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LSU Doctoral Dissertations Graduate School

2009

DNA structural selectivity of binding by the Pol I DNA polymerases DNA structural selectivity of binding by the Pol I DNA polymerases

from Escherichia coli and Thermus aquaticus from Escherichia coli and Thermus aquaticus

Andy James Budiman Wowor Louisiana State University and Agricultural and Mechanical College

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Recommended Citation Recommended Citation Wowor, Andy James Budiman, "DNA structural selectivity of binding by the Pol I DNA polymerases from Escherichia coli and Thermus aquaticus" (2009). LSU Doctoral Dissertations. 179. https://digitalcommons.lsu.edu/gradschool_dissertations/179

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DNA STRUCTURAL SELECTIVITY OF BINDING BY THE POL I DNA POLYMERASES

FROM ESCHERICHIA COLI AND THERMUS AQUATICUS

A Dissertation

Submitted to the Graduate Faculty of the

Louisiana State University and

Agricultural and Mechanical College

in partial fulfillment of the

requirements for the degree of

Doctor of Philosophy

in

The Department of Biological Sciences

by

Andy James Budiman Wowor

B. S., Louisiana State University, 2004

December 2009

ii

DEDICATION

I dedicate this dissertation to my family: Djony Wowor, Jenny Nirmalasari Sholihin,

Ellen Cynthia Wowor, Eva Kirana Wowor, and Elizabeth Clarissa Wowor. Each one of them has

taught me many life lessons, and I am eternally grateful for their love and support. Thank you for

all your prayers. God bless!!!

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ACKNOWLEDGMENTS

I would like to express my sincere gratitude to everyone who gave me the possibility to

complete this dissertation. I am deeply thankful to my advisor Dr. Vince J. LiCata whose

guidance and suggestions helped me in the research and the writing of this dissertation. I also

would like to thank Vince for giving me the opportunity to pursue my artistic endeavors.

I want to thank the past and present members of the LiCata lab. Thanks to Dr. Kausiki

Datta for teaching me the fluorescence anisotropy and isothermal titration calorimetry

techniques. I thank Carmen R. Ruiz, Dr. Chin-Chi Liu, Dr. Allison J. Richard, Gregory S.

Thompson, Hiromi S. Brown, Daniel J. Deredge, Jaycob D. Warfel, and Yanling Yang for their

help and friendship in the lab. I also extend my gratitude to the Grove lab, especially to Sreerupa

Ray for helping me to collect some of the gel shift data.

I acknowledge Dr. Anne Grove, Dr. Fareed M. Aboul-ela, Dr. Joseph F. Siebenaller, Dr.

Raymond W. Schneider, Dr. Terry M. Bricker, and Dr. W. David Constant for serving in my

dissertation committee.

I am grateful to the National Science Foundation and the Louisiana Biomedical Research

Network for the financial support of my dissertation research and to the Department of

Biological Sciences for the teaching assistantships.

Especially, I would like to give my special thanks to my family and friends whose patient

and love enabled me to complete this work. I give my special thanks to God, Wong Liong Yun

and family, Tjoa Sik Tjoen and family, J. Michael O’Connor and family, Yulia Sandra and

family, Vara Mayers and family, Cilly and family, Wirat Buarattanakarn and family, and the

priests and nuns of Santa Maria de Fatima (Toasebio) Catholic Church, Jakarta, Indonesia. I am

sincerely grateful for their endless love, support, and prayers.

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TABLE OF CONTENTS DEDICATION …………………………………………………………………...……………... ii ACKNOWLEDGMENTS ……………………………………………………..………..……… iii LIST OF TABLES ………………………………………………………………….………….. vii LIST OF FIGURES ………………………………………………………………………..…… ix LIST OF ABBREVIATIONS ………………………………………………………………… xiii ABSTRACT …………………………………………………………………………………... xiv CHAPTER 1. GENERAL INTRODUCTION ……………………………………………...….. 1 1.1 DNA Polymerases ….………………………………………………………............... 1

1.2 Structure of Klenow and Klentaq Polymerases ……………………………………... 4 1.3 Polymerization Activity …………………………………………………...……….... 9 1.4 Proofreading Activity …………………………………………………………...….. 11 1.5 Thermodynamics of DNA Binding by DNA Polymerases ……………............…… 13 1.6 DNA Sequence versus DNA Structure Selectivity ……………………………….... 17 1.7 Thermodynamic and Structural Investigations of the DNA

Structural Selectivity of Klenow and Klentaq Polymerases ..……………..……….. 19 CHAPTER 2. APPLICATIONS OF FLUORESCENCE ANISOTROPY

TO THE STUDY OF PROTEIN-DNA INTERACTIONS ………...……….….. 21 2.1 Introduction and General Background …………………………………….……….. 21 2.2 Advantages and Disadvantages of Anisotropy in Monitoring DNA Binding …………………………………………..……...……. 23 2.3 Equipment …………………………………………………………………....…….. 27 2.4 Experimental Design and Performance ……………………………...………….…. 28

2.4.1 Reagents ……………………………………………………………...…... 29 2.4.2 Excitation and Emission Parameters ………………………..…………..…31 2.4.3 Data Collection …………………………………………………….…….. 33 2.4.4 Data Analysis ………………………………………….……………….… 34 2.4.5 Other Controls ………………………………………………..………...… 37 2.4.6 Competition Experiments …………………………………………....…... 39

2.5 Other Applications of Fluorescence Anisotropy to the Study of Protein-DNA Interactions ……………………….……………….…... 39

CHAPTER 3. THERMODYNAMICS OF DIFFERENT DNA STRUCTURES

BINDING BY KLENOW AND KLENTAQ POLYMERASE …………..…..… 41 3.1 Introduction …………………………………………………………………..…….. 41 3.2 Materials and Methods ………………………………………………..……………. 43

3.2.1 Materials ………………………………………………………….…….... 43 3.2.1.1 Preparation of Oligonucleotides …………………………….…. 43

v  

3.2.1.2 Preparation of Klenow and Klentaq Polymerases ………….….. 45 3.2.2 Methods …………………………………………………...………...……. 45

3.2.2.1 Fluorescence Anisotropy ………………………...….……….… 45 3.2.2.2 Isothermal Titration Calorimetry (ITC) ..………............….…… 48 3.2.2.3 Electrophoretic Mobility Shift Assay …………...…………..…. 49

3.3 Results and Discussion ..…………………………………………………….....…... 49 3.3.1 The Binding Stoichiometry of Klenow and Klentaq Polymerases to Different DNA Structures ……………………... 49 3.3.2 DNA Structural Selectivity ……………..………….…………………….. 52 3.3.3 KCl Dependence of DNA Binding by Klenow and Klentaq Polymerases …………………….…………….... 54 3.3.4 Contributions of the Single-Stranded Region of the Template DNA to Klenow Binding .…………………….………… 57 3.3.5 Enthalpies and Heat Capacities of Binding of Different DNA Structures by Klenow and Klentaq ……..…………………….….... 60 3.3.6 The Length of DNA Effect on DNA and Polymerase Binding .………… 68 3.3.7 The Magnesium Chloride Effect on DNA and Polymerase Binding ………………………………….……………… 69 3.3.8 The RRRY Motif and ss-DNA Binding by Klenow and Klentaq …....….. 71

3.4 Summary ……………..………...…………………………………………….…….. 71 CHAPTER 4. TWO MODES OF BLUNT-END DNA BINDING BY KLENOW DNA POLYMERASE …………………………………...…..… 73

4.1 Introduction ……………………………………………………………..………….. 73 4.2 Materials and Methods ………………………………………………………..……. 75

4.2.1 Materials …………………………………………………….………….... 75 4.2.2 Methods …………………………………………………...………...……. 76

4.2.2.1 Electrophoretic Mobility Shift Assay (EMSA) .....….………..… 76 4.2.2.2 Analytical Ultracentrifugation ………………………....….…… 76 4.2.2.3 Circular Dichroism ………………….…………...…………..…. 77

4.3 Results and Discussion ……..…………………………………………….…....…... 77 4.3.1 Klenow/DNA versus Klentaq/DNA Complexes …………………….….. 77 4.3.2 The Sizes of DNA/KF and DNA/KTQ Complexes ..……….…………… 80 4.3.3 Assaying for Secondary Structure Changes Upon

Primer-Template and Blunt-End DNA Binding by Klenow and Klentaq ……………………………………………..…… 87 4.3.4 The Complex S Is 2:1 and Complex F Is 1:1 Hypothesis …………..……. 94 4.3.5 Complex S for ds-DNA/KF Is a Transient Species ….…………..………. 98 4.3.6 Potential Effects of the Kinetic Shift for KF Binding to ds-DNA on Equilibrium Titrations …………..…………..………...… 102 4.3.7 Is the ds-DNA/KF Kinetic Shift Due to an Exonuclease Activity? ………………………...……………….……….. 102 4.3.8 Is the ds-DNA/KF Kinetic Shift Due to a Shift Between the Polymerization and Editing Modes of Binding? ………………...….. 103 4.3.9 Attempts to Capture pt-DNA/KF in Complex S ……….…………..…… 105

4.4 Summary …………..…………...……………………………………...………….. 110

vi  

CHAPTER 5. DISCUSSION OF MOLECULAR MODELS FOR THE INTERACTIONS BETWEEN DNA POLYMERASE AND DIFFERENT DNA SUBSTRATES …………...…………….……….....…..… 112 5.1 Introduction ……………………………...….…………………………..……….... 112 5.2 ss-DNA Binding by Klenow and Klentaq ………………………………………... 112 5.3 Molecular Models to Explain pt-DNA versus ds-DNA Binding by Klenow and Klentaq ….…………………………………..……..…..……….... 114

5.3.1 The Oligomerization Model ……………………….………………..…... 115 5.3.2 The Polymerase-Editing Modes Model …..……….……………………. 119 5.3.3 The Unique ds-DNA/KF Binding Model ……..….………………….…. 121

5.4 Summary ………………..…………………………………..…………..……….... 123 REFERENCES …………………………………………………………………………..…… 125 APPENDIX: ELSEVIER LICENSE TERMS AND CONDITIONS ……...…………………. 141 VITA ………………………………………………………………………..……...…………. 148

vii

LIST OF TABLES

1.1. DNA constructs used for the effect of DNA sequence binding experiments …………....... 18

3.1. DNAs used for binding experiments ………………………………………….…………… 44

3.2. Stoichiometric ratios of protein:DNA binding determined using

fluorescence anisotropy and Isothermal Titration Calorimetry (ITC)a ……………………. 51

3.3. The binding constants (Kd) and free energies (∆G) of binding of

Klenow polymerase to different DNA structures at 25°C

in 10 mM Tris, 5 mM MgCl2, and 300 mM KCl at pH 7.9 ……………………………...... 52

3.4. The binding constants (Kd) and free energies (∆G) of binding of

Klentaq polymerase to different DNA structures at 25°C

in 10 mM Tris, 5 mM MgCl2, and 75 mM KCl at pH 7.9 …………….………………....... 54

3.5. The binding constants (Kd) and free energies (∆G) of binding of

Klenow polymerase to different DNA structures at 25°C

in 10 mM Tris, 5 mM MgCl2, and 200 mM KCl at pH 7.9 ………………….……………. 54

3.6. The binding constants (Kd) and free energies (∆G) of binding of

Klentaq polymerase to different DNA structures at 25°C

in 10 mM Tris, 5 mM MgCl2, and 25 mM KCl (pt- and ds-DNA)

or 5 mM KCl (ss-DNA) at pH 7.9 ………………………………………………….…....... 54

3.7. The binding constants (Kd) and the number of ions released

when Klenow polymerase binds different DNA structures (ss-13, pt-13/20,

and ds-20/20) in the 200 – 300 mM KCl concentration range ………………………….... 56

3.8. The binding constants (Kd) and the number of ions released

when Klentaq polymerase binds different DNA structures

in the 5 – 50 mM KCl concentration range for single-stranded DNA (ss-63)

binding and the 50 – 150 mM KCl concentration range

for double-stranded DNA (ds-63/63) binding …………………………….…..………....... 56

3.9. The differences in free energies between primer-template DNA

and blunt-end ds-DNA binding by Klenow polymerase assayed

via both fluorescence anisotropy and gel shift,

and using both duplex and hairpin DNA constructs ………………….……………….…... 59

3.10. Calorimetric enthalpy of binding (ΔH) and heat capacity change (ΔCp)

when Klenow polymerase binds single-stranded DNA (ss-20 and ss-63) ……………...…. 60

3.11. Calorimetric enthalpy of binding (ΔH) and heat capacity change (ΔCp)

when Klenow polymerase binds double-stranded DNA (ds-20/20 and ds-63/63)……….... 60

viii

3.12. Calorimetric enthalpy of binding (ΔH) and heat capacity change (ΔCp)

when Klentaq polymerase binds different DNA structures

(ss-20, pt-13/20, and ds-20/20) ………………………………………...…………….......... 63

3.13. Thermodynamic parameters for temperature dependence of

single-stranded DNA (ss-63) binding by Klentaq DNA polymerase …………..…………. 63

3.14. Thermodynamic parameters for temperature dependence of

double-stranded DNA (ds-63/63) binding by Klentaq DNA polymerase …………...……. 63

3.15. ΔCp values (cal / mol K) of Klenow and Klentaq binding to

different structures with different lengths of DNA,

including ΔCp data from references 96 and 97 ..………………………………...………… 67

3.16. The binding constants (Kd) and the Gibbs free energy (∆G) of

the binding of Klenow and Klentaq polymerases to primer-template DNA

(hp-39) and double-stranded DNA (hp-32 and hp-46)

in the absence and presence of MgCl2 ………………………………….…………………. 69

4.1. DNAs used for EMSA binding experiments ………………….……………....…………... 75

4.2. The measured s20,w values and the calculated molecular weights of

the globular and highly asymmetric proteins ……………………………………………… 80

4.3. The measured and predicted s20,w values and the calculated molecular weights

of the DNA/KF complexes …………………………………………………………...….... 82

4.4. The measured and predicted s20,w values and the calculated molecular weights

of the DNA/KTQ complexes ……………………………………………………………… 85

4.5. Matched and mismatched DNAs used for EMSA binding experiments

by von Hippel and associates ………………………………………………..……….……. 97

5.1. Summary of data that support and contradict the oligomerization and

the polymerase and editing modes model ……………………………….…..……….…... 124

ix

LIST OF FIGURES

1.1. X-ray crystal structures of Klenow (1KFD) and Klentaq (1KTQ) polymerases ………...…. 4

1.2. X-ray crystal structures of Klenow (1KLN) and Klentaq (4KTQ) polymerases

bound to DNA …………………………………………………………………………...….. 5

1.3. The intermediate state of the two metal ion mechanism

for polymerization activity ……………………………………………………..………….. 10

1.4. The proposed transition state of the two metal ion mechanism

for proofreading activity ……………………………………………………………..……. 12

1.5. The binding free energy of Klentaq/pt-DNA as a function of temperature …….………..... 15

1.6. Salt linkage of Klenow and Klentaq binding to pt-DNA ………………..………………… 16

1.7. Effect of DNA sequence in the single-stranded region of primer-template

DNA on the binding affinity to Klenow polymerase …...……………………..……...…… 18

2.1. Schematic illustration of the effect of rotational diffusion rate (tumbling

and spinning on axis) on the anisotropy of emitted light from

fluorescently labeled DNA ………………………………………………………….…….. 22

2.2. The temperature dependence of ROX labeled single-stranded DNA (63-mer)

binding to Klentaq DNA polymerase, illustrating the ability of to resolve

binding reactions with very similar affinities …………………………………….……….. 25

2.3. The effects of EDTA on the binding of Klentaq DNA polymerase to

primer-template DNA (13/20-mer DNA) ……………………………………………….… 26

2.4. Schematic of the sample compartment and polarizers in

a fluorometer measuring anisotropy …………………………………………….………… 27

2.5. The structure of Rhodamine-X shown attached to the α phosphate at

the 5’ end of a DNA oligomer (top panel). The bottom panel shows

the excitation and emission spectra of the fluorophore ……………………………….…... 30

2.6. Determination of binding stoichiometry of Klentaq polymerase to

double-stranded DNA (63/63-mer) …………………………………………….…………. 36

2.7. Fluorescently labeled primer-template DNA (a 13/20mer labeled with ROX)

being displaced from Klentaq DNA polymerase by identical,

but unlabeled, primer-template DNA (circles) and an unlabeled

hairpin DNA structure (squares) ………………………………………………………….. 38

x

3.1. Determination of binding stoichiometry for Klentaq polymerase

binding to double-stranded DNA (ds-20/20) …………………………….………………... 47

3.2. Determination of binding stoichiometries for Klenow and Klentaq

polymerases binding to double-stranded DNA (ds-63/63) ……………………….………. 50

3.3. DNA structure dependence of binding by Klenow and Klentaq polymerases ..………..… 53

3.4. KCl linkages (∂ln1/Kd versus ∂ln[salt]) for the binding of

Klenow (A) and Klentaq (B), and polymerases to

ss-DNA, pt-DNA, and ds-DNA …………………….…………………………….………. 55

3.5. Top panel: representative gel shift assay showing hp-32 binding

by Klenow polymerase. Bottom panel: digitized gel shift data

from the top panel, fit to a single-site isotherm (Equation 3.2) .……………………...…… 57

3.6. Temperature dependence of the enthalpy change (∆H) upon binding of

Klenow to shorter and longer DNA structures determined by calorimetry …….................. 61

3.7. Temperature dependence of the binding of Klentaq to different DNA structures ………… 62

3.8. A. Calorimetric ΔH values for Klenow-DNA binding at 30°C

in buffers with different ionization enthalpies.

B. Temperature dependences of the calorimetric ΔH of Klenow binding

to 63/70-mer DNA in the presence (■) and absence (□) of MgCl2 …………………...…… 65

3.9. Mean ΔCp values (kcal/mol K) for the binding of Klenow (A) and

Klentaq (B) to different DNA structures .…………………………………..………...…… 66

3.10. The effect of Mg2+

on the free energy of pt-DNA and

ds-DNA binding by Klenow (A) and Klentaq (B) ………………………………………… 70

4.1. Klenow (KF) and Klentaq (KTQ) binding to different DNA structures

after 10 minutes incubation time …………………………………………………………... 79

4.2. The correlation between the logarithm of the molecular weight of globular

proteins and the logarithm of their sedimentation coefficients (solid line) ……………….. 81

4.3. Continuous sedimentation (c(s)) distributions of pt-DNA/KF and

ds-DNA/KF complexes ……………………………………………………………………. 83

4.4. The measured s20,w values of the DNA/KF complexes versus

the calculated molecular weights ………………………………………………………….. 84

4.5. Continuous sedimentation (c(s)) distributions of pt-DNA/KTQ and

ds-DNA/KTQ complexes ………………………………………………………………….. 86

xi

4.6. The measured s20,w values of the DNA/KTQ complexes versus

the calculated molecular weights ………………………………………………………….. 87

4.7. Klentaq shows slightly larger secondary structure changes

upon DNA binding than Klenow ………………………………………………………….. 88

4.8. DNA spectral changes are similar upon binding of

Klenow and Klentaq to pt or ds-DNA …………………………………………………….. 89

4.9. Comparing spectra of polymerases bound to pt- vs. ds-DNA …………………………….. 90

4.10. Both polymerases show similar CD signals when binding to the different DNAs ………. 91

4.11. The signal differences between ds-DNA and pt-DNA binding

by the polymerases ………………………………………………………………………… 92

4.12. Matched and mismatched DNA binding by Klenow (KF) ………………………………. 97

4.13. Migrations of isolated Klenow (KF) on a native gel …………………………………….. 98

4.14. Klenow (KF) and Klentaq (KTQ) binding to different DNA structures

after 8 hours incubation time ………………………………………………………..……. 99

4.15. Klenow (KF) binding to ds-DNA as a function of time ……………………………....... 100

4.16. The kinetic plots of ds-DNA binding to KF ………………...……………...…………... 101

4.17. hp-39 binding by Klenow polymerase without (A) and with (B)

an additional 5 mM MgCl2 …..……………………………………………………….…... 104

4.18. The low temperature effect on Klenow (KF) binding to pt-DNA

(Lanes 1-6) and ds-DNA (Lanes 7-12) ………………………………………………….. 105

4.19. Klenow binding to pt- and ds-DNA in Mg2+

and EDTA buffers

after 10 minutes and 3 hours incubation time ………………………………………….... 106

4.20. Klenow binding to pt- and ds-DNA in EDTA, Ca2+

, no divalent metal

(no Me2+

), and Mg2+

buffers at 10 minutes incubation …………………………………. 107

4.21. Klenow (KF) (lanes 1-10) and Klentaq (KTQ) (lanes 11-20)

binding to different DNA structures in the presence and absence

of ddNTP after 10 minutes incubation time ……………………………..………………. 108

4.22. Klenow (KF) (lanes 1-4) and Klentaq (KTQ) (lanes 5-8)

binding to different DNA structures in the presence of ddNTP

after 8 hours incubation time ……………………………………………………………. 109

xii

5.1. A proposed model for single-stranded DNA binding

by Klenow (A) and Klentaq (B) polymerases ……...…………………………………….. 114

5.2. Schematic showing the oligomerization model for DNA binding

by Klenow and Klentaq polymerases ……………………………………………………. 116

5.3. The proposed polymerase and editing modes model for DNA binding

by Klenow and Klentaq polymerases ……………………………………………………. 120

5.4. The proposed unique ds-DNA/KF binding model ………………………….……………. 122

xiii

LIST OF ABBREVIATIONS

[salt] salt concentration

ΔCp Heat capacity change of binding

ΔG Free energy change of binding

ΔH Enthalpy change of binding

ΔHcal Calorimetric enthalpy change of binding

ΔHvH van’t Hoff enthalpy change of binding

Asp Aspartate

AU Analytical Ultracentrifugation

CD Circular Dichroism

D Aspartate

DNA Deoxyribonucleic Acid

EMSA Electrophoretic Mobility Shift Assay

FA Fluorescence Anisotropy

Glu Glutamate

ITC Isothermal Titration Calorimetry

Kd Equilibrium dissociation constant

Klenow “Large fragment” of E. coli Pol I

Klentaq “Large fragment” of Taq polymerase

PCR Polymerase Chain Reaction

Pol I Escherichia coli DNA polymerase I

SAXS Small Angle X-ray Scattering

Taq Thermus aquaticus DNA polymerase I

xiv

ABSTRACT

Understanding the thermodynamics of substrate selection by DNA Polymerase I is

important for characterizing the balance between replication and repair for this enzyme in vivo.

Due to their sequence and structural similarities, Klenow and Klentaq, the “large fragments” of

the Pol I DNA polymerases from Escherichia coli and Thermus aquaticus, are considered

functional homologues. Klentaq, however, does not have a functional proofreading site.

Examination of the DNA binding thermodynamics of Klenow and Klentaq to different

DNA structures: single-stranded DNA (ss-DNA), primer-template DNA (pt-DNA), and blunt-

end double-stranded DNA (ds-DNA) show that the binding selectivity pattern is similar when

examined across a wide range of salt concentration, but can differ significantly at any individual

salt concentration. For both proteins, binding of ss-DNA shifts from weakest to tightest binding

of the three structures as the salt concentration increases. Both Klenow and Klentaq release 2-3

more ions when binding to pt-DNA and ds-DNA than when binding to ss-DNA. Both of these

non-sequence specific binding proteins exhibit relatively large heat capacity changes (ΔCp) upon

DNA binding, however, Klenow exhibits significant differences in the Cp of binding to pt-DNA

versus ds-DNA, while Klentaq does not, suggesting that Klenow and Klentaq discriminate

between these two structures differently. Taken together, the G, Cp, and salt dependence

patterns suggest that the two polymerases bind ds-DNA very differently, but that both bind pt-

DNA and ss-DNA similarly, despite the absence of a proofreading site in Klentaq.

Structural data from the electrophoretic mobility shift assay (EMSA) also support a

striking difference between ds-DNA binding for Klenow and Klentaq. In EMSA, all ds-

DNA/Klenow complexes show a time dependent shift from a slower to a faster moving complex

while pt-DNA/Klenow complexes (both matched and mismatched) are found only in the fast

xv

moving complex. In contrast, all DNA/Klentaq complexes are observed in a slower moving

complex only. Several potential molecular models for correlating the thermodynamics and the

structural data are discussed. The thermodynamic differences among the different DNA

structural preferences for the two polymerases suggest that the in vivo functions of these two

largely homologous polymerases are somewhat different and respond differently to

environmental conditions.

1

CHAPTER 1

GENERAL INTRODUCTION

1.1 DNA Polymerases

One of the fundamental processes occurring in living organisms is DNA replication.

DNA polymerases are responsible for the replication of genetic information in any organism by

incorporating nucleotides complementary to a template strand of DNA. Since the isolation of the

first polymerase, DNA Polymerase I from Escherichia coli, structural and biochemical studies of

DNA polymerases have provided information about DNA replication and repair. DNA

polymerases are also used as a tool in the polymerase chain reaction (PCR) technique in

biotechnological applications such as cloning, gene analysis/fingerprinting, and detection of

diseases.

The replication and maintenance of the genome involve many different proteins and

different substrates. DNA polymerases have different roles in DNA replication and repair. In

order to understand the roles of different polymerases in DNA replication, mismatch repair,

inter-strand crosslink repair, non-homologous end joining, base excision repair, and nucleotide

excision repair, the substrate selectivity of DNA polymerases must be more clearly understood.

Enzymes are specific, and enzyme catalysis begins with the binding of the substrate by the

enzyme. Although DNA polymerases have been studied for several decades, the substrate

selection process of these enzymes is not well elucidated (1-3).

Structural studies have revealed some information about the interactions of these

polymerases with a few different DNA substrates (4-11). In order to understand the precise

functions of DNA polymerases from the same family, biochemical and thermodynamic studies

are performed because DNA polymerases from the same family often have different

2

characteristics (i.e. substrate selectivity, processivity (continuous nucleotide incorporation), and

fidelity) as a result of small structural differences (1, 2, 12-14).

In a cell, one type of DNA polymerase is mainly responsible for most of the

chromosomal DNA replication (15). The other DNA polymerases perform DNA repair and

primer removal. DNA polymerase δ catalyzes most DNA replications in Homo sapiens (16)

while DNA polymerase III carries out most of the replication in Escherichia coli (17). DNA

polymerase I, the most abundant polymerase in Escherichia coli (~400 molecules per cell),

replaces Okazaki fragments during lagging strand synthesis and repairs DNA damage (18, 19),

and the other three DNA polymerases (DNA polymerase II, IV, and V) in Escherichia coli are

involved in DNA repair (20). In addition to DNA polymerase I and III, DNA gyrase, SSB

(single-stranded DNA-binding protein), DnaA, helicase (DnaB), DnaC, DnaT, primase (DnaG),

DNA ligase, and Tus are also part of the DNA replication machinery in Escherichia coli (21).

Although many DNA polymerases have been discovered since, DNA Polymerase I from

Escherichia coli remains a central model for understanding the general mechanism of DNA

replication.

Based on amino acid sequence and crystal structure comparisons, DNA polymerases are

categorized into seven families: A, B, C, D, X, Y, and RT (22-28). Besides 5' → 3'

polymerization activity, these polymerases also possess 5' → 3' nuclease activity (family A), 3'

→ 5' exonuclease activity (families A, B, and D), lyase activity (X family), and RNaseH activity

(RT family) (28). The type I DNA polymerases from Escherichia coli (Pol I) and Thermus

aquaticus (Taq polymerase) are A family DNA polymerases. The characterizations of Pol I and

Taq polymerases are often extrapolated to emphasize “common” features of all A family DNA

polymerases and are sometimes even extended to other DNA polymerase families to identify

general properties of DNA polymerases.

3

In 1885, Theodor Escherich discovered a mesophilic bacterium, Escherichia coli, in the

human colon. Arthur Kornberg and colleagues isolated DNA polymerase I (Pol I) from

Escherichia coli in 1955 (29, 30). The Pol I of Escherichia coli is a single polypeptide with 928

amino acids and a 103 kDa molecular weight (31). E. coli’s Pol I possesses three functional

domains: the N-terminal 5' → 3' nuclease domain (residues 1 - 326), the intermediate 3' → 5'

exonuclease (proofreading) domain (residues 326 - 519), and the C-terminal 5' → 3' polymerase

domain (residues 520 - 928) (32, 33).

T. D. Brock and H. Freeze discovered a thermophilic bacterium, Thermus aquaticus, in

1969 at a hot spring in Yellowstone National Park (34). The DNA Polymerase I from Thermus

aquaticus (Taq polymerase) was first isolated by Chien et al. in 1976 (35). The cloning and over-

expression of the encoding gene for Taq polymerase in Escherichia coli produces a high yield of

Taq polymerase (33, 36). The Pol I of Thermus aquaticus is also a single polypeptide with 832

amino acids and a molecular weight of 94 kDa. T. aquaticus’s Pol I possesses three different

structural domains: the N-terminal 5' → 3' nuclease domain (residues 1 - 291), a nonfunctional

proofreading domain (residues 292 - 423), and the C-terminal 5' → 3' polymerase domain

(residues 424 - 832) (37).

The Pol I DNA polymerases from Escherichia coli and Thermus aquaticus share 38%

sequence identity based on their amino acid sequence alignment (33). The “large fragment”

domains of these proteins have 49% sequence identity. Chemical modification studies have

suggested that residues Met 512, Arg 682, Asp 705, Lys 758, Tyr 766, Arg 841, His 881, and

Asp 882 are important for polymerase activity in E. coli Pol I (38-42). Only residue Met 512 is

not conserved in Taq polymerase. Crystal structure studies have shown that Asp 705 and Asp

882 bind to the metal ions that catalyze polymerization activity (43, 44). Biochemical and

crystallographic studies have shown that residues Asp 355, Glu 357, Leu 361, Asp 424, Phe 473,

4

and Asp 501 are essential for proofreading activity in E. coli Pol I (45, 46). Only residue Asp

424 has an exact homolog in Taq polymerase (33).

The 5' → 3' nuclease domain is responsible for the removal of RNA primers from the

Okazaki fragments during lagging strand synthesis. The removal of the 5' → 3' nuclease domain

from the full length Pol I DNA polymerases from Escherichia coli and Thermus aquaticus yield

the “large fragments”: Klenow (68 kDa) and Klentaq (62 kDa) (47, 48). Klenow (from a

mesophile) denatures between 40-62°C depending on salt and pH while Klentaq (from a

thermophile) is stable up to 100°C (49, 50). X-ray crystal structures of this mesophilic-

thermophilic pair show that these polymerases have similar structures (Figures 1.1 and 1.2) (27,

32, 51).

1.2 Structure of Klenow and Klentaq Polymerases

Figure 1.1: X-ray crystal structures of Klenow (1KFD) and Klentaq (1KTQ) polymerases. Both

polymerases have “half-open right hand” topologies for their polymerase domains (51, 52). The

proofreading domain in Klentaq polymerase is inactive.

Several X-ray crystal structures of Klenow and Klentaq polymerase with and without

bound DNA substrate have been determined (43, 46, 51, 53). The polymerase domains of

Klenow and Klentaq polymerases share a common architectural feature that resembles a “half-

5

open” right hand; with “fingers,” “thumb,” and “palm” subdomains (46). The “fingers” and the

“thumb” subdomains consist of mostly α-helices while the “palm” subdomain is mainly

antiparallel β-sheet (46). The “fingers” subdomain binds the incoming dNTP while the “thumb”

subdomain binds the duplex region of the DNA. The “palm” subdomain, consisting of the

conserved active site residues, orients the primer strand and carries out phosphodiester bond

formation.

Figure 1.2: X-ray crystal structures of Klenow (1KLN) and Klentaq (4KTQ) polymerases bound

to DNA. Klenow polymerase is shown binding DNA in the editing mode (left figure) while

Klentaq polymerase is shown binding DNA in polymerization mode (right figure) (43, 53).

Klenow polymerase melts 3-4 base pairs at the primer-template DNA junction, and the

nucleotides at the 3'-end of the primer strand bind to the proofreading domain. On the other

hand, Klentaq polymerase binds the primer-template DNA junction at the “palm” subdomain of

the polymerase domain.

6

Besides copying DNA, DNA polymerases also have a 3' → 5' exonuclease domain that

functions as the proofreading or editing domain. The proofreading domain removes

misincorporated nucleotides from the 3'-end of the primer strand. Figure 1.2 shows the co-crystal

structures of a DNA/Klenow complex in editing mode and a DNA/Klentaq complex in

polymerization mode. Klenow has polymerase and editing activities, however, the only co-

crystal structure available is Klenow bound to DNA in editing mode. The duplex region of DNA

binds between the “thumb” subdomain and the proofreading domain in both binding modes. The

single-stranded template region binds at the “fingers” subdomain in the polymerization mode,

but not in the editing mode (53-56). The 3'-end of the primer strand is near the catalytic residues

of the polymerase domain in the polymerization mode of binding. In the editing mode of

binding, the polymerase melts the DNA duplex at the primer-template junction and 3-4

nucleotides at the 3'-end of the primer strand are repositioned to bind to the proofreading

domain.

The proofreading domain of Taq/Klentaq polymerase is non-functional because the

essential carboxylate residues (residues Asp 355, Glu 357, and Asp 501 in E. coli Pol I) required

for editing activity are missing in the proofreading domain (33). Sequence and structural

comparisons also show that the proofreading domain of Klentaq has many hydrophobic side

chains and does not have the residues thought responsible for single-stranded DNA binding (51).

Interestingly, however, Klentaq is still able to bind single-stranded DNA (Chapter 3), possibly

because Klentaq has an RRRY motif. The recently identified RRRY motif is located near the

base of the “fingers” subdomain. This motif is conserved across the Pol I family and has been

shown to bind the single-stranded template portion of a melted duplex DNA during proofreading

(57).

7

Since Klenow can bind DNA in both polymerization and editing modes, a number of

studies have examined the partitioning of DNA between the polymerization and editing modes

using methods such as time-resolved anisotropy (58-60) and circular dichroism (61, 62). Bailey

et al. have measured the binding energetics of Klenow/DNA interactions using steady-state and

time-resolved fluorescence experiments (58, 59). The binding affinity of Klenow to DNA was

determined using steady-state fluorescence while the equilibrium fractions of Klenow binding to

DNA in polymerization and editing modes were quantified using time-resolved fluorescence

anisotropy. This method, adapted for studying Klenow polymerase by David Millar and

associates, relies on the following assumptions: when Klenow binds dansyl-labeled DNA in the

polymerization site, the dansyl probe will have a faster fluorescence anisotropy decay because

the probe is exposed to the solvent. On the other hand, the probe will have a slower fluorescence

anisotropy decay when Klenow binds the DNA in proofreading site because the probe is buried

within Klenow. Using this approach, Bailey et al. suggested that even for matched primer-

template DNA, binding of the 3'-end of the primer strand of duplex DNA to the proofreading

domain occurs ~14% of the time, indicating that there is an equilibrium between polymerase and

proofreading binding sites (59, 63).

By using matched and mismatched primer-template DNA constructs, Millar and

associates have reported that 3-4 mismatched bases increases DNA duplex melting and shifts the

binding equilibrium predominantly to the proofreading site (58, 60). Carver et al. have quantified

that one G·G mismatch at the primer terminus caused a 3-4 fold increase in binding to the

proofreading site while two or more G·G mismatches caused at least a 250 fold increase in

binding at the proofreading site (60). When an internal single mismatch DNA was used instead

of terminal mismatch DNA, Carver et al. observed a ΔΔG of -1.1 to -1.3 kcal/mol between

matched and internal single mismatch DNA binding by Klenow (60).

8

Datta et al. have also examined the equilibrium distribution of Klenow binding to

matched and mismatched DNA using a newly developed circular dichroism (CD) approach (64).

Conformational changes upon Klenow/DNA binding were observed using DNA labeled with

dimers of 2-aminopurine (2-AP). The unstacking of 2-AP dimers causes a blue shift in its CD

signal, indicating the melting of duplex DNA and correlating with the editing mode of DNA

binding. Datta et al. have reported that Klenow binds DNA with 3 mismatches in editing mode

and suggested that DNA with 4 mismatches might not bind properly in the editing site of Klenow

(64). While differing in the details of whether 3 or 4 bases unwind and exactly what percentage

of perfectly matched primer-template DNA partitions to the editing site, these studies report an

equilibrium between polymerase and editing mode binding that shifts toward editing mode as the

number of mismatched bases increases (58-60, 63-65). Previous studies have suggested that

Klenow binds matched primer-template DNA primarily in the polymerase site (> 85%) and three

to four mismatched DNA predominantly in the editing site (> 80%) (58, 60).

Solution conditions also affect the modes of DNA binding in Klenow. Datta et al. have

suggested a role for divalent metal ions in the partitioning of Klenow binding modes (64).

Klenow binds matched primer-template DNA in the polymerase mode when Mg2+

is not present

and in the editing mode when Mg2+

is present (64). In 2 mM EDTA, Klenow is thought to bind

DNA primarily in the polymerase mode because EDTA helps scavenge divalent metal ions (64).

Because calcium ions (Ca2+

) do not catalyze either polymerization or proofreading

activities effectively, Ca2+

can be used to further examine this process. Recent studies have

suggested that Ca2+

eliminates proofreading activity and reduces polymerization activity in the

presence of various dNTPs (66, 67). When Ca2+

was used instead of Mg2+

, the rate of nucleotide

incorporation by Klenow was significantly reduced (68, 69). Klenow binds matched primer-

template DNA in the polymerase mode (57%) when Ca2+

is present (64). In both Ca2+

and Mg2+

,

9

Klentaq only binds DNA in polymerase mode because Klentaq lacks 3' → 5' exonuclease

activity (70) and presumably a proofreading binding site (51, 71).

1.3 Polymerization Activity

DNA polymerases add dNTPs to the 3'-OH of the primer strand of DNA in the 5' → 3'

direction during the polymerization reaction. The polymerization reaction is highly

unidirectional in vivo because the unstable pyrophosphate (PPi) effectively hydrolyzes into

inorganic phosphate (Pi) after PPi is released upon nucleotide incorporation (72). The first step

in the polymerization reaction is the binding of DNA polymerase to the primer-template DNA,

forming a binary complex. Next, the binding of dNTP to the binary complex creates an “open”

ternary complex. The complex changes its conformation to a “closed” ternary complex, and then

the nucleotide is added to the 3'-OH of the growing primer strand (the chemical step). After a

second conformational change, the pyrophosphate product is released, and the DNA polymerase

either performs another round of polymerization reaction or releases the DNA substrate (73).

The chemical step of incorporating nucleotides occurs through a two metal ion

mechanism (74). The mechanism for catalysis is illustrated in Figure 1.3. Crystallographic

studies have shown that residues Asp 705 and Asp 882 bind to Mg2+

and Mn2+

metal ions at the

active site of the polymerase domain in Klenow (43, 44). The active site serves to 1) deprotonate

the 3'-OH of the primer, 2) stabilize the pentavalent transition state, and 3) facilitate the removal

of pyrophosphate (25, 43, 44). Although crystallographic studies have shown that both Mg2+

and

Mn2+

filled the metal ion binding sites of the polymerase domain of Klenow, the exact metal ions

used in vivo have not been identified (75). The interaction between the duplex part of DNA and

the “thumb” subdomain of the polymerase may be important for processivity because deleting

the tip of the “thumb” subdomain causes a 4 fold reduction in processivity (76). Both

polymerases bind 5-8 bp of the duplex part of DNA in polymerization mode based on the photo-

10

crosslinking, chemical footprinting, and fluorescence studies of Klenow/DNA and the crystal

structure of Klentaq/DNA (77).

Figure 1.3: The intermediate state of the two metal ion mechanism for polymerization activity.

This figure is adapted from Figure 3 from reference 27 and was created using the program

BKchem. The two divalent metal ions (Me2+

) stabilize the pentavalent transition state. The two

metal ions are in contact with the two conserved aspartate (D705 and D882) residues, the

phosphates of the dNTP, the main chain oxygen (carbonyl), and two water molecules (black

circles). Metal ion A induces the attack of the 3'-OH of the primer on the α-phosphate of the

dNTP while metal ion B chelates the β- and γ-phosphates of the dNTP and stabilizes the negative

charge of the oxygen.

11

1.4 Proofreading Activity

Klenow and Klentaq primarily differ in their 3' → 5' proofreading domain (33, 51). 3' →

5' exonuclease catalysis functions in the opposite direction of DNA synthesis (19, 25). In

addition to its 5' → 3' polymerase activity, Klenow polymerase also has a 3' → 5' exonuclease

(proofreading) activity (28). The 3'-end of the primer strand of DNA can translocate between the

polymerization and proofreading domains without the dissociation of the DNA (43, 46, 78). The

polymerase and the proofreading active sites are 30 – 35 Å apart in Klenow polymerase. The

presence of an active 3' → 5' exonuclease domain increases replication fidelity (19). A

mismatched nucleotide at the 3'-end of the primer promotes its partitioning to the proofreading

site, where the mismatched nucleotide will be removed through hydrolysis of the phosphodiester

bond.

The proofreading activity also follows a two metal ion mechanism (Figure 1.4) (75, 79).

The divalent metal ions facilitate 1) the stabilization of the transition state, 2) the deprotonation

of a water molecule, 3) the nucleophilic attack to cleave the phosphodiester bond, and 4) the

dissociation of the 3'-OH (75, 78, 79). One metal ion interacts with Asp 355, Glu 357, Asp 501,

and the 5'-phosphate of the dNMP while the other metal ion coordinates with Asp 355 and the 5'-

phosphate of the dNMP (75).

The wild-type Klenow polymerase is able to degrade DNA substrate via its exonuclease

activity, even under “ideal” polymerization conditions in vitro. Because this can significantly

interfere with studies of the polymerization reaction, Joyce and associates thus constructed a

Klenow derivative called Klenow exo minus (exo-) and which contains a D424A mutation (45,

80). Klenow exo- lacks 3' → 5' exonuclease activity, but it can still bind DNA substrate in the

proofreading site (45, 81). Crystallographic studies have shown that single-stranded DNA is the

substrate for the exonuclease domain (43, 75, 78, 82, 83).

12

Figure 1.4: The proposed transition state of the two metal ion mechanism for proofreading

activity. This figure is based on Figure 10 from reference 75 and was created using the program

BKchem. Beese and Steitz solved the co-crystal structure of Klenow using the crystals of the

D424A mutant of Klenow polymerase (75). The attack of a hydroxide ion on the phosphorus is

facilitated by the interactions with tyrosine (Y497), glutamate (E357), and the metal ion A. Metal

ion B stabilizes the O-P-O bond and facilitates the leaving of the 3'-hydroxyl group. Metal ions

A and B (Me2+

) interact with the aspartate residues (D355 and D501).

13

Proofreading activity increases the level of accuracy in DNA replication. Full-length

DNA Polymerase I from Escherichia coli synthesizes DNA with an error rate of 1.6 x 10-7

– 1.5

x 10-6

/ bp (84). The average error rate for Klenow exo- is 2.5 x 10-5

– 1 x 10-4

/ bp (85, 86),

which is 7 – 30 times higher than the error rate of wild-type Klenow (85, 86). Full-length DNA

Polymerase I from Thermus aquaticus’ error rate ranges from 8.9 x 10-5

to 1.1 x 10-4

/ bp (47, 70,

87-90). Klentaq polymerase has an error rate of 5.1 x 10-5

/ bp (47). The method of assay used

and the specific substrates investigated may cause the substantial error rate variations observed

(91).

1.5 Thermodynamics of DNA Binding by DNA Polymerases

Thermodynamic measurements help determine the strength, the molecular driving forces,

and the role of the solvent and temperature in the control of protein/DNA interactions. The

physical environment (i.e. salt concentration and type, temperature, and pH) strongly affects the

non-covalent driving forces of these interactions (92). Fluorescence anisotropy, isothermal

titration calorimetry, the electrophoretic mobility shift assay, and filter binding are the techniques

that have often been used to obtain the binding energetic of protein/DNA interactions accurately

(93-96).

Klenow and Klentaq polymerases have similar structures, and they are traditionally

thought to have similar functions in vivo. They are also generally assumed to behave almost

interchangeably at the molecular level, such that experimental or structural results obtained for

one of them is assumed true for both. However, Klenow and Klentaq polymerases exhibit both

similarities and differences in their DNA binding thermodynamics. Klenow and Klentaq both

show increasing DNA binding affinity with increasing temperature up until ~40°C when they

bind to primer-template DNA (97, 98). After about 40°C, Klenow denatures and Klentaq binding

affinity decreases (Figure 1.5). The two polymerases bind DNA with submicromolar affinities in

14

very different salt concentration ranges (Figure 1.6). At similar [KCl], Klenow binding is ~3

kcal/mol (150x) tighter than Klentaq binding to primer-template DNA (99). The DNA binding of

both Klenow and Klentaq polymerases are enthalpy driven at their physiological temperatures.

Klenow and Klentaq undergo relatively small conformational changes when binding to DNA

(97, 98).

The heat capacity change (ΔCp) is a thermodynamic parameter that reflects structural

properties of the interaction between DNA and protein. ΔCp is most often attributed to changes

in hydrophobic interactions, although Sturtevant has shown that at least six different types of

non-covalent molecular processes will contribute to ΔCp (100). ΔCp is the temperature

dependence of the enthalpy of a reaction. If a reaction’s ΔH does not change with temperature,

the Cp for that reaction is zero. Cp for polymerase/DNA binding can be determined from

measuring ΔH as a function of temperature (the slope is Cp) using isothermal titration

calorimetry or by measuring ΔG as a function of temperature (the curvature is Cp, see Figure

1.5) using fluorescence anisotropy (97, 98). At the molecular level, this Cp has often been

correlated to the changes in the accessible surface area ( ASA), and primarily hydrophobic

surface area, upon complex formation, with the assumption that Cp is temperature independent.

This assumption is based on small protein folding results and the correlation of the number of

water molecules around apolar molecules with Cp (101-104). Datta et al. have shown that

surface area burial can only account for about half of the relatively large negative heat capacity

change for Klenow/primer-template DNA binding (98). In addition to ASA, protein-DNA

conformational changes (105, 106), restriction of vibrational modes upon complex formation

(100, 107, 108), linked proton uptake upon complex formation (109, 110), multiple cooperative

weak interactions (111), and coupled folding/unfolding (103, 112-117) have all been shown to be

contributors in the measured heat capacity change of protein/DNA interactions.

15

Figure 1.5: The binding free energy of Klentaq/pt-DNA as a function of temperature. Klentaq

show increasing binding affinity towards primer-template DNA with increasing temperature up

until about 40°C and decreasing binding affinity after 40°C (97). If Cp were zero, this plot

would be linear. Such curvature of ΔG versus temperature plots is common in site-specific

protein-DNA interactions.

16

Figure 1.6: Salt linkage of Klenow and Klentaq binding to pt-DNA. The two polymerases bind

DNA with submicromolar affinities in different salt concentration ranges (99). At similar [KCl],

Klenow binding is ~3 kcal/mol (150x) tighter than Klentaq binding to primer-template DNA

(99).

High ΔCp’s of binding have been shown to frequently be associated with sequence

specific DNA binding (118). However, Klenow and Klentaq are non-sequence specific binding

proteins. Non-sequence specific DNA binding was originally postulated to have a zero Cp

(118). However, Klenow and Klentaq exhibit relatively large negative Cp values when binding

to DNA (97, 98). A small number of non-sequence specific DNA binding proteins have also

exhibited unexpectedly large negative Cp values (109, 119).

Some studies have also revealed temperature dependent heat capacity (Δ Cp) effects and

proposed that Δ Cp is caused by linked structural changes with temperature and anion dependent

of binding (120-122). Kozlov and Lohman examined the effect of monovalent salt

concentrations and types over a wide temperature range (5-60°C) and found a significant

17

temperature dependent heat capacity for the binding of E. coli SSB (single-stranded DNA

binding) protein to DNA (122). A recent study showed that over half of 48 protein-DNA binding

pairs from the scientific literature can be fitted with temperature dependent heat capacity effects

and 90% of these pairs have negative Δ Cp values (123). Solution structural data for Klentaq

polymerase suggests that a coupled folding event does not cause the observed heat capacity

effects (123). The inclusion of the temperature dependent heat capacity (Δ Cp) does not

significantly improve the fits to my thermodynamic data. Assessing if Δ Cp effects exist for

protein-DNA interactions is important because all current models proposing that ΔCp and ASA

are correlated require that Δ Cp = 0. Thus, finding that Δ Cp ≠ 0 suggests that ΔCp may not

always correlate with ASA (123).

1.6 DNA Sequence versus DNA Structure Selectivity

Because, as noted earlier, large ΔCp values have been frequently correlated with sequence

specific DNA binding, we decided to examine if our fluorescence anisotropy assay might detect

some amount of previously unnoticed sequence preferences of primer-template DNA/polymerase

binding. Joyce and colleagues have reported that at least the first four nucleotides of the single-

stranded part of primer-template DNA are important for Klenow binding (56). We have

measured the effect of DNA sequence in this single-stranded region of primer-template DNA

using homogeneous seven nucleotide 5'-overhangs. We used poly-T, poly-C, and poly-A

overhangs (Table 1.1).

Poly-G was not examined in this series due to its tendency to form secondary structures

(124). Fluorescence anisotropy assays of Klenow binding to these different DNA sequences were

performed at 25°C. The binding affinities of Klenow to 13/20-mer with TCCCAAA overhang

(mixed sequence), with a poly-T overhang, and with a poly-C overhang are within error (ΔG = -

11.1 ± 0.2 kcal/mole, -11.2 ± 0.2 kcal/mole, and -11.1 ± 0.2 kcal/mole, respectively), while the

18

binding affinity of Klenow to 13/20-mer with poly-A overhang is slightly weaker (ΔG = -10.6 ±

0.2 kcal/mole) (98). Base-stack in single-stranded poly-A has been suggested to alter its ΔCp of

interaction with protein (98). These data strongly indicate that there is little or no effect of the

DNA sequence of the single-stranded portion of the primer-template DNA on the binding

affinity of Klenow to DNA (Figure 1.7). If Pol I polymerases are not sequence specific, are they

structure specific binders? How specific is the primer-template DNA interface and is its

specificity or selectivity the same for both polymerases? These are some of the major questions

addressed in this dissertation.

Table 1.1: DNA constructs used for the effect of DNA sequence binding experiments.

DNA Sequence

13/20-mer

(mixed)

5’-TCGCAGCCGTCCA-3’

3’-AGCGTCGGCAGGTTCCCAAA-5’

13/20-mer

(poly-T)

5’-TCGCAGCCGTCCA-3’

3’-AGCGTCGGCAGGTTTTTTTT-5’

13/20-mer

(poly-C)

5’-TCGCAGCCGTCCA-3’

3’-AGCGTCGGCAGGTCCCCCCC-5’

13/20-mer

(poly-A)

5’-TCGCAGCCGTCCA-3’

3’-AGCGTCGGCAGGTAAAAAAA-5’

Figure 1.7: Effect of DNA sequence in the single-stranded region of primer-template DNA on

the binding affinity to Klenow polymerase. There is no or little effect of DNA sequence on

Klenow-DNA binding (98). Error bar is based on multiple measurements.

19

1.7 Thermodynamic and Structural Investigations of the DNA Structural Selectivity of

Klenow and Klentaq Polymerases

In this dissertation, I have characterized the binding of Klenow and Klentaq to different

DNA structures: single-stranded, primer-template, and blunt-end double-stranded DNA to

further understand the DNA structural selectivity of these polymerases. Fluorescence anisotropy

technique and method of analysis are outline in Chapter 2. From a thermodynamic point of view,

this study shows that Klenow binds primer-template and blunt-end DNA differently while

Klentaq binds these DNAs similarly (Chapter 3). The binding of Klenow and Klentaq to

different DNA structures show that the binding selectivity patterns are similar when examined

across a wide range of salt concentration, but can significantly differ at any individual salt

concentration (Chapter 3). Single-stranded DNA binding for both polymerases shifts from

weakest to tightest binding of the three DNA structures as salt concentration increases (Chapter

3). These thermodynamic studies are the main focus of Chapter 3 of this dissertation. These

thermodynamics suggest that the corresponding binding complexes will differ at the structural

level although the magnitude of such structural changes is unclear. This is the main focus of

Chapter 4 of this dissertation.

Although Klenow can bind DNA in both polymerase and editing modes while Klentaq

can only bind DNA in polymerase mode, this can only partially explain their differing

thermodynamics and differing DNA structural selectivity. A recent review suggests that DNA

polymerases can form both 1:1 and 2:1 protein:DNA complexes (3). Klenow polymerase, T4

polymerase, and mammalian polymerase β (Pol β) have been suggested to form 2:1 complexes

(125-132). The different stoichiometric forms of these complexes may have some potential

functional significance in vivo and in vitro (125-132).

Only 1:1 Klenow/DNA complexes have been observed using fluorescence anisotropy and

isothermal titration calorimetry (Chapter 3; references 64 and 99) while 1:1 Taq/DNA complexes

20

have been seen using small-angle neutron scattering (130). On the other hand, the recently

postulated 2:1 Klenow/DNA complexes were characterized using gel shift and analytical

ultracentrifugation (131).

In Chapter 4 of this dissertation, I show that two types of Klenow/DNA complexes

(slower and faster complexes) and one type of Klentaq/DNA complex (slower complex) are

observed using the electrophoretic mobility shift assay (EMSA). Analytical ultracentrifugation

(AU) and circular dichroism (CD) were used to attempt to discover the potential identities of

these complexes and to examine if they reflected either different conformations or different

stoichiometries of polymerase:DNA complexes. Several previously unobserved binding

behaviors for Klenow polymerase with blunt-end DNA were characterized in this study. Neither

primer-template DNA binding nor blunt-end DNA binding by Klenow or Klentaq clearly fits

into a monomer-dimer equilibrium framework. Several potential molecular models for

correlating the thermodynamics and the structural data are discussed.

21

CHAPTER 2

APPLICATIONS OF FLUORESCENCE ANISOTROPY

TO THE STUDY OF PROTEIN-DNA INTERACTIONSA

2.1 Introduction and General Background

Since its introduction in 1990 (133), the use of fluorescence anisotropy as a method for

monitoring protein-DNA interactions has been steadily on the rise. As a real-time, solution based

assay, it has several advantages over its closest “competitors”: filter binding (134) and the

electrophoretic mobility shift assay (135, 136). The methodology has been reviewed several

times, both in the context of general overviews of fluorescence-based methods (96, 137, 138),

and as the sole focus of particular reviews (95, 139-141). The method is also the subject of three

US patents (142-144). This chapter will review the use of fluorescence anisotropy to obtain the

data for this dissertation.

For fluorescence anisotropy, the two key fundamental properties to keep in mind are: 1)

that fluorescent molecules have both an excitation and an emission dipole, and 2) that there is a

short time delay between absorbance of the exciting photon and release of the fluorescent photon

(the fluorescent lifetime).

The excitation dipole of the molecule dictates that, for any solution of fluorophores,

polarized light will only excite those molecules in the solution that are in the proper orientation,

i.e. those fluorophores that just happen to be oriented so that their excitation dipole aligns with

the polarized incident light. Since illumination is constant in fluorescence anisotropy,

fluorophores in the solution will continuously be tumbling into and out of alignment with the

_____________________________

AParts of this chapter have appeared in Methods Cell Biology, 84, V. J. LiCata and A. J.

Wowor, Applications of fluorescence anisotropy to the study of protein-DNA interactions, 243-

262, Copyright 2008, and are used with permission of Elsevier.

22

incident polarized light. Similar to the excitation dipole, the emission dipole of a fluorophore

determines the polarity of the light released by that fluorophore. The exact time delay between

excitation and emission follows an exponential decay law, and the average time delay is denoted

the fluorescence lifetime for that fluorophore (145). Fluorescent lifetimes for common

biochemical fluorophores are typically in the 1-25 nanosecond time range.

If an excited fluorophore molecule tumbles (rotationally diffuses) within its fluorescent

lifetime, then the polarization of its emitted light will be determined by its new position. If a

whole population of fluorophores tumbles within the fluorescent lifetime, the emitted light will

become completely depolarized, because the positions of all the emission dipoles will have

effectively been randomized.

Figure 2.1: Schematic illustration of the effect of rotational diffusion rate (tumbling and spinning

on axis) on the anisotropy of emitted light from fluorescently labeled DNA. Both the free DNA

and the complex-bound DNA are illuminated by polarized light. Since the DNA in the complex

tumbles more slowly, a larger proportion of its emitted light remains polarized or anisotropic.

This figure is reprinted from reference 167 with permission.

23

The central basis for using fluorescence anisotropy to monitor molecular interactions is

based on the fact that larger molecules tumble more slowly (and thus retain more emission

polarization), while smaller molecules tumble more quickly (and thus depolarize the emission

more effectively). A small stretch of fluorescently labeled DNA will tumble (and spin on its axis)

faster when alone in solution than when bound to a protein. Figure 2.1 summarizes this effect in

cartoon form. Thus, the increase in anisotropy due to slower rotational diffusion of the protein-

DNA complex relative to the free DNA is the dependent signal that translates directly into the

fraction of DNA bound in this technique.

2.2 Advantages and Disadvantages of Anisotropy in Monitoring DNA Binding

The advantages and disadvantages of monitoring protein-DNA interactions by

fluorescence anisotropy have been extensively discussed in different reviews and several original

research papers (95, 141). The main advantage of the technique is the fact that it is a solution

based equilibrium technique. This allows one to make measurements without fear that the

detection technique is altering the reaction equilibrium. Furthermore, it allows one to alter

solution conditions for measurements quite easily without fear of altering the direct relationship

between the signal provided by the detection method and the progress of the reaction. Separation

methods, such as filter binding (134) or electrophoretic mobility shift assays (135, 136), may

perturb the reaction equilibrium. The separation process itself pulls reactants (DNA and protein)

away from products (complex). This creates concentration gradients for each component, and so

each sub-environment (i.e. region of specific concentrations of all reactants and products) during

the separation will be thermodynamically pushed toward its own equilibrium, unless such

rearrangement can be fully quenched during the separation process. The result is that the

fractions of each component seen separated on the final gel, or retained on the filter, may not be

24

the same as the fractions of each component in the original equilibrium mixture. In fact, some

researchers prefer to refer to these assays as “non-equilibrium” methods in general.

A further problem with separation-based assays arises if one changes solution conditions

in the sample mixture (salt, temperature, pH, osmolytes, etc.). One must then perform controls to

insure that such changes have not perturbed the separation method itself. For example, varying

the amount of salt, or adding an osmolyte to a protein-DNA reaction can directly alter the

efficiency with which the protein-DNA complex sticks to a nitrocellulose filter or enters a gel. If

one changes the reaction conditions in fluorescence anisotropy, one may alter the value for the

absolute anisotropy, but one will not alter the fact that the normalized change in anisotropy (ΔA)

will still scale directly with fractional saturation ( ).

Other advantages of fluorescence anisotropy include: 1) The fact that it is a real time

assay. One does not wait for a gel to run or radioactivity on filters to be counted to obtain the

result. 2) The data produced are almost always of much higher precision (much lower random

data scatter) than those typically obtained via filter binding or electrophoretic mobility shift

assays. This allows discrimination among reactions of very similar affinity. Figures 2.2 and 2.3

show the clean, clear resolution among four binding reactions that differ from one another by

less than 0.5 kcal/mole. The technique can easily and reproducibly resolve between binding

reactions that differ from one another by < 10% (e.g. a binding curve with a 9 nM Kd is distinct

from one with a 10 nM Kd) (98). Much current biophysical research on protein-DNA interactions

is focused on how solvent components (ions, protons, osmolytes, water activity, etc.) act to

regulate these interactions. Due to its precision and its reliability across a wide range of solution

conditions, fluorescence anisotropy is an extremely well suited method for studying such aspects

of protein-DNA interactions. 3) The titration process can readily be automated.

25

Figure 2.2: The temperature dependence of ROX labeled single-stranded DNA (63-mer) binding

to Klentaq DNA polymerase, illustrating the ability of to resolve binding reactions with very

similar affinities. Equilibrium titrations are shown at 25°C (●), 35°C (■), 45°C (♦), and 55°C

(▲). All titrations were performed in 10 mM Tris, 5 mM MgCl2, 5 mM KCl, at pH 7.9.

Increasing temperature decreases the binding affinity of Klentaq polymerase to single-stranded

DNA. At 25°C the Kd is 33.6 nM (ΔG = -10.2 kcal/mole). At 35°C the Kd is 47.6 nM (ΔG = -

10.3 kcal/mole). At 45°C the Kd is 77.6 nM (ΔG = -10.3 kcal/mole). At 55°C the Kd is 97.0 nM

(ΔG = -10.5 kcal/mole). This figure is reprinted from reference 167 with permission.

26

0

0.2

0.4

0.6

0.8

1

0 50 100 150 200 250 300

0 mM EDTA

5 mM EDTA

10 mM EDTA

20 mM EDTA

No

rma

lize

d A

nis

otr

op

y

[Klentaq] (nM)

Figure 2.3: The effects of EDTA on the binding of Klentaq DNA polymerase to primer-template

DNA (13/20-mer DNA). As with Figure 2.2, the data illustrate the high precision possible when

monitoring binding with fluorescence anisotropy. Equilibrium titrations are shown in the absence

of EDTA (●) and in the presence of EDTA 5 mM EDTA (■), 10 mM EDTA (♦), and 20 mM

EDTA (▲). EDTA, a metal chelator, decreases the affinity of Klentaq polymerase to DNA. In

the absence of EDTA, the Kd is 7.5 nM (ΔG = -11.1 kcal/mole). In 5 mM EDTA, the Kd is 18.0

nM (ΔG = -10.6 kcal/mole). In 10 mM EDTA, the Kd is 41.3.0 nM (ΔG = -10.1 kcal/mole). In 20

mM EDTA, the Kd is 89.9 nM (ΔG = -9.6 kcal/mole). All titrations were performed at 25°C in

10 mM Tris, 50 mM KCl, at pH 7.9. This figure is reprinted from reference 167 with permission.

27

One of the main disadvantages of the technique stems from the instrumental constraints.

Fluorescence anisotropy is infrequently used to measure binding constants that are tighter than 1

nM, simply because the anisotropy signal from < 1 nM concentration of most fluorophores dips

below the detection limit for most commercial fluorometers. This problem is discussed further

below in “Equipment”.

2.3 Equipment

Figure 2.4: Schematic of the sample compartment and polarizers in a fluorometer measuring

anisotropy. The excitation polarizer remains at vertical at all times, while the emission polarizer

switches between vertical and horizontal in order to measure I|| and I┴, respectively. Anisotropy

is calculated (automatically in most instruments) using Equation 2.1 in the text. This figure is

reprinted from reference 167 with permission.

The experimental setup for measurement of fluorescence anisotropy involves significant

loss of light intensity at numerous points along the optical path. Incident light first passes

through a monochromator to select the incident wavelength(s) (even at the selected wavelength,

28

30% losses in intensity are not uncommon in a standard monochromator). The incident light then

passes through the vertical polarizer. Figure 2.4 shows a potential arrangement of polarizers in a

fluorometer measuring anisotropy. Just as with polarized sunglasses, the loss in intensity through

a polarizer is tremendous as all incident light except that in the vertical plane is filtered out by

the polarizer. As the light passes through the sample, now only those fluorophores with properly

aligned absorbance dipoles will absorb the vertically polarized light. This is a very small fraction

of the total fluorophore population at any “steady state” instant. Only those fluorophores that

absorb can fluoresce, so the total outgoing fluorescence intensity can be orders of magnitude

lower than if all the fluorophores in the solution had absorbed. Finally, as the light leaves the

sample it must pass through another polarizer and usually another monochromator. This loss of

intensity at so many steps can mean that, in some cases, a fluorophore that might have a steady

state emission intensity of several million photons per second in a particular machine can display

a steady state anisotropic emission intensity in either the vertical or horizontal detection mode (I||

and I┴ in Equation 2.1, respectively) of only a few thousand photons per second. Thus common

biological fluorophores such as fluorescein or rhodamine based dyes, which might give strong

standard fluorescence signals at 1 nM concentration, may give no discernable signal at all once

the polarizers are placed in the light path. For this reason, photon-counting fluorometers are the

instrument of choice for fluorescence anisotropy relative to analog fluorometers, due to their

enhanced sensitivity to low intensity emission.

Quantitatively, anisotropy, denoted A, is measured as:

(Eq. 2.1)

2.4 Experimental Design and Performance

Since the DNA is much more easily fluorescently labeled, the usual titration mode for a

fluorescence anisotropy experiment is to titrate protein into a solution of labeled DNA. Also,

29

generally, the DNA fragment is smaller than the protein, and thus one will see a larger relative

change in rotational diffusion (and hence anisotropy) for the DNA when it forms complex. The

DNA concentration in many published studies is near 1 nM (often the lowest usable

concentration, as discussed above), and titrations are performed by adding protein to a partially

filled cuvette containing the DNA solution. After each incremental addition of protein, the

solution is allowed to stir for several minutes, then the fluorescence anisotropy is measured. A

good plot of anisotropy (A) versus protein concentration has a clear plateau at high protein

concentration, and contains several points below the Kd value for the reaction. The following

sections contain some information on the individual steps in the experimental procedure.

2.4.1 Reagents

Commercially obtained DNA oligomers should be HPLC or PAGE purified. DNA is

generally labeled at a point farthest from the anticipated protein binding site, to avoid protein

interactions with the fluorophore. Most commercial DNA oligomer synthesis companies will

attach a fluorophore onto either end of an oligomer, and many fluorophores can be attached in

the middle of an oligomer. Some researchers have used in-house attachments of different

fluorophores to DNA when commercial preparation is undesirable or unavailable (95, 146).

A number of different fluorophores have been successfully used. Fluorescein based dyes

remain the popular favorite. Rhodamine-X (ROX), introduced for use in this application by

Beechem and associates (147), has the advantage of displaying a lower tendency to interact

directly with the protein. Figure 2.5 shows the structure of rhodamine-X and its excitation and

emission spectra. The complication of protein-fluorophore interactions has been observed for

several systems with fluorescein labeled DNA (see “Controls” below). Rusinova et al. (2002)

describe the use of newer Alexa and Oregon Green fluorophores as fluorescent labels in

fluorescent anisotropy (148).

30

Figure 2.5: The structure of Rhodamine-X shown attached to the α phosphate at the 5'-end of a

DNA oligomer (top panel). The bottom panel shows the excitation and emission spectra of the

fluorophore. This figure is reprinted from reference 167 with permission.

31

The DNA should not be too large relative to the protein. Otherwise, binding of the

protein will not alter its rotational diffusion enough to produce an observable change in the

anisotropy signal. Several reviews quote 40 bp as an upper limit on the size of the DNA, but our

lab has used oligomers in the 70 bp range without problems, and Heyduk et al. (1996) predict

that oligomers up to approximately 105

Da (about 140bp) should work based on estimated

rotational diffusion rates (95, 148).

Since most fluorescence anisotropy titrations used to examine protein-DNA interactions

involve titrating protein into DNA, it is best if the DNA concentration is far below the Kd value

(however, sometimes the detection limit of the fluorometer precludes this). If the DNA is kept

far below the Kd, then even significant errors in the DNA concentration will not propagate into

the final data.

Any error in the protein concentration, however, will be directly reflected in the Kd, since

the protein concentration comprises the x-axis of the titration. For this reason we typically

determine our protein concentrations by two different methods (in our laboratory we use

absorbance at 280 nm and the Bradford assay) (149).

Many enzyme solutions, upon long term storage, especially frozen, will accumulate

insoluble particulates, often invisible to the naked eye, but which can significantly interfere with

both standard fluorescence measurements and fluorescence anisotropy measurements. A 5

minute, full speed spin in a standard microfuge will usually clear such particulates. The enzyme

concentration must be re-determined after such treatment.

2.4.2 Excitation and Emission Parameters

Because so much of both the incident and emitted light is lost in an anisotropy

measurement, the setup of the excitation and emission parameters requires some caution and

iterative empirical testing. If monochromators are used, the band pass should be opened as

32

widely as possible, to capture as much of the excitation and emission peaks as possible, without

risking overlap of the two. Since most steady state fluorescence signals are so strong, the use of

band pass widths < 1nm are typical in normal fluorescence measurements, but for anisotropy this

is not the case. For example, with rhodamine-X, the excitation peak is at 583 and the emission

peak is at 605 nm (see Figure 2.5), and we use 8 nm band pass width around each peak

maximum. Similar considerations should be used if choosing band pass filters: letting through as

much light as possible without cross contaminating the excitation and emission signals.

Again, because the signal is so low, integration times should be maximized. Instead of

the typical 0.1-1 second integration times used with steady state fluorescence, we use 10 second

integration times with a minimum of 5 averaged measurements to obtain maximal precision

under low signal conditions.

An odd, but experimentally necessary element in fluorescence anisotropy measurements

in protein-DNA interactions is the need for an exceedingly long “wait time” after each addition

of protein for the anisotropy signal to stabilize. For protein-DNA interactions with nanomolar

Kd’s the actual time till equilibrium will be far less than a second. It is typical, however, to wait

4-10 minutes after each addition of protein before the next anisotropy measurement is taken to

achieve maximal precision and stability of the measurement (in our laboratory, 8 minutes is

used). This might be either a mixing effect or a temperature effect. Even in a well-stirred cuvette

there is only a slow approach to absolute homogeneity of mixing. Evidence for such an effect

can be seen if one adds titrant to the top of the solution in the cuvette versus inserting the pipette

farther into the cuvette and adding titrant near the bottom. Additionally, since precise

temperature control is so tightly linked to signal precision in anisotropy, adding even small

amounts of titrant to the cuvette may necessitate a slow return to the set temperature. Similar

“solution settling” effects are seen in dynamic light scattering measurements, and since any stray

33

light scattering will interfere with anisotropy measurements, the exact mechanism for this effect

in fluorescence anisotropy may be similar. One can empirically determine the best “wait” time

for one’s own system, but virtually all published studies simply use a consistent wait time of at

least 4 minutes (95, 150).

2.4.3 Data Collection

For any ligand binding titration, one wants to achieve maximal consistency of spacing of

data along the y-axis. It is naturally much easier to achieve equal spacing along the independent

axis, since one knows how much protein one is adding at each step. However, uniform spacing

along the dependent axis is significantly more crucial for successful data analysis. A typical total

anisotropy change (ΔAT) for a protein-DNA interaction might be in the range of 0.1 to 0.15. One

typically wants 15-20 points spanning that range for a single-site binding isotherm. More points

may be necessary for more complex multi-site binding situations, where subtle curve shape

changes need to be accurately quantitated. Because y-axis spacing is more important to data

analysis, the amount of protein added to the cuvette will typically increase as the titration

continues.

On the x-axis, it is important that some points be below the Kd value, otherwise one is

determining a Kd value using data that does not even overlap the Kd. Although such calculation

is possible, it significantly reduces the reliability of the Kd determination. Because one generally

does not know the actual Kd value when starting a titration, this frequently means that “one”

titration will actually involve collecting an iterative set of titrations until a data set is obtained

that includes about 3-5 data points below the Kd value, and about 4 points on the plateau.

In our laboratory the titration is performed in a 4 ml cuvette, starting with 2 ml of labeled

DNA in buffer, and then adding protein until the reaction plateaus or until the capacity of the

cuvette is reached. One has to correct the protein concentration at each point by the dilution

34

factor. Generally, however, one does not have to worry about dilution effects on the fluorophore.

An alternate strategy is also commonly used, where one removes volume from the cuvette as the

titration proceeds, in order to avoid an overfilled cuvette. With this procedure, however, it is

easier to dilute the fluorophore to the point where it might become problematic. One diagnostic

for such a problem is obtaining a sloped plateau region. A simple background titration (with

buffer but no protein) will confirm whether a sloped plateau is a fluorophore dilution problem. If

fluorophore dilution seems to be the source of the problem, it can be corrected by simultaneously

adding fluorophore with each protein addition so that the fluorophore concentration remains

constant. A positively sloped plateau may, however, be indicative of higher order

oligomerization or aggregation, while a negatively sloped plateau may be indicative of a

contaminating nuclease activity.

2.4.4 Data Analysis

We typically normalize all data prior to analysis. This makes it more straight-forward to

simultaneously graph and compare isotherms from different conditions, where the ΔAT might

change slightly. Large changes in ΔAT, however, should not be ignored, as they can be

diagnostic of linked processes such as oligomerization of the DNA or protein or large

conformational changes. So, for example, one might examine a protein-DNA interaction over an

800 mM salt range and observe a range of ΔAT values between 0.1 and 0.15. If one observes

large changes in ΔAT as one changes solution conditions, one should suspect a linked reaction.

Most published studies fit the resultant isotherm (ΔA versus [protein]) to the full

quadratic expansion of the binding polynomial derived for total concentrations of reactants:

ΔA= ΔAT/2DT{(ET+DT+ Kd) –[(ET+DT+ Kd)2–4ETDT]

1/2} (Eq. 2.2)

where ΔA is the change in anisotropy, ΔAT is the total anisotropy change, ET is the total

polymerase concentration at each point in the titration, DT is the total DNA concentration, and Kd

35

is the dissociation constant (this is a slight rearrangement of the equation as used by Heyduk and

Lee) (133).

In many protein-DNA binding titrations, however, the concentration of the DNA is far

below the Kd for the reaction. If this is the case, it is easier to use the binding polynomial derived

for free reactant concentrations, and assume that Efree = Etotal.

ΔA={ΔAT (E/ Kd)/(1+ E/ Kd)} (Eq. 2.3)

where ΔA is the change in fluorescence anisotropy, ΔAT is the total change in anisotropy, E is

the total polymerase concentration at each point in the titration, and Kd is the dissociation

constant for polymerase-DNA binding. If the [DNA] is 10X lower than the Kd, the error incurred

by using this equation and making this approximation is 10%. If the [DNA] is 100X below the

Kd, the incurred error is 1%, etc. One advantage of using Equation 2.3 is that it is easily modified

to include a Hill coefficient to test for cooperative/multi-site binding:

ΔA={ΔAT (EnH

/ Kd nH

)/(1+ EnH

/ Kd nH

)} (Eq. 2.4)

where nH is the fitted Hill coefficient. It is also easily modified for competitive binding. A

particular hazard of using Equation 2.2 is its too frequent use to obtain Kd values for a binding

reaction under stoichiometric binding conditions. If the concentrations of both reagents (protein

and DNA) are far above the Kd value for their association, the binding is stoichiometric. When

Equation 2.2 is used to analyze stoichiometric binding curves, both DT and Kd are allowed to

vary, and the binding stoichiometry is determined as the ratio of the fitted DT and the known DT

(see Figure 2.6).

While determining the stoichiometry of the reaction, as in Figure 2.6, is an important

control, the Kd values obtained from fits to stoichiometric data are frequently unreliable. This is

because even small errors in the concentrations of the reactants (DT and ET) are propagated into

large errors on Kd. For example, if the concentrations of reactants are in the 10 micromolar

36

range, and the true Kd is in the nanomolar range, and the error on determining the protein

concentration is a standard ± 10%, then the error on the fitted Kd is not ± 10% of the Kd, it is ±

10% of the protein concentrations. So one might obtain a fitted Kd of 10 nM, but the true error on

that value is ± several micromolar. Error in the DNA concentration propagates into the Kd in the

same manner.

0

0.2

0.4

0.6

0.8

1

0 20 40 60 80 100 120 140

No

rmalized

An

iso

tro

py

[Klentaq] ( M)

Figure 2.6: Determination of binding stoichiometry of Klentaq polymerase to double-stranded

DNA (63/63-mer). The titration was performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 75

mM KCl at pH 7.9. The DNA concentration used in the titration was 20 μM ([DNA] >> Kd). The

binding constant for Klentaq polymerase binding to 63/63-mer under “equilibrium titration”

conditions is 29.2 ± 1.8 nM. The data were fit to Equation 2.2 in the text. The ratio of bound

Klentaq polymerase to the 63/63-mer double-stranded DNA is 1.2. This figure is reprinted from

reference 167 with permission.

Variants of Equation 2.2 are frequently used to obtain Kd values under stoichiometric

titration conditions in titration calorimetry. Such Kd values must be eyed with extreme caution in

that technique as well. Under conditions where the reactant concentrations are near the Kd, or

more typically where one reactant is below the Kd and the other is titrated through the Kd, then

Equation 2.2 is perfectly applicable.

37

Equation 2.2 was used to determine the stoichiometry of Klenow and Klentaq binding to

different DNA structures (Figures 3.1 and 3.2, and Table 3.2) while Equation 2.3 was used to

determine the Kd values of different DNA structures binding by Klenow and Klentaq

polymerases in Chapter 3.

2.4.5 Other Controls

Ideally, there should be no change in steady state fluorescence when the labeled DNA

binds protein. This is easy to test, and affords assurance that one is monitoring DNA-protein

binding and not protein-dye interactions. Changes in steady state fluorescence could be either

due to protein-dye interactions, or due to propagated conformational changes in the DNA upon

binding. One does not want to be in the situation of studying binding to the fluorophore instead

of binding to the DNA. One way to troubleshoot this possibility is to label the DNA with

different dyes. Often a fluorophore can be found that shows an anisotropy change upon complex

formation, but does not show a steady state fluorescence change. Alternately, if the same DNA

labeled with several chemically different fluorophores yield the same results (i.e. the same Kd), it

is highly likely that one is observing protein-DNA interactions and not protein-dye interactions.

We have found that the storage life of fluorescently labeled DNA oligomers can vary

from several months to several years. The most reliable diagnostic of a problem with a stock of

labeled DNA is a change in the initial fluorescence anisotropy (before addition of any protein). If

the initial anisotropy changes by more than about 20% from when the labeled DNA was first

used, it probably should be discarded.

Another useful control, but one often not mentioned in published studies, is a test of the

ability of unlabeled DNA to compete effectively with the fluorescently labeled DNA. This test

can be performed as a stoichiometric titration or an equilibrium titration. In the stoichiometric

competition, labeled DNA is supplemented by exactly the same concentration of unlabeled

38

DNA, and then this mixture is titrated with protein. If the protein binds both DNAs, the apparent

stoichiometric breakpoint will exactly double (99).

This unlabeled DNA competition control can also be performed as an equilibrium

titration. Figure 2.7 shows an example where unlabeled DNA is titrated into an equilibrium of

labeled DNA + protein that is at its Kd value. The fitted KI for the unlabeled DNA should be

similar to the previously determined Kd for the labeled DNA.

0

0.2

0.4

0.6

0.8

1

0 50 100 150 200 250 300

pt-DNAhp-DNA

No

rma

lized

An

iso

tro

py

[DNA] (nM)

Figure 2.7: Fluorescently labeled primer-template DNA (a 13/20mer labeled with ROX) being

displaced from Klentaq DNA polymerase by identical, but unlabeled, primer-template DNA

(circles) and an unlabeled hairpin DNA structure (squares). The competition titrations were

performed at 25°C in 10 mM Tris and 5 mM MgCl2 at pH 7.9. The DNA sequences of the

primer-template DNA (pt-DNA) and the hairpin DNA (hp-DNA) are very similar. The cuvette

initially contained 1 nM labeled primer-template DNA (13/20-mer DNA) plus Klentaq

polymerase at the Kd. The unlabeled competitor DNA was then titrated into the cuvette.

Additional Klentaq polymerase and 1 nM labeled primer-template DNA are included in each

addition so that their concentrations remain constant throughout the titration. KI values obtained

from fits of the data are 17.5 nM (ΔG = -10.6 kcal/mole) for the unlabeled primer-template and

29.1 nM (ΔG = -10.25 kcal/mole) for the hairpin DNA. This figure is reprinted from reference

167 with permission.

39

There are numerous published studies where these simple unlabeled DNA competition

controls are either not performed or not mentioned. One risks studying the binding of protein to

the fluorophore instead of the DNA in such cases. For example, in the case of the DNA

polymerases studied in our laboratory, fluorescein-labeled DNA is not equivalently displaced by

unlabeled DNA, whereas ROX-labeled DNA is (99).

2.4.6 Competition Experiments

A natural extension of the unlabeled DNA competition controls described above is the

ability to use the fluorescence anisotropy assay in competitive mode to measure the Kd’s of a

series of unlabeled DNA oligomers for a protein. In this application, the cuvette initially contains

1 nM labeled DNA, plus protein either at its Kd value (50% saturation) or just at saturation (

≈0.95). The unlabeled competitor DNA is then titrated into the cuvette. Additional protein and 1

nM labeled pt-DNA are included in each titrant addition so that their concentrations remain

constant at all times. The anisotropy will decrease as the unlabeled DNA competes with labeled

DNA to bind the protein. Figure 2.7 also shows an example of such competitive binding

experiments, where labeled DNA is displaced by an unlabeled oligomer with a different

sequence or structure. The method of analysis depends upon the exact procedure used (i.e.

whether protein was at the Kd or near saturation with the labeled DNA, whether labeled DNA

and protein concentrations maintained constant, etc.) (151, 152). Competition exaperiments were

used to measure the Kd’s of hairpin DNA binding by Klenow and Klentaq (Tables 3.9 and 3.16).

2.5 Other Applications of Fluorescence Anisotropy to the Study of Protein-DNA

Interactions

Several groups have reported use of fluorescence anisotropy in a high throughput mode

for use in drug screening by adapting the method for use in fluorescent plate readers (153-156).

DNA as well as other small, fluorescently labeled ligands have been used as the anisotropic

probe in such assays.

40

Time-resolved anisotropy has also been used, mostly by Millar and associates (58, 157)

to study protein-DNA interactions. In this application, decay of the anisotropy is monitored

versus time after a single pulse of polarized light, similar to the way one might perform a

fluorescence lifetime experiment. The decay of the anisotropy versus time is monitored.

Deviations from a single exponential decay can be indicative of multiple binding modes.

Resolution of the number of different anisotropic decays observed, and their relative fractions of

the total decay, can provide information on the relative populations of the different binding

modes or mixed conformer sub-populations. This potentially promising application is only

beginning to see widespread use. The method used by Millar and co-workers is discussed in

Sections 1.2 and 4.1.

A relatively new application of fluorescence anisotropy to ligand binding is its adaptation

into solid phase assays (158). Since DNA or RNA that is immobilized at one end can still rotate

and pivot, one can still obtain an increase in anisotropy signal if a protein binds to the

immobilized nucleic acid. This new adaptation is already seeing widespread application.

41

CHAPTER 3

THERMODYNAMICS OF DIFFERENT DNA STRUCTURES BINDING

BY KLENOW AND KLENTAQ POLYMERASEB

3.1 Introduction

Escherichia coli DNA polymerase I (Pol I) possesses three enzymatic activities: a 5' → 3'

DNA polymerase activity, a 3' → 5' exonuclease activity that mediates proofreading, and a 5' →

3' nuclease activity required for nick translation during DNA repair. Pol I fills in the large gaps

between the Okazaki fragments during lagging strand synthesis (19). The ability of DNA Pol I to

synthesize DNA at nicks and short gaps are part of Pol I’s role in short patch DNA repair

(Nucleotide Excision Repair) (159). Pol I’s primary in vivo role is believed to be gapped DNA

repair (159). Pol I has long served as a central model for understanding the general mechanism

of DNA replication.

Removal of the 5' → 3' nuclease domains from the full length Pol I DNA polymerases

from Escherichia coli and Thermus aquaticus yields the Klenow and Klentaq large fragment

domains (47, 48). Structural and biochemical studies have shown that Klenow possesses both 5'

→ 3' polymerase and 3' → 5' exonuclease (editing) activities (32, 45, 65, 160) while Klentaq

only possesses the 5' → 3' polymerase activity (53). The polymerase and editing activities of

Klenow are located in different structural domains of the protein, separated by approximately 30-

35Å. Co-crystal structures are available for Klenow in the editing mode (43, 78) and for Klentaq

in several individual steps of the polymerization mode (51, 53). A great deal of recent

investigation of Klenow has centered on how the bound DNA transitions between the

_____________________________

BParts of this chapter have appeared in Biophysical Journal, 90, K. Datta, A. J. Wowor, A. J.

Richard, and V. J. LiCata, Temperature dependence and thermodynamics of Klenow

polymerase binding to primed-template DNA, 1739-1751, Copyright 2006, and are used with

permission of Elsevier.

42

polymerization and proofreading sites (58-62), and on which individual step in the Klenow

polymerization reaction is the authentic rate-limiting step (69, 161-165).

X-ray crystal structures of Klenow and Klentaq polymerases show that these polymerases

have very similar structures (27, 32, 51), although Klenow is a mesophilic protein that denatures

between 40-62°C, depending on solution conditions, while Klentaq is a thermophilic protein and

is stable to >100°C (49, 50). The polymerase domains of these proteins share a common

architectural feature that resembles a "half-open right hand”; with “fingers,” “thumb,” and

“palm” subdomains (46). The “thumb” subdomain binds the duplex region of DNA while the

“fingers” subdomain binds the incoming dNTP (32, 43, 53). The “palm” subdomain, consisting

of the conserved active site residues, orients the primer strand for phosphodiester bond formation

(32, 43). Several biochemical, crystallographic, and spectroscopic studies have examined the

interactions of DNA with the editing domain of Klenow (57, 75, 78, 83, 166). The co-crystal

structure of Klenow in editing mode shows the last four nucleotides of the primer strand melted

out of a duplex DNA, and bound to the editing domain (75, 78). A few recent studies, however,

have questioned whether this number is absolute, or if it can be shorter (3 base pairs melted) (64)

or longer ( 6 base pairs melted) (Richard and LiCata, unpublished data).

Previous studies in our laboratory have shown that 1) Klenow binds approximately 150X

tighter to primer-template DNA than Taq/Klentaq across a wide range of salt conditions and

temperatures; 2) the KCl and MgCl2 “sensitivities” and linkages (∂ln(1/Kd)/∂ln[KCl]) differ for

the two polymerases; 3) the two proteins both have unusually high ΔCp’s of binding to pt-DNA;

and 4) at their physiological temperatures, the DNA binding of both proteins is enthalpy driven

(97, 98, 167).

The heat capacity change (ΔCp) is the temperature dependence of the enthalpy of a

reaction. Higher ΔCp of binding has often been shown to be associated with sequence specific

43

DNA binding, however, we have previously shown that Klenow and Klentaq are two of several

non-sequence specific DNA binding proteins that show a substantial heat capacity change upon

binding (98, 167). To understand further the DNA binding thermodynamics of these

polymerases, we have characterized the binding of Klenow and Klentaq to different DNA

structures: including single-stranded, primer-template, and blunt-end double-stranded DNA.

The thermodynamic profiles for a protein-DNA interaction can include changes in free

energy (ΔG), enthalpy (ΔH), entropy (ΔS), heat capacity (∆Cp), and linked ion release upon

binding. Klenow binds primer-template DNA (pt-DNA) and blunt-end double-stranded DNA

(ds-DNA) with different thermodynamic profiles while Klentaq binds these DNAs similarly.

Klenow binds pt-DNA more tightly than ds-DNA at all salt concentrations, while Klentaq binds

these two DNA structures identically at all salt concentrations. For both proteins, binding of

single-stranded DNA shifts from weakest to tightest binding of the three structures as the salt

concentration increases. The fact that Klenow can bind DNA in both polymerase and editing

modes while Klentaq can only bind DNA in polymerase mode does not completely explain their

different thermodynamics and DNA structural selectivity.

3.2 Materials and Methods

3.2.1 Materials

3.2.1.1 Preparation of Oligonucleotides

Oligo(deoxyribo)nucleotides were obtained from Integrated DNA Technologies

(Coralville, IA). Oligonucleotides concentrations were determined spectrophotometrically using

the ε260 values provided by the company. “Primer/template” and “double stranded” DNAs were

prepared by combining an equal volume of 20 µM of each strand. These samples were heated at

94°C for 5 minutes and allowed to cool to room temperature. Hairpin-DNAs were also heated to

94°C for 5 minutes and allowed to cool to room temperature before use. Primer-template DNA

44

(pt-DNA) is a duplex DNA with 5'-overhang while double-stranded DNA (ds-DNA) in this study

is a duplex DNA with blunt-ends. hp-39 is a hairpin DNA with 5'-overhang (pt-DNA) while hp-

46 is a blunt-end hairpin (ds-DNA). pt-13/20 is comparable in size (molecular weight) to hp-39,

and ds-20/20 is comparable to hp-46. The DNA constructs used for experiments are shown in

Table 3.1.

DNA is labeled with Rhodamine-X NHS (ROXN) Ester at 5'-end of the primer for

fluorescence anisotropy (97-99, 167) while unlabeled DNA is used for isothermal titration

calorimetry and competition assays.

Table 3.1: DNAs used for binding experiments.

Single-Stranded DNA (ss-DNA)

13-mer 5’-TCGCAGCCGTCCA-3’

20-mer 5’-TCGCAGCCGTCCAAGGGTTT-3’

63-mer 5’-TACGCAGCGTACATGCTCGTGACTGGGATAACCGTGCCGTTTGCCGACTTTCGCAGCCGTCCA-3’

Primer-Template DNA (pt-DNA)

13/20-mer 5’-TCGCAGCCGTCCA-3’

3’-AGCGTCGGCAGGTTCCCAAA-5’

63/70-mer 5’-TACGCAGCGTACATGCTCGTGACTGGGATAACCGTGCCGTTTGCCGACTTTCGCAGCCGTCCA-3’

3’-ATGCGTCGCATGTACGAGCACTGACCCTATTGGCACGGCAAACGGCTGAAAGCGTCGGCAGGTTCCCAAA-5’

hp-39 AAGGCTACCTGCATGA-3’

AGCCGATGGACGTACTACCCCCC-5’

Blunt-End Double-Stranded DNA (ds-DNA)

20/20-mer 5’-TCGCAGCCGTCCAAGGGTTT-3’

3’-AGCGTCGGCAGGTTCCCAAA-5’

63/63-mer 5’-TACGCAGCGTACATGCTCGTGACTGGGATAACCGTGCCGTTTGCCGACTTTCGCAGCCGTCCA-3’

3’-ATGCGTCGCATGTACGAGCACTGACCCTATTGGCACGGCAAACGGCTGAAAGCGTCGGCAGGT-5’

hp-32 AAGGCTACCTGCATGA-3’

AGCCGATGGACGTACT-5’

hp-46 AAGGCTACCTGCATGATAATTGG-3’

AGCCGATGGACGTACTATTAACC-5’

45

3.2.1.2 Preparation of Klenow and Klentaq Polymerases

Klenow Fragment (KF) and Klentaq (KTQ) were purified in our laboratory (99). The

Klenow clone used in this study contains the D424A mutation (Klenow exo-) and was provided

by Catherine Joyce from Yale University. This mutation eliminates Klenow’s 3' → 5'

exonuclease activity, but does not abolish DNA binding ability to the proofreading site (45).

Klenow exo- does not degrade DNA substrate unlike its wild-type. Klentaq does not have an

active 3' → 5' exonuclease activity. The Klentaq clone was obtained from the American Type

Culture Collections, constructed by Wayne Barnes from Washington University. The

purifications of Klenow and Klentaq polymerases followed some published procedures (47,

168). Klenow exo- was overexpressed in Escherichia coli strain CJ376 and induced by heat

(169). The protein was purified similar to the full-length Pol I without the DEAE-cellulose

column step. Klentaq polymerase was purified using Barnes procedure (47, 168) without the

ammonium sulfate precipitation step and with the addition of Bio-Rex 70 column at pH 9.1 after

the heparin column step (99). No surfactants were added in the purifications, storages, and

experiments of both polymerases. Protein concentrations were determined

spectrophotometrically using the measured 280 nm absorptions and ε280 values of 5.88 x 104 M

-1

cm-1

for Klenow and 7.04 x 104 M

-1 cm

-1 for Klentaq. Klenow and Klentaq were stored at -20°C

before use.

3.2.2 Methods

3.2.2.1 Fluorescence Anisotropy

DNA constructs used for equilibrium DNA binding titrations are shown in Table 3.1. For

all titrations, DNA is fluorescently labeled and its concentration is 1 nM in the cuvette. The

proteins are titrated into the DNA. The anisotropy increases as the protein-DNA complexes are

formed. The data are analyzed using a single site isotherm using the program Kaleidagraph to

46

obtain the dissociation constant (Kd) (167). In addition to these direct titrations, competitive

titrations were also performed using fluorescence anisotropy, and are described below.

Stoichiometric binding curves, when [DNA] > Kd, are fitted to the equation (Figure 3.1),

ΔA = ΔAT/2DT {(ET + DT + Kd) – (ET + DT + Kd)2 – 4 ET DT]

1/2} (Eq. 3.1)

where ΔA is the change in anisotropy, ΔAT is the total anisotropy change, ET is the total

polymerase concentration at each point in the titration, DT is the total DNA concentration,

and Kd

is the dissociation constant (133, 170). DT and Kd are allowed to vary,

and the binding

stoichiometry is determined as the ratio of the fitted DT and the known DT.

It should be noted that obtaining Kd values from fits to such stoichiometric binding

curves is not advisable as all the error on measured protein and DNA concentrations is reflected

in the obtained Kd value in this equation (since the effective free DNA and protein

concentrations are << ET or DT) and these error windows are generally significantly larger than

the nanomolar range of the Kd values.

Kd values are thus obtained from equilibrium binding curves, when the [DNA] << Kd,

which are fit to the equation,

ΔA = {ΔAT (ET/Kd)/(1 + ET/Kd)} (Eq. 3.2)

where ΔA is the change in fluorescence anisotropy, ΔAT is the total change in anisotropy, ET is

the total polymerase concentration at each point in the titration, and Kd

is the dissociation

constant for polymerase-DNA binding. All fluorescence anisotropy experiments are replicated at

least three times.

Linked ion release upon binding of the polymerases to DNA is calculated using a basic

linkage relationship (92, 171-173),

{∂ln(1/Kd)}/{∂ln[KCl]} = Δnions = ΔnK+ + ΔnCl

- (Eq. 3.3)

47

The slope of a plot of ln (1/Kd) versus ln [KCl] gives the net number of ions (Δnions) that are

bound or released when the protein-DNA complex is formed.

Figure 3.1: Determination of binding stoichiometry for Klentaq polymerase binding to double-

stranded DNA (ds-20/20). Shown are fluorescence anisotropy-monitored stoichiometric titrations

of Klentaq polymerase into 100 nM ROX-labeled DNA (●) and an unlabeled DNA competition

control titration containing 100 nM ROX-labeled plus 100 nM unlabeled DNA (■). The titrations

were performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 75 mM KCl at pH 7.9. The data is

fitted to Equation 3.1.

Competition assays were used to assess the effects of Mg2+

on binding to the different

DNA constructs (Figure 3.10 and Table 16). Competition assays were performed in 10 mM Tris

and 5 mM KCl at pH 7.9, in the absence or presence of 5 mM MgCl2. The cuvette initially

contains 1 nM labeled pt-DNA 13/20-mer and protein either at the Kd or at saturation. The

unlabeled competitor DNA is then titrated into the cuvette. Additional protein and 1 nM labeled

pt-DNA are included in each addition so that their concentrations remain constant at all times.

48

The anisotropy decreases as the unlabeled DNA competes with labeled DNA to bind the protein.

Competition curves are fit to the equation,

ΔA = {ΔAT ([I]/KI)/(1 + [I]/KI + ET/Kd)}

(Eq. 3.4)

where, ΔA is the change in fluorescence anisotropy, ΔAT is the total change in anisotropy, [I] is

the total competitor concentration at each point in the titration, KI is the inhibition constant for

the polymerase-competitor binding, ET is the total polymerase concentration at each point in the

titration, and Kd is the dissociation constant for polymerase-DNA binding.

The Gibbs free energy is calculated using the equation,

ΔG = –RT ln (1/Kd)

(Eq. 3.5)

where, ΔG is the Gibbs free energy, R is the gas constant (1.987 cal K-1

mol-1

), T is the

temperature in Kelvin, and Kd is the dissociation constant for polymerase-DNA binding.

The temperature dependence of the free energy of DNA binding is analyzed using an

integrated Gibbs-Helmholtz equation with a temperature independent ΔCp (heat capacity change)

as described previously (98),

ΔG(T) = ΔHrefT – TΔSrefT + ΔCp [T – TrefT – T ln(T / TrefT)]

(Eq. 3.6)

where, ΔG(T) is the free energy at each temperature, T is the temperature in Kelvin, ΔCp is the

heat capacity, and ΔHrefT and ΔSrefT are the fitted van’t Hoff enthalpy and entropy values at any

chosen reference temperature TrefT.

3.2.2.2 Isothermal Titration Calorimetry (ITC)

ITC is used to measure the enthalpy (ΔH) associated with intra- and inter-molecular

interactions. DNA used for ITC DNA binding titrations are shown in Table 3.1. For ITC

titrations, the DNA is not labeled and its concentration is 2.5 μM in the sample cell. The

concentration of protein used for most titrations is 75 μM. Protein is titrated into the DNA. Heat

49

is produced as the protein-DNA complexes are formed. The data are analyzed using a single-site

model using the program Origin.

The enthalpies of binding (ΔHcal) obtained from the fit are plotted as a function of

temperature to directly obtain the calorimetric heat capacity change for the binding process

(∂ΔHcal/∂T = ΔCpcal). All ΔHcal values gathered using ITC are the average of a minimum of three

experiments.

3.2.2.3 Electrophoretic Mobility Shift Assay

DNAs used in these experiments are labeled with -32

P-ATP at the 5'-end using T4

polynucleotide kinase. Each 10 µL reaction contains 5 nM DNA and increasing amounts of

protein. The composition of the binding buffer is 10 mM Tris, 5 mM MgCl2, and 50 mM KCl, at

pH 7.9, 25°C. The incubation time is 30 minutes. After incubation, the samples were loaded onto

an 8% acrylamide (39:1 acrylamide:bis) gel and electrophoresed in 0.5X TBE buffer (45 mM

Tris Borate, 1 mM EDTA, pH 8.0). The gel was run at constant voltage of 175 volts for 1.5 hours

at 4°C. The image was obtained using a Storm PhosphorImager and bands were quantified using

the program Image Quant. Background was subtracted from the band intensities measured.

Fractional complex formation as a function of protein concentration was analyzed using a single-

site isotherm with the program Kaleidagraph. All gel shift experiments were repeated at least

twice.

3.3 Results and Discussion

3.3.1 The Binding Stoichiometry of Klenow and Klentaq Polymerases to Different DNA

Structures

Figure 3.2 shows representative stoichiometric binding curves of ds-63/63 binding by

Klenow and Klentaq polymerases. Unlike equilibrium titrations where one reactant is kept well

below the Kd, here both protein and DNA concentrations are >> Kd to ensure saturation /

50

stoichiometric binding. Data are then fit with Equation 3.1 in Materials and Methods to

determine the binding stoichiometry, which is 1:1 for all complexes examined. The steady-state

fluorescence of ROX does not change when protein is added indicating that the protein is not

interacting with the dye. Furthermore, unlabeled DNA added to such stoichiometric titrations

competes directly with the ROX-labeled DNA (data not shown).

0

0.2

0.4

0.6

0.8

1

0 20 40 60 80 100 120 140

KlenowKlentaq

No

rmalize

d A

nis

otr

op

y

[Protein] M

Figure 3.2: Determination of binding stoichiometries for Klenow and Klentaq polymerases

binding to double-stranded DNA (ds-63/63). Shown are fluorescence anisotropy-monitored

stoichiometric titrations of and Klentaq (■) and Klenow (●) polymerases into 20 µM and 30 µM

ROX-labeled DNA, respectively. The titrations were performed at 25°C in 10 mM Tris, 5 mM

MgCl2, and 300 mM KCl (Klenow) or 75 mM KCl (Klentaq) at pH 7.9 and fitted with Equation

3.1 in the text.

51

Table 3.2: Stoichiometric ratios of protein:DNA binding determined using fluorescence

anisotropy and Isothermal Titration Calorimetry (ITC)a. ND: Not Determined.

Fluorescence Anisotropy ITC

DNA Klenow Klentaq Klenow Klentaq

pt-13/20 0.89b 1.0

b 0.8 0.9

pt-63/70 1.1 1.15b 1.0 1.35

ss-20 ND ND 0.9 1.35c

ss-63 ND 0.8 0.8 ND

ds-20/20 0.6 1.0 1.2 1.1c

ds-63/63 0.75 1.25 0.9 ND

aAll errors are < ±0.1 except as noted by superscript “c”.

bFrom reference 99.

cKlentaq ss-20: ±0.12, ds-20/20: ±0.25.

The stoichiometries of Klenow and Klentaq polymerases binding to different DNA

structures were also obtained from isothermal titration calorimetry. Numerical values are given

in Table 3.2. Both fluorescence anisotropy and isothermal titration calorimetry indicate that both

Klenow and Klentaq polymerases form 1:1 complexes with these DNAs. Unusually low or high

values returned by one technique (e.g. Klenow + ds-20/20 by fluorescence anisotropy, or Klentaq

+ pt-63/70 by ITC) are generally offset by the other. Even considering such outliers, there is no

strong evidence for protein:DNA ratios greater than 1:1. These results agree with earlier direct

determinations of binding stoichiometry by our lab (99) and from von Hippel and associates

(64), but conflict with reports of Klenow dimerization from Millar and associates (131).

Analytical ultracentrifugation and small angle X-ray scattering also report that Klenow and

Klentaq bind these DNAs with 1:1 stoichiometry (Wowor and LiCata, and Richard and LiCata in

preparation). It is particularly noteworthy that 1:1 binding stoichiometry is maintained even on

the longer constructs (ds-63mer and pt-63/70mer), where one might expect that proteins could

bind to both ends of the construct. This, however, is clearly not the case (Figure 3.2). DNA

constructs longer than 63/70-mer have not yet been examined with either polymerase, but no

52

complexes with higher than 1:1 stoichiometry were detected by fluorescence anisotropy,

calorimetry, or gel shift assays for either polymerase with any DNA constructs in this study.

3.3.2 DNA Structural Selectivity

The DNA structures used in this study are single-stranded DNA (ss-DNA), primer-

template DNA with a 7 base ss-overhang (pt-DNA), and blunt-end double-stranded DNA (ds-

DNA). The fluorescence anisotropy binding assay, for these proteins, will resolve binding

affinities and produce well behaved titration curves across the nanomolar range (~10 nM to 1

µM) (35). Because the binding affinity of Klentaq for DNA is consistently weaker than that of

Klenow, to obtain data in the same relative affinity range requires titrating the two proteins

across different salt concentration ranges. Figure 3.3 shows Klenow and Klentaq binding to these

DNA structures at 25°C at two different salt concentrations for each protein. At both KCl

concentrations shown in Figure 3.3, the binding affinity trend for Klentaq is ds-DNA ≈ pt-DNA

>> ss-DNA. For Klenow, increasing the salt concentration across the experimentally accessible

range produces a change in the binding hierarchy, and curves are shown at 200 mM KCl and 300

mM KCl (see Tables 3.3-3.6 for Kd values). Although examination of the binding curves within

these experimentally accessible windows suggests differing structural selectivity for the two

polymerases, such substrate affinity hierarchies are dependent on the salt concentration. In the

next section, we show that one observes very similar binding hierarchy patterns for the two

proteins when one examines binding trends over very wide salt concentration ranges.

Table 3.3: The binding constants (Kd) and free energies (∆G) of binding of Klenow polymerase

to different DNA structures at 25°C in 10 mM Tris, 5 mM MgCl2, and 300 mM KCl at pH 7.9.

Titrations for ss-13, pt-13/20, and ds-20/20 are shown in Figure 3.3A.

DNA Kd (nM) ΔG (kcal/mol) DNA Kd (nM) ΔG (kcal/mol)

ss-13 4.9 ± 0.1 -11.3 ± 0.01 ss-63 1.9 ± 0.2 -11.8 ± 0.06

pt-13/20 9.8 ± 0.3 -10.9 ± 0.02 pt-63/70 3.6 ± 0.1 -11.5 ± 0.02

ds-20/20 29.5 ± 0.4 -10.3 ± 0.01 ds-63/63 6.4 ± 0.4 -11.2 ± 0.04

53

Figure 3.3: DNA structure dependence of binding by Klenow and Klentaq polymerases. Shown

are representative equilibrium titrations of the polymerases and single-stranded DNA (ss-13) (●),

primer-template DNA (pt-13/20) (■), and double-stranded DNA (ds-20/20) (♦). Panels A and C:

Klenow titrations performed at 25°C in 10 mM Tris, 5 mM MgCl2 and 300 mM KCl (Panel A)

or 200 mM KCl (Panel C) at pH 7.9. Panels B and D: Klentaq titrations performed at 25°C in 10

mM Tris, 5 mM MgCl2 and 75 mM KCl (all curves in Panel B), 25 mM (pt-DNA and ds-DNA in

Panel D), or 5 mM KCl (ss-DNA in Panel D) at pH 7.9. Lines show the fits to single-site

isotherms (Equation 3.2).

54

Table 3.4: The binding constants (Kd) and free energies (∆G) of binding of Klentaq polymerase

to different DNA structures at 25°C in 10 mM Tris, 5 mM MgCl2, and 75 mM KCl at pH 7.9.

Titrations for ss-13, pt-13/20, and ds-20/20 are shown in Figure 3.3B.

DNA Kd (nM) ΔG (kcal/mol) DNA Kd (nM) ΔG (kcal/mol)

ss-13 1378.7 ± 154.3 -8.0 ± 0.06 ss-63 441.5 ± 27.7 -8.7 ± 0.04

pt-13/20 43.6 ± 0.9 -10.0 ± 0.01 pt-63/70 39.3 ± 1.2 -10.1 ± 0.02

ds-20/20 43.5 ± 0.9 -10.0 ± 0.01 ds-63/63 29.2 ± 1.8 -10.3 ± 0.03

Table 3.5: The binding constants (Kd) and free energies (∆G) of binding of Klenow polymerase

to different DNA structures at 25°C in 10 mM Tris, 5 mM MgCl2, and 200 mM KCl at pH 7.9.

Titrations are shown in Figure 3.3C.

DNA Kd (nM) ΔG (kcal/mol)

ss-13 2.1 ± 0.1 -11.8 ± 0.03

pt-13/20 1.4 ± 0.1 -12.1 ± 0.04

ds-20/20 2.9 ± 0.2 -11.6 ± 0.04

Table 3.6: The binding constants (Kd) and free energies (∆G) of binding of Klentaq polymerase

to different DNA structures at 25°C in 10 mM Tris, 5 mM MgCl2, and 25 mM KCl (pt- and ds-

DNA) or 5 mM KCl (ss-DNA) at pH 7.9. Titrations are shown in Figure 3.3D.

DNA Kd (nM) ΔG (kcal/mol)

ss-13 113.7 ± 22.6 -9.5 ± 0.11

pt-13/20 4.6 ± 0.4 -11.4 ± 0.05

ds-20/20 6.9 ± 0.4 -11.1 ± 0.03

3.3.3 KCl Dependence of DNA Binding by Klenow and Klentaq Polymerases

Figures 3.4A and 3.4B show the thermodynamic linkage plots for binding of different

DNA structures by Klenow and Klentaq polymerases as a function of KCl concentration

(∂ln(1/Kd) versus ∂ln[salt]). The negative slopes of the linkage plots indicate the net ion release

upon protein-DNA complex formation. The linkage plots for Klenow show that binding to ss-

DNA is linked to the release of 2.1 ions while binding to pt-DNA and ds-DNA is linked to the

release of 4.4 and 5.4 ions, respectively (Tables 3.7 and 3.8). The linkage plots for Klentaq

indicates that binding to ss-DNA releases 1.0 ion, binding to pt-DNA releases 2.8 ions, and

binding to ds-DNA releases 3.2 ions. The Kd values and associated error windows are reported in

Tables 3.7 and 3.8, and in reference 99 for pt-DNA. For each protein, the ion releases for pt- vs.

55

ds-DNA are similar, while the ion release upon ss-DNA binding is significantly smaller. Klentaq

consistently releases fewer ions when binding the same DNA, suggesting either a smaller

binding footprint on the DNA for Klentaq, or a linked ion uptake by the protein in Klentaq.

Figure 3.4: KCl linkages (∂ln1/Kd versus ∂ln[salt]) for the binding of Klenow (A) and Klentaq

(B), and polymerases to ss-DNA, pt-DNA, and ds-DNA. The slopes of the plots give the

thermodynamic net average number of ions released upon complex formation. Klenow’s

titrations were performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 200-300 mM KCl at pH 7.9

while Klentaq’s titrations were performed at 25°C and pH 7.9 in 10 mM Tris, 5 mM MgCl2, and

5 – 50 mM KCl concentration range for ss-DNA binding and 50 – 150 mM KCl concentration

range for pt-DNA and ds-DNA binding. pt-DNA data for Klenow and Klentaq include data from

reference 99. Panel C shows the salt linkages for both polymerases, re-plotted together and

extrapolated over the same salt concentration range. Because the binding affinity of Klentaq for

DNA is consistently weaker than that of Klenow, to obtain data in the same relative affinity

range requires titrating the two proteins across different salt concentration ranges. ss- and ds-

DNA binding by Klentaq data were obtained by Gregory S. Thompson.

56

Table 3.7: The binding constants (Kd) and the number of ions released when Klenow polymerase

binds different DNA structures (ss-13, pt-13/20, and ds-20/20) in the 200 – 300 mM KCl

concentration range. All titrations were performed at 25°C in 10 mM Tris and 5 mM MgCl2 at

pH 7.9. Data are those plotted in Figure 3.4A.

[KCl]

(mM)

ss-DNA Ions

Released

[KCl]

(mM)

pt-DNA Ions

Released

[KCl]

(mM)

ds-DNA Ions

Released Kd (nM) Kd (nM) Kd (nM)

200 2.1 ± 0.1

2.1 ± 0.4

200 1.4 ± 0.1

4.4 ± 0.6

200 2.9 ± 0.2

5.4 ± 0.7 225 2.2 ± 0.1 225 2.4 ± 0.1 225 8.7 ± 0.2

250 3.3 ± 0.1 250 4.0 ± 0.1 250 9.5 ± 0.4

275 3.4 ± 0.1 275 4.3 ± 0.2 275 19.4 ± 0.2

300 4.9 ± 0.1 300 9.8 ± 0.3 300 29.5 ± 0.4

Table 3.8: The binding constants (Kd) and the number of ions released when Klentaq polymerase

binds different DNA structures in the 5 – 50 mM KCl concentration range for single-stranded

DNA (ss-63) binding and the 50 – 150 mM KCl concentration range for double-stranded DNA

(ds-63/63) binding. All titrations were performed at 25°C in 10 mM Tris and 5 mM MgCl2 at pH

7.9. Data for pt-DNA (2.8 ± 0.2 ions released) are in reference 99. Data for ss- and ds-DNA were

obtained by Gregory S. Thompson. Data are those plotted in Figure 3.4B.

[KCl]

(mM)

ss-DNA Ions

Released

[KCl]

(mM)

ds-DNA Ions

Released Kd (nM) Kd (nM)

5 33.4 ± 1.3

1.0 ± 0.1

50 12.1 ± 0.3

3.2 ± 0.2 10 43.5 ± 1.9 75 29.2 ± 1.8

20 100.3 ± 1.7 100 89.3 ± 4.4

35 157.3 ± 7.2 125 202.3 ± 10.5

50 329.7 ± 7.2 150 387.2 ± 15.7

In Figure 3.4A, blunt-end double stranded-DNA binding is always weaker than pt-DNA

binding for Klenow, however, at salt concentrations lower than 225 mM KCl (< -1.5 ln [KCl]),

the relative affinities of ss-DNA vs. pt-DNA for Klenow switches. Likewise, at salt

concentrations lower than 200 mM (< -1.7 ln [KCl]), the affinity of Klenow for ss-DNA will

cross the ds-DNA line and become the lowest affinity substrate. In contrast, for Klentaq the

binding affinity hierarchy (ds-DNA ≈ pt-DNA >> ss-DNA) does not change with salt

concentration across the range examined. It can be seen in Figure 3.4B, however, that ss-DNA

binding will become the tightest substrate for Klentaq at salt concentrations ≥ 175 mM KCl,

where the binding affinities of Klentaq for DNA have decreased to the micromolar range.

57

If it is assumed that the salt linkages will remain linear, and the experimental data of

Figures 3.4A and 3.4B are extrapolated over correspondingly wide salt concentration ranges, one

immediately finds that the affinity patterns for the two proteins are very similar (see Figure

3.4C). For both polymerases, binding affinities for pt-DNA and ds-DNA are very close to each

other over several ln units of KCl concentration, while ss-DNA binding is significantly weaker at

low KCl and switches to being the tightest binding substrate at salt concentrations near or

slightly above physiological ionic strength.

3.3.4 Contributions of the Single-Stranded Region of the Template DNA to Klenow Binding

Figure 3.5: Top panel: representative gel shift assay showing hp-32 binding by Klenow

polymerase. [DNA] in each lane is 5 nM. Klenow concentrations in lanes 1-15 are: 0, 25 nM, 50

nM, 100 nM, 150 nM, 200 nM, 250 nM, 300 nM, 450 nM, 500 nM, 600 nM, 700 nM, 800 nM,

900 nM, and 1000 nM. Incubation was performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 50

mM KCl at pH 7.9. Gel was run at 4°C. A marks the gel wells, B shows the KF/DNA

complexes, and C is the DNA. Bottom panel: digitized gel shift data from the top panel, fit to a

single-site isotherm (Equation 3.2). Gel shift data were obtained by Sreerupa Ray.

58

Figures 3.4A and 3.4B also show that Klentaq binds pt-DNA and ds-DNA with nearly

identical affinity across a wide range of salt concentrations, and that Klenow binds pt-DNA ≤ 0.8

kcal/mol tighter than ds-DNA across a relatively wide range of salt concentration. This result is

consistent when we examine Klenow binding to a variety of different pt- and ds-DNAs, and with

both fluorescence anisotropy and gel shift assays. Figure 3.4C even suggests that the affinities

for pt- versus ds-DNA will reverse at very low salt.

Table 3.9 shows values for a variety of different direct measurements of the difference in

free energy of Klenow binding to pt- and ds-DNA. The mean for these measurements, with

different constructs and differing methods, is -0.77 kcal/mol. For hairpin structures, the primer-

template vs. blunt-end DNA difference is slightly larger (mean G of -0.96 kcal/mol) relative

to duplex constructs (mean G of -0.52 kcal/mol). There are no significant differences between

G values from fluorescence anisotropy versus gel shift. Klenow binding to hp-32 using gel

shift is shown in Figure 3.5. These differences between pt- and blunt-end bindings are

considerably smaller than that reported in another recent study (56), but that study did not

measure direct binding but estimated binding differences from differences in competitive

nucleotide incorporation into different constructs. The competitive nucleotide incorporation

measurements estimated a 3 kcal/mol greater affinity of Klenow for pt-DNA with a 6 bp ss-

overhang relative to blunt-end DNA. This difference would predict that if Klenow encountered

equal concentrations of pt-DNA and blunt-end DNA, it would be 170 times more likely to bind

to the pt-DNA. The direct binding results in this study, however, show that there is at most a 0.8

kcal/mol difference in binding between these two structures, which translates into only a 4 fold

greater likelihood to bind pt-DNA. This small(er) difference lends more support to the potential

physiological role of Klenow and Klentaq polymerases can participate in the protection of the ds-

end of the DNA prior to ds-DNA break repair (174, 175).

59

Table 3.9: The differences in free energies between primer-template DNA and blunt-end ds-

DNA binding by Klenow polymerase assayed via both fluorescence anisotropy and gel shift, and

using both duplex and hairpin DNA constructs. Gel shift data were obtained by Sreerupa Ray.

DNA Structurea

G

(kcal/mol)b

G

(kcal/mol)c

pt-13/20 FA -10.90 ± 0.02 -0.65

ds-20/20 FA -10.25 ± 0.01

pt-63/70 FA -11.52 ± 0.02 -0.35

ds-63/63 FA -11.17 ± 0.04

pt-13/20 GS -9.86 ± 0.09 -0.55

ds-20/20 GS -9.31 ± 0.14

hp-39 FA -11.65 ± 0.05 -1.21

hp-32 FA -10.44 ± 0.17

hp-39 FA -11.65 ± 0.05 -1.15

hp-46 FA -10.50 ± 0.13

hp-39 GS -9.64 ± 0.12 -0.84

hp-32 GS -8.80 ± 0.24

hp-39 GS -9.64 ± 0.12 -0.62

hp-46 GS -9.02 ± 0.18

aFA = fluorescence anisotropy. GS = gel shift or electrophoretic mobility shift assay. pt-13/20,

pt-63/70, and hp-39 are primer-template DNAs while ds-20/20, ds-63/63, hp-32, and hp-39 are

blunt-end, double-stranded DNAs.

bpt-13/20, ds-20/20, pt-63/70, and ds-63/63 FA titrations were performed at 25°C in 10 mM Tris,

5 mM MgCl2, and 300 mM KCl at pH 7.9 while hp-32, hp-39, and hp-46 FA titrations were

performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 5 mM KCl at pH 7.9. For gel shift data, all

titrations were performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 50 mM KCl at pH 7.9.

c

G = Gpt - Gds and so represents how much tighter the pt-DNA binds in each pair.

60

3.3.5 Enthalpies and Heat Capacities of Binding of Different DNA Structures by Klenow

and Klentaq

The calorimetric enthalpies of binding by Klenow polymerase are plotted as a function of

temperature in Figures 3.6A and 3.6B. The data were fit by linear regression to determine the

heat capacity changes (ΔCp = the slope) for Klenow binding to different DNA constructs. The

displaced dependence for ss-63 mer relative to the ss-20 mer is due to enthalpy of melting

secondary structure in the ss-63 mer construct. Thermal melts show that the ss-63 mer has some

secondary structure in solution (data not shown), and the heat required to melt the secondary

structure raises the H values for this construct, but does not alter ∆Cp (slope in Figure 3.6B).

All individual ∆H and ∆Cp values are reported in Tables 3.10 and 3.11, and in reference 98 for

pt-DNA.

Table 3.10: Calorimetric enthalpy of binding (ΔH) and heat capacity change (ΔCp) when Klenow

polymerase binds single-stranded DNA (ss-20 and ss-63). All titrations were performed in 10

mM Tris, 5 mM MgCl2, and 75 mM KCl at pH 7.9.

ss-20 ss-63

Temperature

(ºCelsius)

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

Temperature

(ºCelsius)

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

8 3.4 ± 1.1

-487 ± 53

8 13.5 ± 0.1

-430 ± 7 10 3.9 ± 1.3 10 12.8 ± 0.4

30 -6.6 ± 0.7 30 4.1 ± 0.4

Table 3.11: Calorimetric enthalpy of binding (ΔH) and heat capacity change (ΔCp) when Klenow

polymerase binds double-stranded DNA (ds-20/20 and ds-63/63). All titrations were performed

in 10 mM Tris, 5 mM MgCl2, and 300 mM KCl at pH 7.9.

ds-20/20 ds-63/63

Temperature

(ºCelsius)

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

Temperature

(ºCelsius)

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

8 2.7 ± 0.9

-329 ± 51

8 2.7 ± 0.1

-440 ± 2 10 3.3 ± 0.7 10 1.8 ± 0.3

30 -4.0 ± 0.2 30 -7.0 ± 0.01

61

The calorimetric enthalpies of binding by Klentaq polymerase are plotted as a function of

temperature in Figure 3.7A. Again the data are linearly fitted to determine the heat capacity

changes (ΔCp). The ∆H and the ∆Cp values are reported in Table 3.12. In Figure 3.7B, the

temperature dependence of the Gibbs free energy of DNA binding, determined using

fluorescence anisotropy titrations, is plotted as a function of temperature and then analyzed using

Gibbs-Helmholtz equation to obtain corresponding Gibbs-Helmholtz/van’t Hoff ΔCp values for

binding the different constructs. It should be noted that introduction of a temperature dependent

ΔCp for the ds-63/63 or ss-63 DNA does not significantly improve the fits to these data (123).

The Kd values and the thermodynamics (ΔG, ΔH, TΔS, and ∆Cp values) are reported in Tables

3.13 and 3.14, and in reference 97 for pt-DNA. The van’t Hoff enthalpies of DNA binding by

Klenow and Klentaq polymerases are larger than the calorimetric enthalpies, and this

discrepancy has previously been linked to protonation/deprotonation processes detected

calorimetrically, but not in the anisotropy assay (167).

Figure 3.6: Temperature dependence of the enthalpy change (∆H) upon binding of Klenow to

shorter and longer DNA structures determined by calorimetry. A) Binding enthalpies (∆Hcal) for

ss-20 (●), pt-13/20 (■), and ds-20/20 (♦). B) Binding enthalpies for ss-63 (●), pt-63/70 (■), and

ds-63/63 (♦). Linear fits to the calorimetric data are used to obtain the calorimetric ∆Cp. The

titrations were performed in 10 mM Tris, 5 mM MgCl2, and 75 mM KCl for ss-DNA or 300 mM

KCl for pt-DNA and ds-DNA at pH 7.9. Error bar is based on multiple measurements (see

Section 3.2.2.2). Data for both lengths of pt-DNA are from reference 98.

62

Figure 3.7: Temperature dependence of the binding of Klentaq to different DNA structures.

Panel A shows calorimetrically determined enthalpies (∆Hcal) upon binding of Klentaq to ss-20

(●), pt-13/20 (■), and ds-20/20 (♦). Linear fits to the calorimetric data are used to obtain the

calorimetric ∆Cp values. The titrations were performed in 10 mM Tris, 5 mM MgCl2, and 75 mM

KCl at pH 7.9. Panel B shows the temperature dependence of the free energy (∆G) of binding of

Klentaq to of ss-63 (●), pt-63/70 (■), and ds-63/63 (♦), determined in equilibrium titrations by

fluorescence anisotropy. Data for pt-63/70mer include data from reference 97. Lines are the fits

to the Gibbs-Helmholtz equation. A much lower salt concentration was used for ss-DNA in order

to bring the affinity of Klentaq for ss-DNA into a similar Kd range as for pt-DNA and ds-DNA.

The titrations were performed in 10 mM Tris, 5 mM MgCl2, and 5 mM KCl for ss-DNA or 75

mM KCl for pt-DNA and ds-DNA at pH 7.9. Panel C shows the van’t Hoff enthalpies (∆HvH) as

a function of temperature obtained from the Gibbs-Helmholtz analysis of the data in Panel B for

ss-63 (●), pt-63/70 (■), and ds-63/63 (♦). Shorter and longer DNA constructs yield the same

pattern.

63

Table 3.12: Calorimetric enthalpy of binding (ΔH) and heat capacity change (ΔCp) when Klentaq

polymerase binds different DNA structures (ss-20, pt-13/20, and ds-20/20). All titrations were

performed in 10 mM Tris, 5 mM MgCl2, and 75 mM KCl at pH 7.9. ND = Not Determined.

Temperature

(ºCelsius)

ss-20 pt-13/20 ds-20/20

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

ΔH

(kcal/mol)

ΔCp

(cal/mol K)

10 -1.5 ± 0.1

-75 ± 22

10.0 ± 0.8

-531 ± 129

17.1 ± 0.8

-684 ± 90 20 ND 7.5 ± 0.4 9.4 ± 0.4

30 -2.2 ± 0.1 6.6 ± 0.5 7.3 ± 0.1

40 ND -0.6 ± 0.1 -0.5 ± 0.3

50 -4.5 ± 0.1 -12.5 ± 0.3 -12.1 ± 0.2

Table 3.13: Thermodynamic parameters for temperature dependence of single-stranded DNA

(ss-63) binding by Klentaq DNA polymerase. All titrations were performed in 10 mM Tris, 5

mM MgCl2, and 5 mM KCl at pH 7.9.

Temperature

(ºCelsius)

Kd

(nM)

ΔG

(kcal/mol)

ΔH

(kcal/mol)

TΔS

(kcal/mol)

ΔCp

(cal/mol K)

5 21.8 ± 0.7 -9.8 ± 0.02 2.4 ± 2.4 12.1 ± 2.3

-400 ± 71

15 25.4 ± 1.0 -10.0 ± 0.02 -1.5 ± 1.7 8.6 ± 1.7

25 33.6 ± 1.1 -10.2 ± 0.02 -5.3 ± 1.1 5.0 ± 1.2

35 47.6 ± 1.2 -10.3 ± 0.02 -9.1 ± 0.7 1.2 ± 0.7

45 77.7 ± 1.0 -10.3 ± 0.01 -13.1 ± 0.8 -2.8 ± 0.8

55 97.0 ± 2.5 -10.5 ± 0.02 -16.7 ± 1.3 -6.5 ± 1.3

65 356.0 ± 16.8 -9.9 ± 0.03 -20.5 ± 1.9 -10.5 ± 1.9

75 908.9 ± 40.5 -9.6 ± 0.03 -24.4 ± 2.6 -14.8 ± 2.6

Table 3.14: Thermodynamic parameters for temperature dependence of double-stranded DNA

(ds-63/63) binding by Klentaq DNA polymerase. All titrations were performed in 10 mM Tris, 5

mM MgCl2, and 75 mM KCl at pH 7.9.

Temperature

(ºCelsius)

Kd

(nM)

ΔG

(kcal/mol)

ΔH

(kcal/mol)

TΔS

(kcal/mol)

ΔCp

(cal/mol K)

15 27.5 ± 2.4 -10.0 ± 0.05 13.9 ± 4.3 23.7 ± 4.2

-993 ± 152

25 29.2 ± 1.8 -10.3 ± 0.04 4.0 ± 2.8 14.4 ± 2.8

35 31.7 ± 2.3 -10.6 ± 0.04 -5.9 ± 1.6 4.8 ± 1.6

45 28.0 ± 1.9 -11.0 ± 0.04 -15.9 ± 1.2 -5.1 ± 1.2

55 94.6 ± 1.6 -10.5 ± 0.01 -25.8 ± 2.3 -15.3 ± 2.3

60 190.1 ± 11.9 -10.2 ± 0.04 -30.8 ± 3.0 -20.6 ± 3.0

65 414.4 ± 43.3 -9.9 ± 0.07 -35.7 ± 3.7 -25.9 ± 3.7

70 1189.2 ± 135.4 -9.3 ± 0.07 -40.7 ± 4.4 -31.3 ± 4.5

The detection and measurement of DNA binding linked protonation/deprotonation effects

can be accomplished by performing parallel calorimetric titrations in buffers with different

ionization enthalpies (176). DNA binding titrations were thus performed in phosphate ( Hion =

+1.22 kcal/mole), borate ( Hion = +3.31 kcal/mole), HEPES ( Hion = +5.0 kcal/mole), MOPS

64

(ΔHion = +5.3 kcal/mole), imidazole (ΔHion = +8.75 kcal/mole), and Tris (ΔHion = +11.4

kcal/mole) (109, 110). Titrations were performed at 30°C with 63/70-mer DNA in 10 mM

“buffer,” 5 mM MgCl2, and 300 mM KCl at pH 7.9. If protons are taken up or released upon

formation of the Klenow-DNA complex, calorimetric measurements should reflect an additional

heat effect due to the linked protonation or deprotonation of the buffer. Figure 3.8A shows the

dependence of Hcal as a function of Hion of the buffer. The slope of this plot yields the number

of protons taken up or released upon complex formation, and the y-intercept gives the enthalpy

of binding in absence of contributions from the buffer ionization (176). Thus, whereas the

directly measured ΔHcal in Tris buffer at this temperature is -14.1 kcal/mole, the extrapolation of

ΔHcal to zero ΔHion reports a Hbinding of -17.3 kcal/mole, indicating that +3.2 kcal/mole of the

measured enthalpy at 30°C in Tris is due to a linked buffer ionization effect. Because the

thermodynamics of 13/20-mer and 63/70-mer binding are a little different, the linked

protonation/deprotonation effect for binding to the 13/20 may be different. Linked buffer

protonation/deprotonation heats are one source of the commonly observed discrepancies between

calorimetric and van’t Hoff enthalpies (169, 176), such as those observed in these data. However,

it is notable here that whereas the ΔHcal and HvH values differ somewhat for the binding of

Klenow to DNA, the ΔCp values determined for binding to the 13/20-mer and 63/70-mer DNAs

by calorimetric and van’t Hoff analysis are quite similar (98).

The slope of plot 3.22 A is +0.25, indicating that there is a small net proton uptake by the

complex when Klenow polymerase binds 63/70-mer DNA (109, 110, 176). The magnitude of

this measured linked proton uptake will be pH dependent (176), so this linkage indicates that at

least one ionizable group becomes more protonated upon complex formation. This value of 0.25

is a thermodynamic average net change in protonation and could reflect either titration of one

65

specific group or the net difference for a number of simultaneous linked

protonation/deprotonation events.

Figure 3.8: A. Calorimetric ΔH values for Klenow-DNA binding at 30°C in buffers with

different ionization enthalpies. Measurements were made in 10 mM phosphate (ΔHion = 1.22

kcal/mole), borate (ΔHion = 3.31 kcal/mole), HEPES (ΔHion = 5.0 kcal/mole), MOPS (ΔHion = 5.3

kcal/mole), imidazole (ΔHion = 8.75 kcal/mole), and Tris (ΔHion = 11.4 kcal/mole) with 5 mM

MgCl2, and 300 mM KCl at pH 7.9. B. Temperature dependences of the calorimetric ΔH of

Klenow binding to 63/70-mer DNA in the presence (■) and absence (□) of MgCl2. The ΔCp

values, obtained from the slopes, are nearly identical (ΔCp with Mg2+

= −1.22 kcal/mole K; ΔCp

without Mg2+

= −1.15 kcal/mole K), indicating the ΔCp of binding is not magnesium dependent.

In both plots, error bars on the measured values are ≤0.6 kcal/mole and are smaller than the plot

symbols. Some data in Figure 3.8 were obtained by Kausiki Datta. This figure is reprinted from

reference 98 with permission.

66

We also assayed for the possible presence of a linked magnesium ion effect on the

measured binding enthalpy for the interaction of Klenow and primed template DNA. Figure 3.8B

shows temperature dependent measurements of the ΔHcal for binding of Klenow to 63/70-mer

DNA in the presence and absence of MgCl2. Klenow is known to require Mg2+

for catalytic

activity but not for DNA binding activity (99). The data in Figure 3.8B indicate that added

magnesium does not significantly alter the ΔCp of binding for Klenow.

Figure 3.9: Mean ΔCp values (kcal/mol K) for the binding of Klenow (A) and Klentaq (B) to

different DNA structures. The results for the longer DNA constructs are shown, and are the

averages and standard deviations of all ΔCp values obtained for each type of DNA with each

polymerase, including ΔCp data from references 97 and 98.

67

Table 3.15: ΔCp values (cal / mol K) of Klenow and Klentaq binding to different structures with

different lengths of DNA, including ΔCp data from references 97 and 98.

Single-Stranded DNA Primer-Template DNA Double-Stranded DNA

20-mer 63-mer 13/20-mer 63/70-mer 20/20-mer 63/63-mer

Klenow -487 ± 53a

-430 ± 7a

-890 ± 28c

-1094 ± 179c

-329 ± 51a

-440 ± 2a

Klentaq -75 ± 22a

-400 ± 71b

-531 ± 129a

-821 ± 1c

-760 ± 107c

-993 ± 152b

aCalorimetric ΔCp values.

bvan’t Hoff ΔCp values.

cThe average of van’t Hoff and calorimetric ΔCp values.

The ∆Cp values for binding the different DNA structures are summarized for both

polymerases in Figure 3.9 and Table 3.15. Cp has often been correlated at the molecular level

with changes in the accessible surface ( ASA) area upon complex formation, and the balance of

polar and non-polar surface within that ASA. In addition, large negative Cp values have often

been associated with site-specific DNA binding. We have shown that neither of these

correlations hold quantitatively for Klenow and Klentaq DNA binding (97, 98). We and others

have strongly suggested that these correlations may hold for some subset of protein-DNA

interactions, but that neither correlation holds quantitatively for DNA-protein interactions in

general (122, 123). As a general qualitative correlation in protein-DNA interactions, ∆Cp is

definitely reflective of ∆ASAnonpolar, but exact quantitative correlation does not universally hold

for any current model for this specific class of interactions (122, 123).

Lack of a universal precise quantitative correlation between ΔCp and molecular properties

for protein-DNA interactions does not mean, however, that such values are devoid of

information. ΔCp changes in protein-DNA interactions are still strong reflections of changes in

the qualitative nature of the molecular interaction and the types of non-covalent forces that

dominate the binding (100). The different ΔCp patterns in Figure 3.9 relative to the binding of the

same DNAs by Klenow and Klentaq indicate that the different affinities (ΔG values for each type

of DNA) are not simply changes in strength of association in the same binding mode, but

68

actually reflect different binding modes. Figure 3.9 shows the mean ΔCp values for all

determinations (calorimetric and Gibb’s Helmholtz) for both polymerases for the longer DNA

constructs. Shorter DNA constructs yield the same patterns: Klenow shows a large ΔCp of pt-

DNA binding and significantly smaller ΔCp’s of binding to ss- and ds-DNA, while Klentaq

shows large ΔCp values for binding both pt- and ds-DNA. (ΔCp values are in Table 3.15).

Klenow’s ability to bind DNA in both polymerase and editing modes does not explain these

thermodynamic patterns, as the primary ΔCp difference between Klenow and Klentaq is

manifested upon binding to blunt-ended ds-DNA.

3.3.6 The Length of DNA Effect on DNA and Polymerase Binding

Both Klenow and Klentaq bind longer ss-DNA tighter than shorter ss-DNA. The ΔΔG

binding values between the shorter and longer ss-DNAs are 0.5 kcal/mol for Klenow (Table 3.3)

and 0.7 kcal/mol for Klentaq (Table 3.4). The longer ss-DNA (63-mer) has some secondary

structure in solution at 25°C (see Section 3.3.5), and both polymerases may bind to those

secondary structures tighter than the completely single-stranded structure of the shorter 13-mer.

If a ΔG of -11.3 kcal/mol corresponds to a ∆Cp value of -0.49 kcal/mol K for shorter ss-DNA/KF

binding, a difference in ΔG of 0.5-0.7 kcal/mol might correspond to an additional ∆Cp value of 0.

Thus, the ∆Cp values of both shorter and longer ss-DNAs binding are similar (Table 3.10).

Both Klenow and Klentaq bind longer pt- and ds-DNA tighter than shorter pt- and ds-

DNA (Tables 3.3 and 3.4). The ΔΔG values between the shorter and longer pt-DNA binding by

Klenow and Klentaq are 0.6 kcal/mol (Table 3.3) and 0.1 kcal/mol (Table 3.4), respectively

while the ΔΔG values between the shorter and longer ds-DNA binding by Klenow and Klentaq

are 0.9 kcal/mol (Table 3.3) and 0.3 kcal/mol (Table 3.4), respectively. These data suggest that

the length of pt- and ds-DNA affects the binding ability of Klenow more significantly. Klenow

binds DNA with higher ion release than Klentaq; hence, Klenow binds DNA with larger

69

footprint than Klentaq. Therefore, Klenow may need longer DNA for proper DNA binding.

Because longer pt- and ds-DNA may give Klenow enough duplex DNA regions for Klenow to

bind DNA more properly, Klenow binds the longer pt- and ds-DNA tighter than the shorter pt-

and ds-DNA. However, the different lengths of pt- and ds-DNA binding by Klenow and Klentaq

do not alter the ∆Cp values (Figures 3.6 and 3.7), within error.

3.3.7 The Magnesium Chloride Effect on DNA and Polymerase Binding

Table 3.16: The binding constants (Kd) and the Gibbs free energy (∆G) of the binding of Klenow

and Klentaq polymerases to primer-template DNA (hp-39) and double-stranded DNA (hp-32 and

hp-46) in the absence and presence of MgCl2.All titrations were performed at 25°C in 10 mM

Tris and 5 mM MgCl2 at pH 7.9.

DNA

Klenow Klentaq

Kd

(nM)

ΔG

(kcal/mol)

Kd

(nM)

ΔG

(kcal/mol)

hp-32 without MgCl2 2.8 ± 0.4 -11.7 ± 0.01 4.3 ± 0.1 -11.4 ± 0.01

hp-32 with MgCl2 25.4 ± 8.7 -10.4 ± 0.2 3.2 ± 0.2 -11.6 ± 0.04

hp-39 without MgCl2 1.9 ± 0.3 -11.9 ± 0.1 4.3 ± 0.1 -11.4 ± 0.02

hp-39 with MgCl2 2.9 ± 0.3 -11.7 ± 0.1 1.7 ± 0.3 -12.0 ± 0.1

hp-46 without MgCl2 2.3 ± 0.4 -11.8 ± 0.1 5.6 ± 0.5 -11.3 ± 0.1

hp-46 with MgCl2 20.8 ± 5.4 -10.5 ± 0.1 6.8 ± 0.4 -11.2 ± 0.03

Magnesium ions are required for both Klenow and Klentaq enzymatic function (35, 75,

78, 99, 177-179). For DNA binding to pt-DNA, while Mg2+

appears not to be absolutely

essential, it does clearly enhance binding affinity. Klenow especially appears to require a

minimal amount of free Mg2+

for high affinity binding (99). Figure 3.10 shows that magnesium

ions help Klenow differentiate primer-template (pt-DNA) from blunt-end (ds-DNA) (see Table

3.16 for Kd values). Klenow binds pt-DNA with similar affinities in the absence and presence of

MgCl2, but the affinity of Klenow for ds-DNA is decreased by almost an order of magnitude (1.2

to 1.3 kcal/mol) when magnesium is present. On the other hand, Klentaq binds ds-DNA with

almost identical affinities in the absence and presence of MgCl2, and binds pt-DNA more tightly

in the presence of Mg2+

. Adding Mg2+

to the solution thus appears to function as an “affinity

70

switch” for Klenow binding to ds-DNA and to a lesser extent for Klentaq binding to pt-DNA, but

the Mg2+

switch acts in opposite directions for the two polymerases. In contrast, added Mg2+

does not have a significant effect on the binding affinity of Klenow for pt-DNA or Klentaq for

ds-DNA.

Figure 3.10: The effect of Mg2+

on the free energy of pt-DNA and ds-DNA binding by Klenow

(A) and Klentaq (B). G is the binding free energy in the absence of MgCl2 minus the free

energy of binding in the presence of 5 mM MgCl2 ( Gabsence- Gpresence). Free energy of binding

in these experiments was measured via competition assays as described in Materials and

Methods. Mean and standard deviations of three determinations are shown.

71

3.3.8 The RRRY Motif and ss-DNA Binding by Klenow and Klentaq

Modak et al. have recently identified a sequence they call the RRRY motif, which is

conserved across the Pol I family, and appears to be involved in ss-DNA binding of the template

portion of a melted DNA duplex during proofreading (57). Located near the base of the “fingers”

subdomain, mutations in this 4-residue sequence in Klenow will reduce the 3'-exonuclease

activity by up to 29 fold (57). The finding that Klentaq binds ss-DNA even though it does not

have a proofreading site suggests that the RRRY site is binding capable in Klentaq, although its

purpose in Klentaq is unclear. The significantly weaker ss-DNA binding by Klentaq relative to

Klenow, and the release of fewer ions upon ss-DNA binding by Klentaq relative to Klenow, may

either be due to additional ss-DNA interactions with the proofreading active site in Klenow

which are absent in Klentaq, or may simply be a part of the overall reduction in binding affinity

for any/all DNA exhibited by Klentaq relative to Klenow. This raises the question if isolated ss-

DNA binds only to the RRRY motif even in Klenow and the melted duplex DNA binds to the

RRRY and exonuclease sites of Klenow.

3.4 Summary

What do these thermodynamics say about the different binding modes of the two

proteins? For ss-DNA binding, both proteins bind ss-DNA with the lowest ion release and the

lowest ΔCp. Yet for both proteins, the position for ss-DNA binding in the binding hierarchy is

the most variable, switching from weakest to tightest binding as salt concentration increases for

both proteins. Although the switch occurs near 200 mM salt for both proteins, within the

hypothetical physiological salt range ss-DNA binding is the tightest in the hierarchy for Klenow,

and weakest in the hierarchy for Klentaq. In summary, it is noteworthy: 1) that ss-DNA binds to

Klentaq at all; 2) that the linked ion release and ΔCp of ss-DNA binding to both proteins is very

similar, suggesting similar binding interfaces; and 3) that the position of ss-DNA binding in the

72

binding hierarchy for both proteins changes very similarly with increasing salt concentrations.

These findings suggest that the recently identified RRRY ss-DNA binding motif may be the

primary binding site for isolated ss-DNA in both polymerases. Klenow’s proofreading site

clearly binds ss-DNA that has been melted out of a bound duplex (14). Klentaq, however, has no

proofreading site, and is missing most or all of the residues that have been biochemically

associated with the proofreading site. If isolated ss-DNA were binding primarily to the

proofreading site in Klenow, one might expect a more striking difference in the thermodynamics

of ss-DNA binding to Klenow versus Klentaq.

For ds-DNA versus pt-DNA binding, the results also highlight some notable binding

characteristics. Klentaq binding to pt-DNA and ds-DNA appear similar, if not identical, by all

thermodynamic criteria: the same ΔCp, the same affinities, and the same ion release. If only

duplex constructs had been used, one might hypothesize that Klentaq avoided the pt-end of the

DNA and bound only to the ds-end, but the binding thermodynamics of Klentaq binding to

primer-template and blunt-end hairpin DNAs shows that Klentaq indeed binds to blunt-end DNA

and primer-template DNA almost identically.

Klenow, on the other hand, binds differently to ds-DNA and pt-DNA thermodynamically:

the ion release for binding the two structures differ by about an ion, their binding free energies

differ by slightly less than a kcal/mol, and their ΔCp changes are very different. From a purely

thermodynamic point of view, the data suggest that the binding of ds-DNA to Klenow is

structurally different from pt-DNA binding; i.e. that a unique ds-DNA binding mode exists for

Klenow’s interaction with blunt-end ds-DNA. Without further structure-based characterization,

however, this hypothesis cannot yet be confirmed.

73

CHAPTER 4

TWO MODES OF BLUNT-END DNA BINDING BY KLENOW DNA POLYMERASE

4.1 Introduction

Some DNA polymerases have both polymerase and editing functionality. Co-crystal

structural studies of DNA/polymerase complexes have identified the DNA binding topologies on

Pol A polymerases during the polymerization and editing reactions (43, 53-55, 66, 67, 180).

Because Klenow possesses both activities, Klenow polymerase from Escherichia coli DNA

Polymerase I has been used to study the interaction between polymerization and editing sites

using techniques such as time-resolved fluorescence anisotropy (58-60) and circular dichroism

(64). Time-resolved fluorescence anisotropy with an internally dansyl-labeled primer-template

DNA (pt-DNA) has been used to differentiate the polymerase and editing binding modes of

DNA/KF complexes in solution (58, 60). Matched and mismatched DNA were used to obtain the

partitioning equilibrium of the DNA between polymerase and editing site binding to Klenow (58,

59). When the dansyl fluorophore is attached to the C5 position of a thymine base that is located

seven bases away from the 3'-end of the primer strand, the dansyl fluorophore is suggested to be

exposed to the solvent when Klenow binds the DNA in polymerase mode (58, 59). On the other

hand, the fluorophore will be buried within the protein when Klenow binds DNA in editing mode

(58, 59). Bailey et al. have measured that the exposed fluorophore has a fast anisotropic decay

while the buried one has a slower decay (58, 59). The anisotropic decay rate was correlated with

the primer binding conformations to determine the fractional binding of polymerase and editing

sites. These studies suggest that matched pt-DNA binds primarily (~85%) to the polymerase site

while 4 mismatched pt-DNA exclusively binds to the editing site (58, 60).

The shuttling of the matched and mismatched primer strand of DNA between polymerase

and editing domains of Klenow and Klentaq polymerases has also been measured by following

74

the fluorescence and circular dichroism of inserted 2-aminopurine dimer probes (64). 2-

aminopurine (2-AP) is a fluorescent isomer of adenine, and the change in the spectroscopic

interactions of two adjacent 2-AP molecules directly reflects conformational changes in the

DNA. When the polymerase binds DNA in the editing mode, the DNA will be unwound and the

2-AP bases are unstacked, resulting in a blue shift of the 2-AP dimer CD signal. Using this

technique, Datta et al. also suggest that Klenow binds matched pt-DNA in a mixed polymerase

plus editing mode and that pt-DNA with three consecutive terminal mismatches binds in editing

mode (64).

The ionic composition of the solution also affects the modes of binding of polymerase to

DNA. Divalent metal cofactors such as Mg2+

, Mn2+

, or Zn2+

are required for both polymerase

and editing activities. Although crystallographic studies have shown that both Mg2+

and Zn2+

filled the metal ion binding sites of Klenow, the exact metal ions used in vivo have not been

identified (75). The presence of Mg2+

has been suggested to shift the binding of matched pt-DNA

to the editing mode of Klenow while the presence of 2 mM EDTA (i.e. the removal of divalent

metal ions) shifts the DNA binding to the polymerase mode of Klenow (64). On the other hand,

the presence or absence of divalent metal ions does not affect the DNA binding conformation

with Klentaq (64).

Although Klenow and Klentaq polymerases share highly conserved residues and motifs

(22, 26), these A family DNA polymerases show somewhat different substrate selectivity (see

Chapter 3). Thermodynamically, Klenow differentiates primer-template from blunt-end DNA

while Klentaq binds both DNAs similarly. Most specifically, the ΔCp values of pt-DNA/KF and

ds-DNA/KF are significantly different, but those of pt-DNA/KTQ and ds-DNA/KTQ are similar.

In this chapter, the primer-template and blunt-end DNA bindings by Klenow and Klentaq

polymerases were examined using the electrophoretic mobility shift assay (EMSA).

75

DNA/Klenow binding reveals two types of complexes while DNA/Klentaq binding shows only

one type of complex. ds-DNA binding by KF results in both types of complexes, but one of them

is transient. Are these complexes indicating the polymerase and editing modes of binding, 1:1

and 2:1 binding, or a unique ds-DNA/KF mode of binding? Analytical ultracentrifugation and

circular dichroism were also used to examine the potential origin of these complexes.

4.2 Materials and Methods

4.2.1 Materials

Oligo(deoxyribo)nucleotides were obtained from Integrated DNA Technologies

(Coralville, IA) (see Chapter 3 for DNA sample preparation). The DNA constructs used for

experiments are shown in Table 4.1. Klenow Fragment (KF) and Klentaq (KTQ) polymerases

were purified in our laboratory (refer to Chapter 3 and/or references 47, 99, 168, and 169).

Table 4.1: DNAs used for EMSA binding experiments.

Primer-Template DNA (pt-DNA)

13/20-mer 5’-TCGCAGCCGTCCA-3’

3’-AGCGTCGGCAGGTTCCCAAA-5’

63/70-mer 5’-TACGCAGCGTACATGCTCGTGACTGGGATAACCGTGCCGTTTGCCGACTTTCGCAGCCGTCCA-3’

3’-ATGCGTCGCATGTACGAGCACTGACCCTATTGGCACGGCAAACGGCTGAAAGCGTCGGCAGGTTCCCAAA-5’

hp-39 AAGGCTACCTGCATGA-3’

AGCCGATGGACGTACTACCCCCC-5’

hp-39 mis 3 AAGGCTACCTGCACAG-3’

AGCCGATGGACGTACTACCCCCC-5’

Blunt-End Double-Stranded DNA (ds-DNA)

20/20-mer 5’-TCGCAGCCGTCCAAGGGTTT-3’

3’-AGCGTCGGCAGGTTCCCAAA-5’

63/63-mer 5’-TACGCAGCGTACATGCTCGTGACTGGGATAACCGTGCCGTTTGCCGACTTTCGCAGCCGTCCA-3’

3’-ATGCGTCGCATGTACGAGCACTGACCCTATTGGCACGGCAAACGGCTGAAAGCGTCGGCAGGT-5’

hp-32 AAGGCTACCTGCATGA-3’

AGCCGATGGACGTACT-5’

hp-46 AAGGCTACCTGCATGATAATTGG-3’

AGCCGATGGACGTACTATTAACC-5’

76

4.2.2 Methods

4.2.2.1 Electrophoretic Mobility Shift Assay (EMSA)

All DNA constructs used in these experiments were unlabeled. Samples are 10 µL, each

with 4 µM DNA and 4 µM protein. The control only contains 4 µM DNA. The composition of

the binding buffer is 10 mM Tris, 5 mM MgCl2, 50 mM KCl, at pH 7.9, 25°C. Incubation times

were varied and are reported in the results for each experiment. After incubation, the samples

were loaded onto an 8% acrylamide (29:1 acrylamide:bis) (gel size: 8.0 cm x 6.5 cm) and

electrophoresed in 1X TBE buffer (89 mM Tris Borate, 2 mM EDTA, pH 8.3). The gel was run

at a constant voltage of 100 volts for 40 minutes at room temperature. Gels were stained with

SYBR Green (Molecular Probes Inc.) or ethidium bromide for 30 minutes. Gel images were

obtained in a Bio-Rad gel imager.

4.2.2.2 Analytical Ultracentrifugation

Analytical ultracentrifugation experiments were performed in a Beckman Optima XL-A

analytical ultracentrifuge. The double-sector cell with quartz windows and charcoal-filled Epon

centerpiece was loaded with 425 μl of buffer and 400 μl of sample solution. Sample solutions

contain 5 µM DNA and/or 5 µM protein. The absorbance of protein-DNA complexes was

monitored at 289 nm to determine the particle distribution within the cell. The absorbance of the

reference sector is subtracted from that of the sample sector for each measurement. All

sedimentation velocity runs were performed at 38,000 rpm in an An-60 Ti rotor for nine hours.

The absorbance scans were recorded with a 0.004 cm step size. Sedimentation coefficients were

measured at 20°C in 10 mM Tris, 5 mM MgCl2, 50 mM KCl, at pH 7.9. Svedberg constants were

determined from fits of the data using the program Svedberg (181, 182). All s values reported

herein have been converted to s20,w values using measured solvent densities and viscosities. The

partial specific volume of the protein in the experimental buffer at 20°C was calculated from the

77

amino acid sequence using the computer program SEDNTERP (183, 184) and measured using

Edelstein and Schachman’s method (185-188). The partial specific volume of the proteins was

0.73 mL/g while the partial specific volume of the DNA was assumed to be 0.52 mL/g (189).

The partial specific volumes of protein-DNA complexes were calculated based on the weighted

averages of the partial specific volumes of protein and DNA alone (190). The density and

viscosity of the experimental buffer solution were calculated using SEDNTERP as

1.001215g/mL and 0.010056 poise, respectively.

4.2.2.3 Circular Dichroism

Circular dichroism (CD) spectral changes were used to assay for protein and DNA

conformational changes upon binding (191, 192). Circular dichroism spectra were measured at

25°C using an AVIV Model 202 circular dichroism spectrophotometer. A dual compartment

mixing cuvette from Starna Cells was used to ensure that any small spectral changes would not

be due to sample to sample concentration errors. The spectra of protein and DNA were obtained

before and after mixing. 2 μΜ DNA was in one compartment while 2 μΜ protein was in the

other. Spectra were collected in 1 nm steps. We examined the changes in protein and DNA

structures in solution using CD spectra of protein-DNA complexes versus protein and DNA

alone. CD signals above 240 nm are mostly due to the DNA, and the 265-290 nm range is

specifically sensitive for B-form DNA (185). The spectral signals below 240 nm are mainly due

to the protein, primarily in the 206-228 nm range (185). Any lack of spectral changes in these

wavelength ranges would indicate the absence of conformational changes upon binding (193).

4.3 Results and Discussion

4.3.1 Klenow/DNA versus Klentaq/DNA Complexes

We have examined the interactions of Klenow and Klentaq polymerases with matched

primer-template DNA (pt-13/20 and hp-39), blunt-end DNA (ds-20/20 and hp-46), and

78

mismatched primer-template DNA (hp-39 mis 3) (see Table 4.1 for DNA constructs used) using

the electrophoretic mobility shift assay (EMSA). A primer-template DNA with three consecutive

mismatches at the 3'-end of the primer strand was used as an editing mode substrate. Such

constructs have previously been shown to bind essentially exclusively in the editing mode to

Klenow (58, 60). Each lane on the gel in Figure 4.1 contains 4 µM DNA and 4 µM protein. The

lowest bands correspond to DNA only. There are two kinds of polymerase:DNA complexes that

appear on the gel: complex S (slower band) and complex F (faster band). Because only DNA is

being stained, only complexes containing DNA are visible. When Klenow polymerase binds

matched primer-template DNA (Lanes 1 and 3) and mismatched DNA (Lane 5), complex F

forms. When Klenow polymerase binds double stranded DNA (Lanes 2 and 4), complex S is

observed. On the other hand, only complex S forms when Klentaq polymerase binds any of these

different DNA structures (Lanes 6-10). It should be noted that hp-46 + both proteins consistently

form a doublet. The doublet shows complex S band and a complex that is slower than complex S

band (Lanes 4 and 9). Some of the complexes may be dissociating, so free DNA is seen on the

gel. A shorter blunt-end hairpin DNA was also examined (hp-32 + Klenow), and also forms

complex S (data not shown).

There are several possible explanations for the results shown in Figure 4.1. This chapter

involves testing the different hypotheses: 1) Complex S is a 2:1 protein:DNA while complex F is

1:1 binding. To test this, sedimentation velocity runs were performed to determine whether the

sizes and shapes of these complexes are different. Circular dichroism experiments were also

conducted to determine if secondary structural changes are seen that correlate with the different

complexes formed on the gel. 2) Since Klentaq/DNA complexes are always in the

polymerization mode, it is possible that complex S might be protein/DNA complex in

polymerization mode and complex F could be the protein/DNA complex in editing mode. An

79

argument against this model is the fact that Klenow binds both matched and mismatched pt-DNA

in complex F form. To test this, low temperature, EDTA, and added ddNTPs were used to try to

capture matched pt-DNA/KF complexes in the complex S form. 3) If both models 1 and 2 are

ruled out, the ds-DNA/KF complex may be a unique, newly identified binding complex, specific

to blunt-end DNA/Klenow binding.

Figure 4.1: Klenow (KF) and Klentaq (KTQ) binding to different DNA structures after 10

minutes incubation time. Lane 1: pt-13/20 + KF; Lane 2: ds-20/20 + KF; Lane 3: hp-39 + KF;

Lane 4: hp-46 + KF; Lane 5: hp-39 mis 3 + KF; Lane 6: pt-13/20 + KTQ; Lane 7: ds-20/20 +

KTQ; Lane 8: hp-39 + KTQ; Lane 9: hp-46 + KTQ; and Lane 10: hp-39 mis 3 + KTQ.

Incubation buffer is 10 mM Tris, 5 mM MgCl2, 50 mM KCl, at pH 7.9, 25°C. pt is pt-13/20, ds is

ds-20/20, pt hp is hp-39, ds hp is hp-46, and pt mis is hp-39 mis 3. Complex S is the slower

moving complex while complex F is the faster moving complex.

80

4.3.2 The Sizes of DNA/KF and DNA/KTQ Complexes

To attempt to learn more about the potential identities of the different protein-DNA

complexes observed on the gels, we conducted analytical ultracentrifugation experiments. In

Figure 4.1, matched pt-DNA/KF is a faster complex while ds-DNA/KF is a slower complex. To

determine the size of the molecules (DNA, KF, and DNA/KF complexes) based on their

sedimentation coefficients, sedimentation velocity analyses were used. Sample solutions contain

5 µM DNA and/or 5 µM protein. The measured sedimentation coefficient is corrected to an s20,w

value, which represents the sedimentation coefficient of the macromolecule in water at 20°C.

Molecules with small molecular weights and/or elongated shapes will have a smaller

sedimentation coefficient values than molecules with large molecular weights and/or globular

shapes. Table 4.2 and Figure 4.2 show the relationships between s20,w values and molecular

weights for a set of protein standards (194).

Table 4.2: The measured s20,w values and the calculated molecular weights of the globular and

highly asymmetric proteins (194).

Globular Proteins Calculated Molecular Weight (Da) s20,w (S)

Cytochrome C 12,400 1.8

Carboanhydrase 34,500 3.2

Bovine Serum Albumin 66,000 4.4

Aldolase 149,000 7.35

Catalase 240,000 11.0

Thyroglobulin 630,000 19.2

Highly Asymmetric Proteins Calculated Molecular Weight (Da) s20,w (S)

Tropomyosin 74,000 2.55

Myosin 470,000 6.4

If complex F is 1:1 binding, complex S is 2:1 binding, and both complexes have globular

shapes, the sedimentation coefficient of complex F will be about 5-6 Svedbergs (S) while that of

complex S will be from 7-8 S. Isolated Klenow and Klentaq (4.6 S on Table 4.3; and 4.1 S on

Table 4.4) have similar sedimentation coefficients to isolated Bovine Serum Albumin (4.4 S on

81

Table 4.2) (Figure 4.2). Therefore, the isolated Klenow and Klentaq are monomers and have

globular shapes. The sedimentation coefficient of the isolated DNA (i.e. hp-32) is 2.3 S (Table

4.3). As discussed in detail below, when KF binds different DNA structures, the sedimentation

coefficients range about 5.4-5.8 S, which corresponds to about 70-80 kDa molecules (Table 4.3).

Figure 4.2: The correlation between the logarithm of the molecular weight of globular proteins

and the logarithm of their sedimentation coefficients (solid line). ●: globular proteins; □: highly

asymmetric proteins; Δ: isolated KF and DNA/KF complexes; and ◊: isolated KTQ and

DNA/KTQ complexes.

82

Figure 4.3 shows the c(s) distributions of pt-DNA/KF complex and ds-DNA/KF complex

from which the sedimentation coefficients of the complexes were determined. The complexes

have large peaks at 5.5 S and 5.7 S, respectively. These results suggest that these protein-DNA

complexes exist in 1:1 stoichiometry. If there were two proteins binding to one DNA, the

measured sedimentation coefficients and molecular weights should have values of about 7-8 S

and 140 kDa, respectively. For example, aldolase, a globular protein, has a molecular weight of

149 kDa and a sedimentation coefficient of 7.35 S (Table 4.2). A very small peak at 9 S can be

seen on the distribution plot of pt-13/20 + KF (Figure 4.3). This higher molecular weight

molecule, which potentially corresponds to a 2:1 complex, is < 3% of the total complexes formed

in that sample. This is in contrast to Millar and associates’ report of ~24% 2:1 complex in a

similar pt-DNA/KF analytical ultracentrifugation experiments (131). The small 2 S peak in pt-

13/20 + Klenow data corresponds to the unbound DNA (Figure 4.3).

Table 4.3: The measured and predicted s20,w values and the calculated molecular weights of the

DNA/KF complexes.

Sample Calculated Molecular Weight (Da) Measured s20,w (S)*

hp-32 9,858 2.3

Klenow 68,094 4.6

hp-32+KF 77,952 5.4

pt-13/20+KF 78,116 5.5

hp-39+KF 80,001 5.6

ds-20/20+KF 81,735 5.7

hp-46+KF 82,276 5.8

Sample Calculated Molecular Weight (Da) Predicted s20,w (S)

ds-20/20+2KF 149,829 7.8

hp-46+2KF 150,370 7.8

*All s20,w values are ± 0.1 S.

83

Figure 4.3: Continuous sedimentation (c(s)) distributions of pt-DNA/KF and ds-DNA/KF

complexes. The maximum distributions of 5.5 S and 5.7 S correspond to 1:1 stoichiometry.

Because the Klenow and Klentaq complexes fall on a good correlation line with isolated

DNA and proteins (Figures 4.4 and 4.6), it suggests that the s20,w values of these complexes are

mostly reflecting the size of the complexes and not any unusual shape. The s20,w values of an

elongated molecule will be smaller than the s20,w values of a globular molecule of equivalent

mass, because the elongated molecule will have a higher frictional coefficient. Figures 4.4 and

4.6 show that the sedimentation coefficient increases as the molecular weight increases for these

DNA/KF and DNA/KTQ complexes, indicating that there is no significant shape perturbation or

information involved in DNA/KF complex and DNA/KTQ complex experiments. The monomer

84

form of an isolated Klenow and the 1:1 DNA/KF complexes data are closer to the correlation

between the logarithm of the molecular weight of globular proteins and the logarithm of the their

sedimentation coefficients (solid line on Figure 4.2) than the potential dimer of Klenow and the

2:1 DNA/KF complexes (Figure 4.2).

Figure 4.4: The measured s20,w values of the DNA/KF complexes versus the calculated molecular

weights. Since the s20,w values of DNA/KF complexes are directly proportional to the molecular

weight, there is no significant shape information involved with these complexes.

4

4.5

5

5.5

6

6.5

65000 70000 75000 80000 85000

s 20

, w

(S

)

Molecular Weight (Da)

85

All DNA/KTQ complexes form complex S on the gel (Lanes 6-10 in Figure 4.1). If

complex S were a 2:1 form of KTQ:DNA complex, the sedimentation coefficient of complex S

would be about 7-8 S. Table 4.4 shows that the s20,w value of isolated Klentaq is 4.1 S while pt-

DNA/Klentaq and ds-DNA/Klentaq complexes have s20,w values of 5.7 S and 6.5 S, respectively.

Figure 4.5 shows the c(s) distributions of pt-DNA/Klentaq and ds-DNA/Klentaq complexes. The

sedimentation coefficients of these complexes are too low to be 2:1 complexes. Since pt-

DNA/KTQ and ds-DNA/KTQ complexes show no difference on the gel (Figure 4.1) and the

sedimentation coefficient of pt-13/20 + KTQ (5.7 S; Table 4.4) is close to the sedimentation

coefficient of pt-13/20 + KF (5.5 S; Table 4.3), it follows that both the S and F complexes on the

gel are 1:1 complexes. It is not known at this time why the s20,w of ds-DNA/KTQ is ~ 14% larger

than pt-DNA/KTQ. It should be noted that these results are complicated by the fact that we have

subsequently found a kinetic shift for the ds-DNA/KF complex in EMSA (see Section 4.3.5), but

these analytical ultracentrifugation results do show that the complexes for all DNA/KF and all

DNA/KTQ are exclusively or predominantly (>95%) 1:1 at 8 hours.

Table 4.4: The measured and predicted s20,w values and the calculated molecular weights of the

DNA/KTQ complexes.

Sample Calculated Molecular Weight (Da) Measured s20,w (S)

Klentaq 62,409 4.1 ± 0.1*

pt-13/20+KTQ 72,431 5.7 ± 0.1

ds-20/20+KTQ 76,049 6.5 ± 0.2

Sample Calculated Molecular Weight (Da) Predicted s20,w (S)

pt-13/20+2KTQ 134,840 7.3

ds-20/20+2KTQ 138,458 7.4

*From reference 129.

86

Figure 4.5: Continuous sedimentation (c(s)) distributions of pt-DNA/KTQ and ds-DNA/KTQ

complexes. The maximum distributions of 5.7 S and 6.5 S correspond to 1:1 stoichiometry.

87

Figure 4.6: The measured s20,w values of the DNA/KTQ complexes versus the calculated

molecular weights. Since the s20,w values of DNA/KTQ complexes are directly proportional to

the molecular weight, there is no significant shape information involved with these complexes.

4.3.3 Assaying for Secondary Structure Changes Upon Primer-Template and Blunt-End

DNA Binding by Klenow and Klentaq

To assay for potential conformational changes of the protein and DNA upon formation of

the S versus F complexes in solution, we measured the secondary structures of the proteins,

DNAs, and protein-DNA complexes using circular dichroism (CD). Since spectral changes are

likely to be small, a dual compartment mixing cell was used to ensure that any signal change

observed was caused by a conformational change of the molecules as opposed to the normal

statistical scatter present even in careful pipetting. Protein and DNA were placed in separate

sides of the dual compartment cuvette, and CD spectra were obtained before and after mixing.

The spectra were collected in 1 nm steps. The CD signals between 206-228 nm correspond to

3

4

5

6

7

60000 65000 70000 75000 80000

S2

0, w

(S

)

Molecular Weight (Da)

88

signal from the protein (Figure 4.7) while the CD signals between 265-290 nm primarily come

from the DNA (Figure 4.8).

Figure 4.7: Klentaq shows slightly larger secondary structure changes upon DNA binding than

Klenow. The spectral regions from 206 to 228 nm of the circular dichroism spectra of the

protein/DNA complexes before and after mixing at 25°C are shown. All CD signal range scales

(y-axes) are identical. These regions primarily correspond to signals from the protein. A. Klenow

polymerase binding to pt-13/20 (pt-DNA) before and after mixing. B. Klentaq polymerase

binding to pt-13/20 (pt-DNA) before and after mixing. C. Klenow polymerase binding to ds-

20/20 (ds-DNA) before and after mixing. D. Klentaq polymerase binding to ds-20/20 (ds-DNA)

before and after mixing.

89

Figure 4.8: DNA spectral changes are similar upon binding of Klenow and Klentaq to pt or ds-

DNA. The spectral regions from 265 to 290 nm of the circular dichroism spectra of the

protein/DNA complexes before and after mixing at 25°C are shown. All CD signal range scales

(y-axes) are identical. These regions primarily correspond to signals from the DNA. A. Klenow

polymerase binding to pt-13/20 (pt-DNA) before and after mixing. B. Klentaq polymerase

binding to pt-13/20 (pt-DNA) before and after mixing. C. Klenow polymerase binding to ds-

20/20 (ds-DNA) before and after mixing. D. Klentaq polymerase binding to ds-20/20 (ds-DNA)

before and after mixing.

90

Figure 4.9: Comparing spectra of polymerases bound to pt- vs. ds-DNA. The spectral regions

from 206 to 228 nm of the circular dichroism spectra of protein/pt-DNA and protein/ds-DNA

complexes at 25°C are shown. All CD signal range scales (y-axes) are identical. These regions

primarily correspond to signals from the protein. A. Klenow polymerase binding to pt-13/20 (pt-

DNA) and ds-20/20 (ds-DNA). B. Klentaq polymerase binding to pt-13/20 (pt-DNA) and ds-

20/20 (ds-DNA). C. Klenow polymerase binding to hp-39 (pt-DNA) and hp-46 (ds-DNA). D.

Klenow polymerase binding to hp-39 (pt-DNA) and hp-46 (ds-DNA). pt-13/20 + KF data on

Panel A of Figure 4.9 is a re-plot from pt-13/20 + KF after mixing data on Panel A of Figure 4.7

while ds-20/20 + KF data on Panel A of Figure 4.9 is a re-plot from ds-20/20 + KF after mixing

data on Panel C of Figure 4.7. pt-13/20 + KTQ data on Panel B of Figure 4.9 is a re-plot from pt-

13/20 + KTQ after mixing data on Panel B of Figure 4.7 while ds-20/20 + KTQ data on Panel B

of Figure 4.9 is a re-plot from ds-20/20 + KTQ after mixing data on Panel D of Figure 4.7.

91

Figure 4.10: Both polymerases show similar CD signals when binding to the different DNAs.

The spectral regions from 265 to 290 nm of the circular dichroism spectra of protein/pt-DNA and

protein/ds-DNA complexes at 25°C are shown. All CD signal range scales (y-axes) are identical.

These regions primarily correspond to signals from the DNA. A. Klenow polymerase binding to

pt-13/20 (pt-DNA) and ds-20/20 (ds-DNA). B. Klentaq polymerase binding to pt-13/20 (pt-

DNA) and ds-20/20 (ds-DNA). C. Klenow polymerase binding to hp-39 (pt-DNA) and hp-46

(ds-DNA). D. Klenow polymerase binding to hp-39 (pt-DNA) and hp-46 (ds-DNA). pt-13/20 +

KF data on Panel A of Figure 4.10 is a re-plot from pt-13/20 + KF after mixing data on Panel A

of Figure 4.8 while ds-20/20 + KF data on Panel A of Figure 4.10 is a re-plot from ds-20/20 +

KF after mixing data on Panel C of Figure 4.8. pt-13/20 + KTQ data on Panel B of Figure 4.10 is

a re-plot from pt-13/20 + KTQ after mixing data on Panel B of Figure 4.8 while ds-20/20 + KTQ

data on Panel B of Figure 4.10 is a re-plot from ds-20/20 + KTQ after mixing data on Panel D of

Figure 4.8.

92

Figure 4.11: The signal differences between ds-DNA and pt-DNA binding by the polymerases.

ΔCD Signal = CD Signal ds-DNA/protein - CD Signal pt-DNA/protein. The spectral regions

from 200 to 300 nm of circular dichroism spectra of protein-DNA complexes at 25°C are shown.

All CD signal range scales (y-axes) are identical. A. Klenow polymerase binding to hp-46 – hp-

39 and ds-20/20 – pt-13/20. B. Klentaq polymerase binding to hp-46 – hp-39 and ds-20/20 – pt-

13/20. C. Klenow and Klentaq polymerases binding to ds-20/20 – pt-13/20. D. Klenow and

Klentaq polymerases binding to hp-46 – hp-39.

93

Small protein secondary structure rearrangements are observed before and after mixing

both Klenow and Klentaq polymerases with pt-DNA and/or ds-DNA (Figure 4.7). Klenow forms

complex F with pt-DNA and complex S with ds-DNA while Klentaq forms complex S with both

pt-DNA and ds-DNA on the gel. Therefore, if the gel migration patterns reflected significant

conformational changes, one might expect the difference between the CD signals of pt-DNA/KF

versus ds-DNA/KF before and after mixing to be larger than the CD signal difference between

pt-DNA/KTQ versus ds-DNA/KTQ before and after mixing at 200-300 nm. However, the

spectral regions from 206 to 228 nm of Klentaq binding to both pt-DNA and ds-DNA (Panels B

and D in Figure 4.7) show larger conformational changes upon binding than that of Klenow

binding to either structure (Panels A and C in Figure 4.7). Similar magnitude CD changes were

reported previously for pt-DNA by Kausiki Datta (97, 98). Figure 4.8 shows that the DNA

spectral changes are similar upon binding of Klenow or Klentaq to pt-DNA or ds-DNA.

Experiments equivalent to Figures 4.7 and 4.8 were also performed using hp-39 (pt-DNA) and

hp-46 (ds-DNA) (data not shown). All these comparisons show that although Klenow clearly

distinguishes pt- from ds-DNA thermodynamically and by EMSA while Klentaq does not, these

differences are not reflected at the secondary structural level.

After mixing, the CD signals of pt-DNA/KF are similar to the signals of pt-DNA/KTQ,

and so are ds-DNA/KF and ds-DNA/KTQ (Figures 4.9 and 4.10). However, the CD signals of

pt-DNA/protein complexes are different from the CD signals of ds-DNA/protein complexes,

especially from the DNA signals (Figures 4.9 and 4.10). Although small spectral changes are

seen, the difference in DNA spectra of Klentaq binding to pt-DNA and ds-DNA after mixing

(Panels B and D in Figure 4.10) is larger than that of Klenow binding to these DNA after mixing

(Panels A and C in Figure 4.10).

94  

Figure 4.11 shows that the secondary structure difference of the ds-DNA/protein and pt-

DNA/protein complexes are the same for both Klenow and Klentaq (Panels C and D in Figure

4.11). Panels A and B in Figure 4.11 show that comparing complexes containing different DNA

(pt-13/20 and ds-20/20 vs. hp-39 and hp-46) easily detects conformational differences.

Therefore, ds-pt/KTQ is different from ds-pt/KF with different types of DNA (blunt-end or

hairpin at one of the ends), but ds-pt/protein are the same when the same type of DNA is

compared (i.e. hp-39 and hp-46). Although it is of interest that ds-DNA/protein vs. pt-

DNA/protein complexes can be distinguished this way, the results add no further information on

the differences between complexes S and F on the gel.

The analytical ultracentrifugation results suggest that the size and shape of Klenow

binding to different DNA structures are quite similar. Circular dichroism shows that secondary

structural changes upon complex formation are small for both proteins binding to different DNA

structures. Therefore, the differences in the effective pI of DNA/protein complexes as the DNA

shifts positions on the protein may be the cause of the different migration of DNA/protein

complexes in the electrophoretic mobility shift assay.

4.3.4 The Complex S Is 2:1 and Complex F Is 1:1 Hypothesis

A recent review has explored the potential functional significance of 2:1 DNA

polymerase:DNA complexes in vivo and in vitro (3). Millar et al. have examined potential

dimerization of Klenow using analytical ultracentrifugation and the electrophoretic mobility shift

assay (131). Their gel shift data suggested that two Klenow molecules bind to matched primer-

template DNA and one Klenow polymerase binds to mismatched primer-template DNA (131).

Some discussion of Millar and associates’ data is pertinent here. After observing the

matched DNA/KF and mismatched DNA/KF on their gels, Millar et al. used analytical

ultracentrifugation to determine the sizes of those complexes. The sedimentation velocity and

95

equilibrium experiments performed by Millar et al. yield mostly 1:1 complexes for both matched

and mismatched pt-DNA binding by Klenow (3 µM DNA + 4.8 µM KF) (131). Using

equilibrium analytical ultracentrifugation, Millar and co-workers observed about 24% higher

molecular weight molecules in the 3 µM matched pt-DNA + 4.8 µM KF samples (131). Their

AU results show that the 2:1 (protein:DNA) complexes were favored when a higher

concentration of KF (3 µM pt-DNA + 10.5 µM KF) was used (131). Even Millar and associates

acknowledge quantitative inconsistencies in their own analytical ultracentrifugation and the

electrophoretic mobility shift assay data (131).

With the dimerization of Klenow interpretation, Millar et al. also suggested that the

second KF binding site may be located at the duplex part of DNA (131). This was because when

the single-stranded template region of DNA was shortened, they still observed the “2:1” complex

(131). However, when Millar and associates used longer matched and mismatched pt-DNA, both

“1:1” and “2:1” complexes were observed for both DNA/KF complexes (131). That result

contradicts their 2:1 complex interpretation, which suggests that the shorter matched pt-DNA

binding by Klenow only forms 2:1 complex. A shorter duplex part of DNA should have been

used in order to show that the stoichiometry becomes 1:1 when the second KF binding site is not

available.

Millar et al.’s and our data have some similarities and differences. Similar to their data,

1:1 complexes are mostly observed with lower concentrations of matched pt-DNA/KF in AU

(Table 4.3). However, some of our observations contradict Millar et al.’s data: 1) Both matched

and mismatched pt-DNA binding by Klenow yield faster moving complexes on our gels (Figure

4.1). 2) Our analytical ultracentrifugation data show < 3% 2:1 complexes in 5 µM matched pt-

DNA + 5 µM KF (Figure 4.3), compared to the 24% reported by Millar and associates.

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The von Hippel lab has also addressed some of these issues. I will summarize them here.

von Hippel and co-workers have observed similar results for matched and mismatched pt-

DNA/KF to our lab using gel shift when DNA and protein are in equimolar concentrations of ≤ 5

µM. The DNA constructs used in von Hippel and associates’ experiments are in Table 4.5. Faster

moving complexes are formed when both matched and mismatched pt-DNA bind to KF (3 µM

DNA + 2.6 µM KF) (Figure 4.12). Both slower and faster moving complexes form when 10.8

µM KF binds to 3 µM matched pt-DNA while mostly faster moving complexes form when 10.8

µM KF binds to 3 µM 3-4 mismatched pt-DNA (Figure 4.12), so slower moving complexes are

formed when higher concentration/stoichiometry of KF is used.

Isolated Klenow in the 0.8-11.8 µM concentration range always shows a slower moving

band (Figure 4.13). This result is similar to findings by our lab (Figure 4.19). Lanes 9 and 10 on

Figure 4.19 are isolated Klenow and Klentaq polymerases, respectively. The native proteins

migrate similarly to complex S (Figure 4.19). On the gel, pt-DNA/KF complex moves faster than

the isolated Klenow.

In summary, the 2:1 binding interpretation contrasts with our stoichiometric data from

fluorescence anisotropy and isothermal titration calorimetry that show 1:1 binding of Klenow

and Klentaq to pt- and ds-DNA (Chapter 3). The faster complex of pt-DNA/KF is 1:1 complex,

and at high stoichiometry, the slower complex of pt-DNA/KF may be the 2:1 complex for pt-

DNA/KF (Figure 4.12). However, the slower complex of all DNA/KTQ is definitely not 2:1

(Figure 4.1). Since pt-DNA/KTQ and ds-DNA/KTQ complexes look similar on the gel (Figure

4.1) and the sedimentation coefficient of pt-13/20 + KTQ (5.7 S; Table 4.4) is close to the

sedimentation coefficient of pt-13/20 + KF (5.5 S; Table 4.3), the DNA/KTQ complexes on the

gel are suggested to be 1:1. The slower band of Klenow only (Figure 4.13) is not a dimer because

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Klenow is a monomer according to both Millar et al. and our analytical ultracentrifugation

experiments (4.6 S in Table 4.3, and references 129 and 131).

Table 4.5: Matched and mismatched DNAs used for EMSA binding experiments by von Hippel

and associates. The mismatch bases are in italics. The AA shows where 2-aminopurine (2-AP)

dimers are inserted when used.

DNA Sequence

Matched 5’-GCAAGAACCGAACCAA-3’

3’-CGTTCTTGGCTTGGTTAAACTAATC-5’

Two

Mismatches

5’-GCAAGAACCGAACCAA-3’

3’-CGTTCTTGGCTTGGACAAACTAATC-5’

Three

Mismatches

5’-GCAAGAACCGAACCAA-3’

3’-CGTTCTTGGCTTGCACAAACTAATC-5’

Four

Mismatches

5’-GCAAGAACCGAACCAA-3’

3’-CGTTCTTGGCTTTCACAAACTAATC-5’

Figure 4.12: Matched and mismatched DNA binding by Klenow (KF). Lanes 4-10 are 3 µM

DNA + 2.6 µM KF while lanes 14-20 are 3 µM DNA + 10.8 µM KF. Lane 1: matched DNA (0

mm) only; Lane 2: three mismatches DNA (3 mm) only; Lane 3: KF only; Lane 4: matched

DNA + KF; Lane 5: two mm (2-AP labeled) + KF; Lane 6: two mm (unlabeled) + KF; Lane 7:

three mm (2-AP labeled) + KF; Lane 8: three mm (unlabeled) + KF; Lane 9: four mm (2-AP

labeled) + KF; and Lane 10: four mm (unlabeled) + KF. These gels were stained using SYBR

Green and were obtained by Kausiki Datta.

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Figure 4.13: Migrations of isolated Klenow (KF) on a native gel. Lanes 2-10 are KF alone with different concentrations. Lane 1: three mm + 11.8 µM KF; Lane 2: 0.8 µM KF; Lane 3: 1.6 µM KF; Lane 4: 2.6 µM KF; Lane 5: 3.3 µM KF; Lane 6: 4 µM KF; Lane 7: 5 µM KF; Lane 8: 6.7 µM KF; Lane 9: 8.2 µM KF; and Lane 10: 11.8 µM KF. This gel was stained using Coomassie and was obtained by Kausiki Datta.

4.3.5 Complex S for ds-DNA/KF Is a Transient Species

The DNA/protein complexes originally observed on Figure 4.1 have also been examined

after 8 hours incubation. After 8 hours, all DNA/KF complexes are in complex F form while all

DNA/KTQ complexes remain in complex S on the gel (Figure 4.14). The analytical

ultracentrifugation experiments show that all DNA/KF and all DNA/KTQ complexes are 1:1 at 8

hours (Tables 4.3 and 4.4). Figure 4.15 shows the binding of KF to ds-DNA as a function of

incubation time, and demonstrates that complex S formed by Klenow and ds-DNA is a transient

species that slowly converts to complex F.

Figure 4.16 shows the kinetic plots of ds-DNA binding by Klenow. The quantification of

complex F formation can be presented in several ways e.g. the density of complex F for each

lane, the normalized complex F by the total complex formation (complexes S and F) for each

lane, and the normalized complex F by the total density (complex S, complex F, and free DNA)

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for each lane. The different ways of quantification include a background subtraction step. Figure

4.16 shows the density of complex F for each lane. Complex S shifts to complex F over time

with a t1/2 of ~2-4 hours (Figure 4.16). However, the quantification of SYBR Green or ethidium

bromide stain may be tricky and less reliable because different complexes may be stained

differently. The fact that the free DNA decreases as the incubation time increases (Figure 4.15)

also adds to the difficulty of this quantification method.

Figure 4.14: Klenow (KF) and Klentaq (KTQ) binding to different DNA structures after 8 hours

incubation time. Lane 1: pt-13/20 + KF; Lane 2: ds-20/20 + KF; Lane 3: hp-39 + KF; Lane 4:

hp-46 + KF; Lane 5: hp-39 mis 3 + KF; Lane 6: pt-13/20 + KTQ; Lane 7: ds-20/20 + KTQ; Lane

8: hp-39 + KTQ; Lane 9: hp-46 + KTQ; and Lane 10: hp-39 mis 3 + KTQ. Incubation buffer is

10 mM Tris, 5 mM MgCl2, 50 mM KCl, at pH 7.9, 25°C. pt is pt-13/20, ds is ds-20/20, pt hp is

hp-39, ds hp is hp-46, and pt mis is hp-39 mis 3. The slower moving complex is labeled complex

S while the faster moving complex is complex F.

100

Figure 4.15: Klenow (KF) binding to ds-DNA as a function of time. Lane 1: ds-20/20 only.

Lanes 2-10 are ds-20/20 + KF at the follow incubation times: Lane 2: 10 minutes; Lane 3: 1

hour; Lane 4: 2 hours; Lane 5: 3 hours; Lane 6: 4 hours; Lane 7: 5 hours; Lane 8: 6 hours; Lane

9: 7 hours; and Lane 10: 8 hours. Lane 11: hp-46 only. Lanes 12-20 contain hp-46 + KF at the

same incubation times as lanes 2-10, respectively. Incubation buffer is 10 mM Tris, 5 mM

MgCl2, 50 mM KCl, at pH 7.9, 25°C.

101

Figure 4.16: The kinetic plots of ds-DNA binding to KF. A. The formation of complex F in ds-

20/20 + KF gel (the left gel in Figure 4.15). B. The formation of complex F in hp-46 + KF gel

(the right gel in Figure 4.15).

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4.3.6 Potential Effects of the Kinetic Shift for KF Binding to ds-DNA on Equilibrium Titrations

In Chapter 3, using fluorescence anisotropy, it was shown that ds-DNA binding is

thermodynamically different from pt-DNA binding for KF. This clearly correlates with the

differences in complex formation for KF with pt- versus ds-DNA found in Figure 4.1. However,

the question of how the kinetic shift for ds-DNA/KF (Figure 4.15) impacts the fluorescence

anisotropy results becomes important. It takes up to three hours to complete a fluorescence

anisotropy or an isothermal titration calorimetry experiment (15-20 points are collected in 8

minute intervals for fluorescence anisotropy). The electrophoretic mobility shift assay results

indicate that at 3 hours, ds-DNA/KF complexes are likely to exist in a mix of the two

conformations (complex S and F). However, variations in the duration of the fluorescence

anisotropy experiments (from 1-3 hours) do not affect the dissociation constant values (data not

shown). The mean time for any point in the titration is 1.5 hours, and titrations are performed so

that later additions contain more protein than early additions. Taken together, the kinetic

observations indicate that the thermodynamic data for ds-DNA/KF should mostly reflect

complex S with, at most, up to 40% complex F. This assumes that the kinetic shift for ds-

DNA/KF seen on the gels occurs at the same rate for the 10-100 times lower reactant

concentrations in fluorescence anisotropy relative to the stoichiometric condition in the

electrophoretic mobility shift assay.

4.3.7 Is the ds-DNA/KF Kinetic Shift Due to an Exonuclease Activity?

Is it possible that Klenow is degrading the DNA to cause the shift from complex S to F

for ds-DNA? We have used Klenow exo- to prevent the degradation of the primer strand in all of

these experiments (45). Moreover, complex S always migrates to a discrete complex F. If there

were residual exonuclease activity, one might expect a continuous degradation of complex S into

a smear of smaller complexes.

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Fluorescence anisotropy experiments also suggest that Klenow may not degrade the DNA

because the anisotropy values, measured from the emission of rhodamine-X on the DNA, do not

decrease after Klenow is added, as is observed if native Klenow or other proofreading active

polymerases are used in the fluorescence anisotropy assay (195).

4.3.8 Is the ds-DNA/KF Kinetic Shift Due to a Shift Between the Polymerization and Editing Modes of Binding?

As noted earlier in this chapter, previous studies have attempted to measure the

partitioning between the polymerization and editing modes of binding in Klenow (58-60, 64,

196), although different groups report different relative partitioning of some DNAs (especially

matched pt-DNA). Despite this, some results have been consistent among laboratories. For

example, primer-template DNA with three - four consecutive mismatches at the junction has

been shown to bind exclusively in the editing mode to Klenow using time-resolved fluorescence

anisotropy and circular dichroism of inserted 2-AP dimer probes  (58, 60). Our gels show that

only complex F forms when Klenow binds DNA with 3 mismatches (Figures 4.1 and 4.14).

Since Klentaq-DNA complexes are always in polymerase mode, it is possible that

complex S might be protein-DNA complex in polymerization mode and complex F could be the

protein-DNA complex in editing mode. This interpretation suggests that Klenow initially binds

primer-template DNA in editing mode and double-stranded DNA in polymerization mode, and

that Klenow/ds-DNA complexes then slowly switch to the editing mode.

Figure 4.15 shows that the ds-DNA/KF complex shifts from complex S to F over time.

Klenow binding in the editing mode requires the melting of three or four base pairs of duplex

DNA (43, 65, 78). The requirement for melting of the duplex and the translocation of the DNA

from the polymerization domain to the 3' → 5' editing domain was suggested to be the rate

limiting step for the Klenow exonuclease reaction with duplex DNA (77, 79). It may be that

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Klenow melts pt-DNA (complex F) faster than ds-DNA (complex S) because pt-DNA has the

single-stranded portion.

The recent 2-AP dimer spectroscopic study by von Hippel and associates suggests that

Klenow binds DNA in polymerase mode in the absence of magnesium ion (in the presence of

EDTA) but binds DNA in the editing mode in the presence of magnesium ion (64). Figure 4.17

shows that complex S forms earlier in the titration without magnesium ions than the one with

magnesium ions. Therefore, Klenow binds pt-DNA (hp-39) tighter in buffer without magnesium

ions than in buffer with magnesium ions. Similar result has been observed using fluorescence

anisotropy. Figure 4.17 partially supports the model suggested by von Hippel and associates

because complex F forms at the same point in the titration with and without magnesium ion

while complex S forms earlier without Mg2+

.

Figure 4.17: hp-39 binding by Klenow polymerase without (A) and with (B) an additional 5 mM

MgCl2. The DNA is labeled with -32

P-ATP at the 5'-end using T4 polynucleotide kinase. [DNA]

in each lane is 5 nM. Klenow concentrations in lanes 1-15 are: 0, 25 nM, 50 nM, 100 nM, 150

nM, 200 nM, 250 nM, 300 nM, 450 nM, 500 nM, 600 nM, 700 nM, 800 nM, 900 nM, and 1000

nM. Incubation was performed at 25°C in 10 mM Tris, 5 mM MgCl2, and 50 mM KCl at pH 7.9.

Gel was run at 4°C. These gel shift data were obtained using a Storm PhosphorImager by

Sreerupa Ray.

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4.3.9 Attempts to Capture pt-DNA/KF in Complex S

Multiple studies show that the complexes formed when matched pt-DNA binds Klenow

should be at least 50% polymerization mode complexes (58-62, 64). A major caveat in the

polymerase and editing model introduced in Section 4.3.8 is that pt-DNA/KF at 1:1 protein:DNA

ratio has not been observed in complex S although complex S of pt-DNA/KF is clearly observed

at high protein:DNA ratios (Figures 4.12 and 4.17). Since pt-DNA/KF complex has not been

seen in complex S form, we cannot confidently assign complex S as the polymerization mode

and complex F as the editing mode. Therefore, low temperature, EDTA, and added ddNTP have

been used to try to capture pt-DNA/KF complex in complex S form.

Low temperature samples were incubated at 0°C. Figure 4.18 shows that lower

temperature slows down the shift of ds-DNA/KF complex such that the transition seen in Figure

4.15 has not even begun after 12 hours at 0°C in Figure 4.18 (Lanes 8-12). However, the pt-

DNA/KF complex is still always in complex F (see Lanes 2-6 on Figure 4.18).

Figure 4.18: The low temperature effect on Klenow (KF) binding to pt-DNA (Lanes 1-6) and ds-

DNA (Lanes 7-12). All samples were incubated at 0°C. Lane 1: hp-39; Lane 2: hp-39 + KF (10

min. incubation); Lane 3: hp-39 + KF (1 hour incubation); Lane 4: hp-39 + KF (4 hours

incubation); Lane 5: hp-39 + KF (8 hours incubation); Lane 6: hp-39 + KF (12 hours

incubation); Lane 7: hp-46; Lane 8: hp-46 + KF (10 min. incubation); Lane 9: hp-46 + KF (1

hour incubation); Lane 10: hp-46 + KF (4 hours incubation); Lane 11: hp-46 + KF (8 hours

incubation); and Lane 12: hp-46 + KF (12 hours incubation). Incubation buffer is 10 mM Tris, 5

mM MgCl2, 50 mM KCl, at pH 7.9, 25°C.

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Using the 2-AP dimer spectroscopic assay, Datta et al. have suggested that Klenow binds

DNA in polymerase mode in 2 mM EDTA buffer (64). 2 mM EDTA also appears to slow down

the shift from complex S to F for ds-DNA binding by Klenow (see Lanes 6 and 8 on Figure

4.19). However, like temperature, EDTA does not capture pt-DNA/KF complex in complex S

(Lanes 2 and 4 on Figure 4.19).

Figure 4.19: Klenow binding to pt- and ds-DNA in Mg2+

and EDTA buffers after 10 minutes and

3 hours incubation time. Mg2+

buffer contains 10 mM Tris, 5 mM MgCl2, and 50 mM KCl while

EDTA buffer is 10 mM Tris and 2 mM EDTA at 25°C, pH 7.9. Lane 1: pt-13/20 + KF in Mg2+

buffer (10 min.); Lane 2: pt-13/20 + KF in EDTA buffer (10 min.); Lane 3: pt-13/20 + KF in

Mg2+

buffer (3 hrs.); Lane 4: pt-13/20 + KF in EDTA buffer (3 hrs.); Lane 5: ds-20/20 + KF in

Mg2+

buffer (10 min.); Lane 6: ds-20/20 + KF in EDTA buffer (10 min.); Lane 7: ds-20/20 + KF

in Mg2+

buffer (3 hrs.); Lane 8: ds-20/20 + KF in EDTA buffer (3 hrs.); Lane 9: KF only; and

Lane 10: KTQ only.

107

Using their spectroscopic assay, Datta et al. have also suggested that Klenow binds

matched primer-template DNA in polymerase mode when calcium (Ca2+

) is substituted for Mg2+

in the binding buffer (64). If EDTA, Ca2+

, and the absence of divalent metal ions enhance the

binding of Klenow to matched pt-DNA in the polymerase site, one might expect to trap some pt-

DNA/KF complexes in complex S under those conditions. Lanes 3, 5, and 7 in Figure 4.20 show

that pt-DNA/KF complexes only form complex F under those conditions. Therefore, complex S

may not be DNA/KF complexes in polymerase mode. Figure 4.20 may suggest that complex F

contains both polymerase and editing modes of matched pt-DNA binding by Klenow.

Figure 4.20: Klenow binding to pt- and ds-DNA in EDTA, Ca2+

, no divalent metal (no Me2+

),

and Mg2+

buffers at 10 minutes incubation. EDTA buffer is 10 mM Tris, 2 mM EDTA, and 50

mM KCl; Ca2+

buffer is 10 mM Tris, 5 mM CaCl2, and 50 mM KCl; no divalent metal (no Me2+

)

buffer is 10 mM Tris and 50 mM KCl; and Mg2+

buffer contains 10 mM Tris, 5 mM MgCl2, and

50 mM KCl. The pH of all buffers is 7.9 at 25°C. Lane 1: pt-13/20 only; Lane 2: ds-20/20 only;

Lane 3: pt-13/20 + KF in EDTA buffer; Lane 4: ds-20/20 + KF in EDTA buffer; Lane 5: pt-

13/20 + KF in Ca2+

buffer; Lane 6: ds-20/20 + KF in Ca2+

buffer; Lane 7: pt-13/20 + KF in no

Me2+

buffer; Lane 8: ds-20/20 + KF in no Me2+

buffer; Lane 9: pt-13/20 + KF in Mg2+

buffer;

and Lane 10: ds-20/20 + KF in Mg2+

buffer. This gel is electrophoresed in 1X TB buffer (89 mM

Tris Borate, pH 8.3).

During polymerization, after formation of the protein:DNA binary complex, dNTP binds

to the protein/DNA complex so polymerization can occur. If ddNTP is used instead of dNTP, the

108

polymerase cannot add any additional nucleotides after the addition of that ddNTP because the 3'

-end of the primer terminus will not have a 3'-OH. We tested if added ddNTP could trap some pt-

DNA/KF in complex S.

ddNTP does not affect most of the shifts (Figures 4.21 and 4.22). Adding 500 µM ddNTP

does not cause any shifts at 10 minutes incubation because the complexes with and without the

addition of ddNTP appear to be similar (For KF, compare Lanes 1 and 7, Lanes 2 and 10, Lanes

3 and 8, and Lanes 5 and 9; For KTQ, compare Lanes 11 and 17, Lanes 12 and 20, Lanes 13 and

18, and Lanes 15 and 19 on Figure 4.21). Adding ddNTP after 8 hours incubation, at the

beginning or the end of incubation time, may slow hp-39/KF and hp-46/KF (Lanes 3 and 4 on

Figure 4.22). Incubating ddNTP with the complex in 10 mM Tris and 2 mM EDTA buffer does

not affect the shift (data not shown). Since the addition of ddNTP does not affect most of the

matched pt-DNA/KF, complex S may not be the DNA/KF complex in polymerase mode.

Figure 4.21: Klenow (KF) (lanes 1-10) and Klentaq (KTQ) (lanes 11-20) binding to different

DNA structures in the presence and absence of ddNTP after 10 minutes incubation time. Lane 1:

pt-13/20 + ddATP + KF; Lane 2: ds-20/20 +ddATP + KF; Lane 3: hp-39 + ddTTP + KF; Lane 4:

hp-46 + ddTTP + KF; Lane 5: pt-63/70 + ddATP + KF; Lane 6: ds-63/63 + ddATP + KF; Lane

7: pt-13/20 + KF; Lane 8: hp-39 + KF; Lane 9: pt-63/70 + KF; and Lane 10: ds-20/20 + KF.

Lanes 11-20 contain KTQ binding to the same DNA sample as lanes 1-10, respectively.

Incubation buffer is 10 mM Tris, 5 mM MgCl2, 50 mM KCl, at pH 7.9, 25°C. pt is pt-13/20, ds is

ds-20/20, pt hp is hp-39, ds hp is hp-46, pt L is pt-63/70, and ds L is ds-63/63. Slower moving

complex is labeled complex S while faster moving complex is complex F. It should be noted that

there is no difference in the complexes with and without ddNTP.

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Interestingly, Klenow binding to pt-63/70 produces both complex S and F (Lanes 5 and 9

on Figure 4.21) while ds-63/63 binding by Klenow only forms complex S (Lane 6 on Figure

4.21), suggesting that pt-63/70 + KF forms complexes in both polymerase and editing modes and

ds-63/63 + KF forms complex in the polymerase mode. On the other hand, Klentaq forms both

complex S and a slower band with pt-63/70 and ds-63/63 (Lanes 15, 16, and 19 on Figure 4.21).

It should be noted that hp-39 + KF sometimes form complex S (Lane 8 on Figure 4.21).

Figure 4.22: Klenow (KF) (lanes 1-4) and Klentaq (KTQ) (lanes 5-8) binding to different DNA

structures in the presence of ddNTP after 8 hours incubation time. Lane 1: pt-13/20 + ddATP +

KF; Lane 2: ds-20/20 +ddATP + KF; Lane 3: hp-39 + ddTTP + KF; Lane 4: hp-46 + ddTTP +

KF; Lane 5: pt-13/20 + ddATP + KTQ; Lane 6: ds-20/20 + ddATP + KTQ; Lane 7: hp-39 +

ddTTP + KTQ; and Lane 8: hp-46 + ddTTP + KTQ. Incubation buffer is 10 mM Tris, 5 mM

MgCl2, 50 mM KCl, at pH 7.9, 25°C. pt is pt-13/20, ds is ds-20/20, pt hp is hp-39, and ds hp is

hp-46.

110

4.4 Summary

The “large fragments” of Escherichia coli and Thermus aquaticus Type I DNA

polymerases, Klenow and Klentaq polymerases, have homologous structures. However, we have

shown that Klenow and Klentaq polymerases bind ds-DNA differently. In EMSA, all ds-

DNA/KF complexes show a time dependent shift from a slower to a faster moving complex

while matched and mismatched pt-DNA/KF complexes are found only in the fast moving

complex. DNA/KTQ complexes are observed in complex S only, and both isolated polymerases

(without DNA bound) co-migrate with complex S. Low temperature (0°C) and 2 mM EDTA

slow down the shift from complex S to F for ds-DNA/KF complexes. However, EDTA, Ca2+

,

and the absence of divalent metal ions do not capture the binding of Klenow to matched pt-DNA

in complex S form. Incubating ddNTP with the complex does not affect most of the shifts. These

different conditions do not capture pt-DNA/KF in complex S form.

Fluorescence anisotropy and isothermal titration calorimetry results reveal that Klenow

and Klentaq polymerases form 1:1 complexes with these DNAs. No significant oligomerization

of any complexes was observed. Circular dichroism finds no significant secondary structure

differences between pt- and ds-DNA binding by Klenow and Klentaq polymerases that correlate

with the gel shift results.

Klenow has been suggested to bind DNA more in the polymerization mode in the

absence of magnesium ions (or in the present of EDTA) and more in the editing mode in the

presence of magnesium ions (64). However, at this time, we cannot confidently assign the

different types of complexes on the gel as polymerase and editing modes of binding complexes.

Since pt-DNA/KF complex has not been observed in complex S form in the absence of

magnesium ions and the presence of EDTA, this result could mean that complex S and F are not

polymerase and editing modes of binding. Complex F may consist of both polymerase and

111

editing modes of Klenow binding while the transient complex S is some other mode of ds-

DNA/KF binding. On the other hand, all DNA/KTQ complexes are in the polymerase mode of

binding.

Similar to the polymerase and editing modes model, some data support the 2:1

oligomerization model while other data conflict. Equimolar pt-DNA and KF complexes yield 1:1

complexes (faster band). At high stoichiometry, pt-DNA/KF does form a slower band (Figure

4.12), which may be a 2:1 complex, however, the co-migrating slower DNA/KTQ complexes are

not 2:1 complexes. Analytical ultracentrifugation results show that all DNA/KF and DNA/KTQ

complexes are 1:1 after 8 hours, even while DNA/KTQ complexes remain as slower complexes

on the gel after 8 hours incubation. If complex S is the 2:1 complex, one might expect to chase

all complex F to be complex S on the gel with more protein. However, complex F cannot shift to

complex S completely (Figures 4.12 and 4.17). Further testing of the 2:1 binding hypothesis is

clearly needed.

In summary, all DNA/KTQ complexes are 1:1 and in polymerase mode. pt-DNA/KF

complex is 1:1 at equimolar concentrations of DNA:KF while ds-DNA/KF is a transient

complex. I have clearly demonstrated a new binding behavior for Klenow with ds-DNA which

correlates with thermodynamic data on Klenow and Klentaq DNA binding. Neither the

oligomerization model nor the polymerase and editing modes model is fully supported by any

current data from any laboratory. We might have to consider a new ds-DNA/KF binding mode.

Certainly, more experimentation exploring the oligomerization and the polymerase and editing

modes models is warranted.

112

CHAPTER 5

DISCUSSION OF MOLECULAR MODELS FOR THE INTERACTIONS BETWEEN

DNA POLYMERASE AND DIFFERENT DNA SUBSTRATES

5.1 Introduction

I have demonstrated that Klenow and Klentaq polymerases show similarities and

differences when they bind to different DNA structures in this dissertation (Chapters 3 and 4). I

will present molecular models of the interactions of Klenow and Klentaq polymerases with

single-stranded, primer-template, and blunt-end DNA in this chapter. Primer-template DNA has

been shown to be the substrate for DNA replication (17, 19) while single-stranded and blunt-end

DNA have been suggested to be substrates for DNA repair (83, 174, 175). Single-stranded DNA

has been shown to be the substrate for editing mode of binding in Klenow (83), and blunt-end

DNA has been suggested to be a substrate for non-homologous end-joining in Klenow (174,

175). In this chapter, I will discuss three potential models for pt-DNA versus ds-DNA binding by

Klenow and Klentaq.

5.2 ss-DNA Binding by Klenow and Klentaq

Single-stranded DNA (ss-DNA) has been suggested to be the substrate for editing mode

binding of DNA to Klenow (43, 75, 78, 82, 83). Both polymerases can bind single-stranded

DNA (Chapter 3), although Klentaq does not have any editing activity (33, 51). Because Klentaq

does not bind DNA in an editing mode, how is it binding ss-DNA? The recently identified

RRRY motif has been shown to be involved in the binding of the 5'-end of single-stranded DNA

to Klenow polymerase in the editing mode (57). The fact that Klentaq binds to single-stranded

DNA may be due to the RRRY motif which is also conserved in Klentaq.

The co-crystal structures of Klentaq binding to pt-DNA 12/13-mer and 12/14-mer

showed that the position of the 5'-end of the template strand is close to the RRRY motif in

Klentaq (53-56). Therefore, a longer 5'-template strand of pt-DNA or an isolated ss-DNA may

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bind to the RRRY motif. Thus in both Klenow and Klentaq, the RRRY site may help to stabilize

primer-template DNA binding during DNA replication by binding to the 5'-end of the template

strand of DNA, and this function may also result in ss-DNA binding activity for both

polymerases. Mutagenesis studies of the RRRY residues in Klentaq are needed to confirm the

hypothesis which suggests that ss-DNA binds to the RRRY motif.

Modak et al. have suggested that ss-DNA binds both the RRRY motif and the editing site

in Klenow and Figure 5.1A shows the schematic binding model proposed by Modak et al. (57). It

is known that Klenow exo- lacks editing activity, but it can still bind DNA substrate in the

editing mode (45, 81). Our hypothesis is that the 5'-end of ss-DNA binds to the RRRY motif in

Klentaq while the 3'-end of ss-DNA does not bind to the editing domain of Klentaq because

Klentaq does not have an editing active site (Figure 5.1B) (33, 51). In this proposed model, the

location of the 3'-end of ss-DNA in Klentaq/ss-DNA complex is not known. I also propose that

the binding of Klentaq polymerase to ss-DNA is weaker (Chapter 3) because Klentaq does not

have an editing binding site.

The fact that heat capacity correlates with ΔASA in many protein:DNA interactions

suggests that if the heat capacity changes for formation of two different protein:DNA complexes

are equal, these complexes could have similar binding interfaces. On the other hand, if the heat

capacity changes for formation of two different protein:DNA complexes are not equal, it is much

less likely that these complexes will have similar binding interfaces. The heat capacity values of

ss-DNA binding by Klenow and Klentaq are very similar (Figure 3.9), so these findings suggest

that the ss-DNA/Klenow and ss-DNA/Klentaq complexes may have similar structures. However,

the smaller ion release in ss-DNA/Klentaq complex (1 ion released) relative to ss-DNA/Klenow

complex (2 ions released) (Tables 3.7 and 3.8) indicates that Klentaq has a smaller footprint

when binding to ss-DNA which may be because Klentaq does not bind DNA in the editing

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domain. This observation correlates with our hypothesis that ss-DNA binds to both the RRRY

motif and the editing binding site in Klenow and only binds to the RRRY motif in Klentaq. Thus,

Klenow and Klentaq may have different structures when they bind to ss-DNA (Figure 5.1).

Figure 5.1: A proposed model for single-stranded DNA binding by Klenow (A) and Klentaq (B)

polymerases. Both polymerases can bind single-stranded DNA. Figure 5.1A is based on

reference 57 and shows the ss-DNA binding model proposed in that publication. I propose that

the binding of Klentaq polymerase to ss-DNA is weaker because Klentaq does not have an

editing binding site. Figure 5.1B proposes that the 5'-end of ss-DNA binds to the RRRY motif in

Klentaq. The exact location of the 3'-end of ss-DNA in Klentaq/ss-DNA complex is not yet

known.

5.3 Molecular Models to Explain pt-DNA versus ds-DNA Binding by Klenow and Klentaq

For primer-template versus blunt-end DNA binding, Klentaq binds these DNAs similarly,

e.g. similar binding affinities, similar ion releases, similar heat capacity changes (Chapter 3), and

similar migrations in the electrophoretic mobility shift assay (Chapter 4). Therefore, one might

115

expect that the primer-template DNA/Klentaq and blunt-end DNA/Klentaq complexes have

similar structures. The co-crystal structure of Klentaq binding to pt-DNA shows that Klentaq

binds pt-DNA in polymerase mode (53). Thus, the energetic results reported here suggest that the

structure of ds-DNA/Klentaq complex may be similar to the structure of pt-DNA/Klentaq

complex solved in reference 53.

On the other hand, similar logic implies that Klenow binds primer-template and blunt-end

DNA differently because the ∆Cp values of pt-DNA and ds-DNA binding by Klenow are very

different (Chapter 3), and also because these two complexes behave differently in the

electrophoretic mobility shift assay (Chapter 4). Therefore, one would expect that the primer-

template DNA/Klenow and blunt-end DNA/Klenow complexes have different structures.

Several potential molecular models for correlating the thermodynamics and the structural

data were introduced in Chapter 4: the oligomerization model (see Section 4.3.4), the polymerase

and editing modes model (see Section 4.3.8), and the unique ds-DNA/KF binding model (see

Section 4.3.1). Each model is described and discussed in further detail below.

5.3.1 The Oligomerization Model

First, I will discuss data that support the oligomerization model. In reference 131, Millar

et al. proposed the oligomerization model for pt-DNA/KF complexes and suggested that two

Klenow molecules might bind to the duplex region of DNA (131). The depiction of complexes S

and F for DNA/KF in Figure 5.2 are thus based on reference 131. Figure 4.12 suggests an

oligomerization model because pt-DNA/KF forms a slower band at high protein:DNA ratios

which may be a 2:1 protein:DNA complex (Figure 4.12). On the other hand, at equimolar ratios

of pt-DNA and KF, only a faster moving complex is formed, which may correspond to a 1:1

complex (Figure 4.12). This observation is similar to Millar et al.’s data (131). So, in this model,

complex S for DNA/KF is a 2:1 complex and complex F for DNA/KF is a 1:1 complex.

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Figure 5.2: Schematic showing the oligomerization model for DNA binding by Klenow and

Klentaq polymerases. All pt- and ds-DNA/KTQ complexes are 1:1 while complex S of DNA/KF

is shown as 2:1 binding and complex F of DNA/KF is shown as 1:1 binding. Complexes S and F

of DNA/KF here are based on similar schematic depictions of the potential KF 2:1 complex

proposed by Millar et al. and described in Table 3 of reference 131.

On the other hand, the oligomerization model is not supported by DNA/Klentaq binding

data. Figure 4.14 shows that all DNA/KTQ complexes remain as slower complexes (complex S)

and all DNA/KF complexes form faster complexes (complex F) on the gel after 8 hours

incubation. According to the oligomerization model, these data would suggest that all DNA/KTQ

117

complexes are 2:1 and all DNA/KF complexes are 1:1 after 8 hours. This interpretation

contradicts our analytical ultracentrifugation data which reveal that all DNA/KF and DNA/KTQ

complexes are 1:1 after 8 hours (Table 4.4). The analytical ultracentrifugation results indicate

that the co-migrating slower DNA/KTQ complexes are not 2:1 complexes. Therefore, complex S

of DNA/KTQ must be different from complex S of DNA/KF in this model. If that is true, that

also means that the 1:1 DNA/KF and DNA/KTQ complexes migrate differently. Complex F of

DNA/KTQ is not observed on the gel (Figure 4.14). Based on these observations, complex S of

DNA/KTQ is 1:1 binding (Figure 5.2).

The 1:1 complexes of DNA/KF and DNA/KTQ may migrate differently on the gel

because of a difference in their effective pI. The isolated proteins co-migrate on the gel because

the pIs of isolated proteins are similar: 5.70 for Klenow and 5.85 for Klentaq (Figures 4.13 and

4.19). If both polymerases bind DNA in the same way, the complexes of Klenow and Klentaq

binding to the same DNA would co-migrate on the gel. However, the pt-DNA/KF (complex F)

and pt-DNA/KTQ (complex S) migrate differently on the gel. This suggests that the topological

positions of the pt-DNA in the Klenow and Klentaq complexes are different.

Electrophoretic samples of ds-DNA/KF complexes on several gels (Figures 4.1, 4.14,

4.15) show a free DNA band eventhough the protein:DNA ratio is 1:1. If complex S for ds-

DNA/KF were a 2:1 complex, one might expect half the added DNA to remain in the free DNA

band. However, this free DNA could also be caused by the dissociation of the complex as the gel

was running. Unintentionally, unequal protein:DNA ratios may also yield the free DNA band

because the DNA concentration was determined by its absorbance value at 260 nm. Alternate

DNA quantification methods, such as picogreen method (197), could address this issue.

However, Figure 4.15 suggests that unequal loading may not be the case. Figure 4.15 shows that

the free DNA disappears as ds-DNA/KF shifts from complex S to complex F. Figure 4.15 thus

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supports the oligomerization model. It suggests that Klenow initially binds DNA in 2:1

stoichiometry and shifts to a 1:1 binding mode over time. However, even Figure 4.15 only

clearly shows this pattern for hp-46 DNA, and not for ds-20/20.

Despite some supporting data, a variety of other data contradict the oligomerization

model. Fluorescence anisotropy and isothermal titration calorimetry experiments indicate that all

DNA/KF and DNA/KTQ complexes are 1:1 stoichiometry even when titrated to high

protein:DNA ratios (Table 3.2). Figure 4.17 shows that with pt-DNA/KF, complex S forms

under higher protein:DNA ratios, however, complex F does not completely shift to complex S at

high protein:DNA ratios. In general, this result contrasts with the oligomerization model because

one might expect to chase all of complex F into complex S at higher protein:DNA ratios.

However, if the ΔΔG between complexes F and S is less than a kcal/mol, complex F may not

completely shift to complex S because the stabilities of these complexes would be quite similar.

Determinations of the Kd and ΔG values of the complexes in Figure 4.17 by either visual

inspection or the program ImageQuant give values of approximately 200 nM (-9.1 kcal/mol) for

complex F, and 600 nM (-8.4 kcal/mol) for complex S. Therefore, complex F may not

completely shift to complex S in pt-DNA/KF binding.

Figures 4.1, 4.14, 4.18, 4.21 and 4.22 show that there is also a band that migrates slightly

slower than complex S. This slower band may be different from complex S and may be the

higher stoichiometry form of the DNA/protein complexes. If this slower band is a 2:1 complex,

then complex S would not be a 2:1 complex. A key question is whether we are mis-identifying

two very similarly migrating bands as the same “complex S.” In other words, does complex S for

pt-DNA/KF seen at high protein:DNA ratios almost co-migrate with the complex S for ds-

DNA/KF and DNA/KTQ complexes? Unfortunately, none of our current data can answer this

119

question. Running pt-DNA/protein and ds-DNA/protein at lower versus higher protein:DNA

ratios on the same gel could resolve this question.

Dynamic light scattering experiments can help determine if the oligomerization model is

possible by comparing the sizes of pt-DNA/KF vs. ds-DNA/KF and DNA/KF vs. DNA KTQ.

Using dynamic light scattering, one can determine the radius of gyration and the size of the

molecules in a shorter period of incubation time than with analytical ultracentrifugation (198-

200). So, for example, if the size of equimolar pt-DNA/KF complex is different from equimolar

ds-DNA/KF complex at shorter incubation times, the oligomerization model would be supported

for ds-DNA.

5.3.2 The Polymerase-Editing Modes Model

In this model, complex S is DNA/protein complex in polymerase mode while complex F

is DNA/protein complex in editing mode (Figure 5.3). This model is supported by the fact that

both pt-DNA/KTQ and ds-DNA/KTQ complexes form complex S. Klentaq will bind DNA in

polymerase mode only, because Klentaq does not have any editing activity (33, 51).

For ds-DNA/KF, the time dependent shift between complex S and F suggests that

Klenow initially binds duplex DNA in polymerase mode, but when no polymerization takes

place after some time, Klenow may assume that an error has occurred and may melt the duplex

DNA and shift to the editing mode (Figures 4.1, 4.14, and 4.15). Crystallographic and

biochemical studies of DNA/Klenow interaction show that Klenow binding in the editing mode

requires the melting of the duplex DNA (43, 65, 77, 78, 79). The co-crystal structure of DNA/KF

itself may also support the polymerase and editing modes model. The crystallography technique,

which requires high concentration of sample and longer procedure times, shows that the co-

crystal structure of DNA/KF is in the editing mode (43, 44). It has, in fact, not yet been possible

to crystallize Klenow in the polymerization mode.

120

Figure 5.3: The proposed polymerase and editing modes model for DNA binding by Klenow and

Klentaq polymerases. All pt- and ds-DNA/KTQ complexes are in polymerase mode while

complex S of DNA/KF complex may be in polymerase mode and complex F of DNA/KF

complex may be in editing mode. The proposed polymerase and editing modes models in this

figure are based on similar schematic binding models illustrated in Figure 8 of reference 57.

On the other hand, other data contradict this model. A major caveat in this model is that I

have not trapped pt-DNA/KF at 1:1 stoichiometry in complex S (although complex S of pt-

DNA/KF is clearly observed at high protein:DNA ratios) (Figures 4.12 and 4.17). Based on other

solution studies (58, 60, 64), one would not expect matched pt-DNA to form 100% editing mode

complexes with Klenow, so the current inability to trap a potential polymerase mode complex

with matched pt-DNA argues against complex S being the polymerization mode for Klenow. For

example, Klenow has been suggested to bind DNA more in the polymerization mode in the

absence of magnesium ions and more in the editing mode in the presence of magnesium ions

121

based on a recent 2-AP dimer spectroscopic study (64). However, at equimolar ratio, pt-DNA/KF

cannot be captured in complex S form even without magnesium (Figure 4.20). Some studies

have also suggested that matched pt-DNA binds primarily (~85%) to the polymerase site of

Klenow while 4 mismatched pt-DNA exclusively binds to Klenow’s editing site (58, 60), so,

Figures 4.1, 4.12, and 4.14 do not support the polymerase and editing modes model because

matched and mismatched DNA binding by Klenow both form complex F. If the matched pt-

DNA/KF complex is in the polymerase mode while the mismatched pt-DNA/KF is in the editing

mode, this data may indicate that complex F represents both polymerase and editing modes of pt-

DNA binding by Klenow, and that ds-DNA binds to Klenow in a “unique” binding mode that is

neither polymerization nor editing mode.

5.3.3 The Unique ds-DNA/KF Binding Model

The oligomerization and the polymerase and editing modes models are the most

conservative models because both modes of binding have been observed previously (58-60).

However, I am the first to examine blunt-end DNA binding, and it is possible that Klenow binds

this DNA uniquely. For the following discussion the “new” Klenow binding mode simply

represents a binding complex that is not identical to either the polymerase or editing mode

complexes. It may be an end-binding complex, an intermediate between the polymerase mode

and editing mode, or a previously unidentified binding topology.

This “new” mode of ds-DNA/KF binding may be similar to the polymerase mode of

binding. This unique ds-DNA/KF binding model proposes that complexes S and F for ds-

DNA/KF correspond to “new” mode and editing mode for ds-DNA/KF. Figure 5.4A is a

schematic showing ds-DNA/KF in such a “new” binding mode while Figure 5.4B shows the

editing mode for ds-DNA/KF binding. Figure 5.4B is based on reference 57 while Figure 5.4A is

hypothesized based on the fact that the 3'-end of the primer strand binds to the polymerase site of

122

Klenow but that the 5'-end of the template strand will not bind to the RRRY site because the

DNA is blunt-end.

Figure 5.4: The proposed unique ds-DNA/KF binding model. Complex S of DNA/KF complex

(A) may be in a “new” ds-DNA binding mode and complex F for DNA/KF complex (B) may be

in editing mode where Klenow may melt the duplex DNA. Figure 5.4A is based on the fact that

the 3'-end of the primer strand binds to the polymerase site of Klenow but the 5'-end of the

template strand cannot bind to the RRRY site because the DNA is blunt-end. Figure 5.4B is

adapted from a similar schematic depicting KF editing mode binding in Figure 8 of reference 57.

123

Figure 4.15 shows that the ds-DNA/KF complex shifts from complex S to F over time.

Similar to the explanation for the polymerase and editing modes model, that data suggest that

Klenow binds ds-DNA in the “new” mode at first. Then, Klenow melts ds-DNA and converts to

the editing mode. However, without structural data and further biochemical experimentation, this

hypothesis cannot be confirmed.

Experiments using 2-aminopurine may help to determine if the polymerase and editing

modes model and/or the unique ds-DNA/KF binding model are the more accurate models. 2-

aminopurine has previously been used to monitor whether DNA is in a duplex or single-strand

(162, 201-205). For 2-aminopurine experiments, one might determine if the ds-DNA/KF is

melted after a longer incubation time by measuring the differences in 2-aminopurine

fluorescence after shorter and longer incubations (162, 201-205). A change in the 2-AP

fluorescence over time would indicate that the DNA duplex is being melted over time in the ds-

DNA/KF complex.

5.4 Summary

In summary, none of the three models (e.g. the oligomerization model, the polymerase

and editing modes model, or the unique ds-DNA/KF binding model) is fully supported by the

current data from any laboratory. Table 5.1 summarizes the supporting and opposing data for the

oligomerization and the polymerase and editing modes models. Overall, the oligomerization

model is primarily supported by the EMSA experiments, and contradicted by the stoichiometry

and heat capacity change data from fluorescence anisotropy and isothermal titration calorimetry

experiments. On the other hand, the polymerase and editing modes model is mostly contradicted

by the “missing” data of not being able to trap pt-DNA/KF in complex S at equimolar

protein:DNA ratios, but is supported by the stoichiometry and heat capacity change data. If

additional dynamic light scattering and 2-AP fluorescence experiments do not confirm any of

124

these models, this may suggest that there could be more than one model occuring

simultaneously. For example, complex S for Klenow may contain both 2:1 and polymerase

binding modes while complex F consists of 1:1 and editing modes.

Table 5.1: Summary of data that support and contradict the oligomerization and the polymerase

and editing modes models. The polymerase and editing modes model for DNA/KTQ is not

considered because Klentaq binds DNA in polymerase mode only. S = supporting data, C =

contradicting data, and N = neither supporting nor contradicting data (neutral).

Experiments

DNA/KF

Oligomerization

Model

DNA/KF

Polymerase and

Editing Modes Model

DNA/KTQ

Oligomerization

Model

Stoichiometry C S C

ΔG S C N

ΔCp C S S

Analytical

Ultracentrifugation N N C

Circular Dichroism N N N

Electrophoretic

Mobility Shift Assay S C S

The data of this dissertation have shown that the interactions of these supposedly

homologous polymerases to different DNA structures are somewhat different, because of

demonstrated thermodynamic and structural differences in the DNA structural preferences of

Klenow versus Klentaq polymerases. In particular, these studies show that Klenow binds pt-

DNA and ds-DNA differently while Klentaq does not, however, the most realistic molecular

model for these protein-DNA interactions remains to be determined.

125

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VITA

Andy James Budiman Wowor was born in Jakarta, Indonesia, in December 1981 to

Djony Wowor and Jenny Nirmalasari Sholihin. Andy attended Kemurnian I School, Jakarta,

Indonesia, for kindergarten and Ricci I Catholic School, Jakarta, Indonesia, for the elementary

and junior high schools. He graduated from Sekolah Menengah Umum Kristen III Badan

Pendidikan Kristen Penabur (Christian Education Foundation Penabur), Jakarta, Indonesia, in

June 2000. That August he entered Southern Arkansas University, Magnolia, Arkansas, and

transferred to Louisiana State University, Baton Rouge, Louisiana, in August 2002. Andy

received the Bachelor of Science in chemistry from Louisiana State University with magna cum

laude in May 2004. He entered the graduate program at Louisiana State University in the

Department of Biological Sciences in the Fall of 2004 and became a doctoral student in the

laboratory of Dr. Vince J. LiCata. Andy is a candidate for the Doctor of Philosophy degree in

biochemistry on December 18, 2009.