). 218. McKinney, S.A., Joo, C. & Ha, T. Analysis of single-molecule FRET trajectories using hidden Markov modeling. Biophys J 91, 1941-51 (2006). 219. Dahl, J.M. et al. Direct observation of translocation in individual DNA polymerase complexes. J Biol Chem 287, 13407-21 (2012). 220. Lieberman, K.R., Dahl, J.M., Mai, A.H., Akeson, M. & Wang, H. Dynamics of the translocation step measured in individual DNA polymerase complexes. J Am Chem Soc 134, 18816-23 (2012). 221. Lieberman, K.R. et al. Kinetic mechanism of translocation and dNTP binding in individual DNA polymerase complexes. J Am Chem Soc 135, 9149-55 (2013). 222. Abbondanzieri, E.A. et al. Dynamic binding orientations direct activity of HIV reverse transcriptase. Nature 453, 184-9 (2008). 223. Liu, S., Abbondanzieri, E.A., Rausch, J.W., Le Grice, S.F. & Zhuang, X. Slide into action: dynamic shuttling of HIV reverse transcriptase on nucleic acid substrates. Science 322, 1092-7 (2008). 224. Beard, W.A. & Wilson, S.H. Structural insights into the origins of DNA polymerase fidelity. Structure 11, 489-96 (2003). 225. Bruskov, V.I., Malakhova, L.V., Masalimov, Z.K. & Chernikov, A.V. Heat- induced formation of reactive oxygen species and 8-oxoguanine, a biomarker of damage to DNA. Nucleic Acids Res 30, 1354-63 (2002). 226. Brautigam, C.A. & Steitz, T.A. Structural and functional insights provided by crystal structures of DNA polymerases and their substrate complexes. Curr Opin Struct Biol 8, 54-63 (1998). 227. Steitz, T.A. DNA polymerases: structural diversity and common mechanisms. J Biol Chem 274, 17395-8 (1999). 228. Garcia-Diaz, M. & Bebenek, K. Multiple functions of DNA polymerases. CRC Crit Rev Plant Sci 26, 105-122 (2007). 138

229. Dodson, M.L., Michaels, M.L. & Lloyd, R.S. Unified catalytic mechanism for DNA glycosylases. J Biol Chem 269, 32709-12 (1994). 230. Yang, W. Structure and mechanism for DNA lesion recognition. Cell Res 18, 184- 97 (2008). 231. Friedman, J.I. & Stivers, J.T. Detection of damaged DNA bases by DNA glycosylase enzymes. Biochemistry 49, 4957-67 (2010). 232. Lehman, I.R. DNA ligase: structure, mechanism, and function. Science 186, 790- 7 (1974). 233. Tomkinson, A.E., Vijayakumar, S., Pascal, J.M. & Ellenberger, T. DNA ligases: structure, reaction mechanism, and function. Chem Rev 106, 687-99 (2006). 234. Nishino, T. & Morikawa, K. Structure and function of nucleases in DNA repair: shape, grip and blade of the DNA scissors. Oncogene 21, 9022-32 (2002). 235. Yang, W. Nucleases: diversity of structure, function and mechanism. Q Rev Biophys 44, 1-93 (2011). 236. Finger, L.D. et al. The wonders of flap endonucleases: structure, function, mechanism and regulation. Subcell Biochem 62, 301-26 (2012). 237. Hlinkova, V. et al. Structures of monomeric, dimeric and trimeric PCNA: PCNA- ring assembly and opening. Acta Crystallogr D Biol Crystallogr 64, 941-9 (2008). 238. Dionne, I., Brown, N.J., Woodgate, R. & Bell, S.D. On the mechanism of loading the PCNA sliding clamp by RFC. Mol Microbiol 68, 216-22 (2008). 239. Kelch, B.A., Makino, D.L., O'Donnell, M. & Kuriyan, J. How a DNA polymerase clamp loader opens a sliding clamp. Science 334, 1675-80 (2011). 240. Millar, D., Trakselis, M.A. & Benkovic, S.J. On the solution structure of the T4 sliding clamp (gp45). Biochemistry 43, 12723-7 (2004). 241. Studier, F.W. Protein production by auto-induction in high density shaking cultures. Protein Expr Purif 41, 207-34 (2005). 242. Fairhead, M. & Howarth, M. Site-specific biotinylation of purified proteins using BirA. Methods Mol Biol 1266, 171-84 (2015). 243. O'Callaghan C, A. et al. BirA enzyme: production and application in the study of membrane receptor-ligand interactions by site-specific biotinylation. Anal Biochem 266, 9-15 (1999). 244. Karshikoff, A. & Ladenstein, R. Ion pairs and the thermotolerance of proteins from hyperthermophiles: a "traffic rule" for hot roads. Trends Biochem Sci 26, 550-6 (2001).

139