dna structural selectivity of binding by the pol i dna
TRANSCRIPT
Louisiana State University Louisiana State University
LSU Digital Commons LSU Digital Commons
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
Follow this and additional works at: https://digitalcommons.lsu.edu/gradschool_dissertations
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
This Dissertation is brought to you for free and open access by the Graduate School at LSU Digital Commons. It has been accepted for inclusion in LSU Doctoral Dissertations by an authorized graduate school editor of LSU Digital Commons. For more information, please [email protected].
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!!!
iii
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.
iv
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.
96
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
97
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.
98
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)
99
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).
102
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.
103
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
104
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.
105
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.
106
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.
109
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
113
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
114
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.
116
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
118
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
REFERENCES
1. Bebenek, K., and Kunkel, T. A. (2004) Functions of DNA polymerases. Adv Protein
Chem 69, 137-165
2. Prakash, S., Johnson, R. E., and Prakash, L. (2005) Eukaryotic translesion synthesis DNA
polymerases: specificity of structure and function. Annu Rev Biochem 74, 317-353
3. Tang, K. H., and Tsai, M. D. (2008) Structure and function of 2:1 DNA polymerase.DNA
complexes. J Cell Physiol 216, 315-320
4. Krahn, J. M., Beard, W. A., Miller, H., Grollman, A. P., and Wilson, S. H. (2003)
Structure of DNA polymerase beta with the mutagenic DNA lesion 8-oxodeoxyguanine
reveals structural insights into its coding potential. Structure 11, 121-127
5. Ling, H., Boudsocq, F., Plosky, B. S., Woodgate, R., and Yang, W. (2003) Replication of
a cis-syn thymine dimer at atomic resolution. Nature 424, 1083-1087
6. Brieba, L. G., Eichman, B. F., Kokoska, R. J., Doublie, S., Kunkel, T. A., and
Ellenberger, T. (2004) Structural basis for the dual coding potential of 8-oxoguanosine by
a high-fidelity DNA polymerase. EMBO J 23, 3452-3461
7. Hsu, G. W., Ober, M., Carell, T., and Beese, L. S. (2004) Error-prone replication of
oxidatively damaged DNA by a high-fidelity DNA polymerase. Nature 431, 217-221
8. Johnson, S. J., and Beese, L. S. (2004) Structures of mismatch replication errors observed
in a DNA polymerase. Cell 116, 803-816
9. Hsu, G. W., Huang, X., Luneva, N. P., Geacintov, N. E., and Beese, L. S. (2005)
Structure of a high fidelity DNA polymerase bound to a benzo[a]pyrene adduct that
blocks replication. J Biol Chem 280, 3764-3770
10. Batra, V. K., Beard, W. A., Shock, D. D., Krahn, J. M., Pedersen, L. C., and Wilson, S.
H. (2006) Magnesium-induced assembly of a complete DNA polymerase catalytic
complex. Structure 14, 757-766
11. Garcia-Diaz, M., Bebenek, K., Krahn, J. M., Pedersen, L. C., and Kunkel, T. A. (2006)
Structural analysis of strand misalignment during DNA synthesis by a human DNA
polymerase. Cell 124, 331-342
12. Kamtekar, S., Berman, A. J., Wang, J., Lazaro, J. M., de Vega, M., Blanco, L., Salas, M.,
and Steitz, T. A. (2004) Insights into strand displacement and processivity from the
crystal structure of the protein-primed DNA polymerase of bacteriophage phi29. Mol Cell
16, 609-618
13. Garcia-Diaz, M., Bebenek, K., Gao, G., Pedersen, L. C., London, R. E., and Kunkel, T.
A. (2005) Structure-function studies of DNA polymerase lambda. DNA Repair (Amst) 4,
1358-1367
126
14. Rodriguez, I., Lazaro, J. M., Blanco, L., Kamtekar, S., Berman, A. J., Wang, J., Steitz, T.
A., Salas, M., and de Vega, M. (2005) A specific subdomain in phi29 DNA polymerase
confers both processivity and strand-displacement capacity. Proc Natl Acad Sci U S A
102, 6407-6412
15. Sadava, D. E., Heller, H. C., Orians, G. H., Purves, W. K., and Hillis, D. (2006) Life.
New York: Sinauer Associates, Inc., and W. H. Freeman and Company. 8th
Edition.
16. McCulloch, S. D., and Kunkel, T. A. (2008) The fidelity of DNA synthesis by eukaryotic
replicative and translesion synthesis polymerases. Cell Res 18, 148-161
17. Kornberg, A. (1988) DNA replication. J Biol Chem 263, 1-4
18. Lehman, I. R., and Uyemura, D. G. (1976) DNA polymerase I: essential replication
enzyme. Science 193, 963-969
19. Kornberg, A., and Baker, T. A. (1992) DNA Replication. New York: W. H. Freeman and
Company. 2nd
Edition.
20. Garrett, R. H., and Grisham, C. M. (2008) Biochemistry. St. Paul: Brooks/Cole
Publishing Company. 4th
Edition.
21. Lodish, H., Berk, A., Zipursky, L. S., Matsudaira, P., Baltimore, D., and Darnell, J.
(2000) Molecular Cell Biology. New York: W. H. Freeman and Company. 4th
Edition.
22. Delarue, M., Poch, O., Tordo, N., Moras, D., and Argos, P. (1990) An attempt to unify
the structure of polymerases. Protein Eng 3, 461-467
23. Ito, J., and Braithwaite, D. K. (1991) Compilation and alignment of DNA polymerase
sequences. Nucleic Acids Res 19, 4045-4057
24. Braithwaite, D. K., and Ito, J. (1993) Compilation, alignment, and phylogenetic
relationships of DNA polymerases. Nucleic Acids Res 21, 787-802
25. Joyce, C. M., and Steitz, T. A. (1994) Function and structure relationships in DNA
polymerases. Annu Rev Biochem 63, 777-822
26. Joyce, C. M., and Steitz, T. A. (1995) Polymerase structures and function: variations on a
theme? J Bacteriol 177, 6321-6329
27. Steitz, T. A. (1999) DNA polymerases: structural diversity and common mechanisms. J
Biol Chem 274, 17395-17398
28. Rothwell, P. J., and Waksman, G. (2005) Structure and mechanism of DNA polymerases.
Adv Protein Chem 71, 401-440
29. Bessman, M. J., Kornberg, A., Lehman, I. R., and Simms, E. S. (1956) Enzymic synthesis
of deoxyribonucleic acid. Biochim Biophys Acta 21, 197-198
127
30. Lehman, I. R., Zimmerman, S. B., Adler, J., Bessman, M. J., Simms, E. S., and Kornberg,
A. (1958) Enzymatic Synthesis of Deoxyribonucleic Acid. V. Chemical Composition of
Enzymatically Synthesized Deoxyribonucleic Acid. Proc Natl Acad Sci U S A 44, 1191-
1196
31. Jovin, T. M., Englund, P. T., and Bertsch, L. L. (1969) Enzymatic synthesis of
deoxyribonucleic acid. XXVI. Physical and chemical studies of a homogeneous
deoxyribonucleic acid polymerase. J Biol Chem 244, 2996-3008
32. Beese, L. S., Friedman, J. M., and Steitz, T. A. (1993) Crystal structures of the Klenow
fragment of DNA polymerase I complexed with deoxynucleoside triphosphate and
pyrophosphate. Biochemistry 32, 14095-14101
33. Lawyer, F. C., Stoffel, S., Saiki, R. K., Myambo, K., Drummond, R., and Gelfand, D. H.
(1989) Isolation, characterization, and expression in Escherichia coli of the DNA
polymerase gene from Thermus aquaticus. J Biol Chem 264, 6427-6437
34. Brock, T. D., and Freeze, H. (1969) Thermus aquaticus gen. n. and sp. n., a
nonsporulating extreme thermophile. J Bacteriol 98, 289-297
35. Chien, A., Edgar, D. B., and Trela, J. M. (1976) Deoxyribonucleic acid polymerase from
the extreme thermophile Thermus aquaticus. J Bacteriol 127, 1550-1557
36. Engelke, D. R., Krikos, A., Bruck, M. E., and Ginsburg, D. (1990) Purification of
Thermus aquaticus DNA polymerase expressed in Escherichia coli. Anal Biochem 191,
396-400
37. Li, Y., and Waksman, G. (2001) Crystal structures of a ddATP-, ddTTP-, ddCTP, and
ddGTP- trapped ternary complex of Klentaq1: insights into nucleotide incorporation and
selectivity. Protein Sci 10, 1225-1233
38. Basu, A., and Modak, M. J. (1987) Identification and amino acid sequence of the
deoxynucleoside triphosphate binding site in Escherichia coli DNA polymerase I.
Biochemistry 26, 1704-1709
39. Basu, A., Williams, K. R., and Modak, M. J. (1987) Ferrate oxidation of Escherichia coli
DNA polymerase-I. Identification of a methionine residue that is essential for DNA
binding. J Biol Chem 262, 9601-9607
40. Mohan, P. M., Basu, A., Basu, S., Abraham, K. I., and Modak, M. J. (1988) DNA
binding domain of Escherichia coli DNA polymerase I: identification of arginine-841 as
an essential residue. Biochemistry 27, 226-233
41. Pandey, V. N., and Modak, M. J. (1988) Affinity labeling of Escherichia coli DNA
polymerase I by 5'-fluorosulfonylbenzoyladenosine. Identification of the domain essential
for polymerization and Arg-682 as the site of reactivity. J Biol Chem 263, 6068-6073
42. Pandey, V. N., Williams, K. R., Stone, K. L., and Modak, M. J. (1987) Photoaffinity
labeling of the thymidine triphosphate binding domain in Escherichia coli DNA
128
polymerase I: identification of histidine-881 as the site of cross-linking. Biochemistry 26,
7744-7748
43. Beese, L. S., Derbyshire, V., and Steitz, T. A. (1993) Structure of DNA polymerase I
Klenow fragment bound to duplex DNA. Science 260, 352-355
44. Kohlstaedt, L. A., Wang, J., Friedman, J. M., Rice, P. A., and Steitz, T. A. (1992) Crystal
structure at 3.5 A resolution of HIV-1 reverse transcriptase complexed with an inhibitor.
Science 256, 1783-1790
45. Derbyshire, V., Freemont, P. S., Sanderson, M. R., Beese, L., Friedman, J. M., Joyce, C.
M., and Steitz, T. A. (1988) Genetic and crystallographic studies of the 3',5'-
exonucleolytic site of DNA polymerase I. Science 240, 199-201
46. Ollis, D. L., Brick, P., Hamlin, R., Xuong, N. G., and Steitz, T. A. (1985) Structure of
large fragment of Escherichia coli DNA polymerase I complexed with dTMP. Nature
313, 762-766
47. Barnes, W. M. (1992) The fidelity of Taq polymerase catalyzing PCR is improved by an
N-terminal deletion. Gene 112, 29-35
48. Klenow, H., and Overgaard-Hansen, K. (1970) Proteolytic cleavage of DNA polymerase
from Escherichia coli B into an exonuclease unit and a polymerase unit. FEBS Lett 6, 25-
27
49. Karantzeni, I., Ruiz, C., Liu, C. C., and Licata, V. J. (2003) Comparative thermal
denaturation of Thermus aquaticus and Escherichia coli type 1 DNA polymerases.
Biochem J 374, 785-792
50. Richard, A. J., Liu, C. C., Klinger, A. L., Todd, M. J., Mezzasalma, T. M., and LiCata, V.
J. (2006) Thermal stability landscape for Klenow DNA polymerase as a function of pH
and salt concentration. Biochim Biophys Acta 1764, 1546-1552
51. Korolev, S., Nayal, M., Barnes, W. M., Di Cera, E., and Waksman, G. (1995) Crystal
structure of the large fragment of Thermus aquaticus DNA polymerase I at 2.5-A
resolution: structural basis for thermostability. Proc Natl Acad Sci U S A 92, 9264-9268
52. Li, Y., Kong, Y., Korolev, S., and Waksman, G. (1998) Crystal structures of the Klenow
fragment of Thermus aquaticus DNA polymerase I complexed with deoxyribonucleoside
triphosphates. Protein Sci 7, 1116-1123
53. Li, Y., Korolev, S., and Waksman, G. (1998) Crystal structures of open and closed forms
of binary and ternary complexes of the large fragment of Thermus aquaticus DNA
polymerase I: structural basis for nucleotide incorporation. EMBO J 17, 7514-7525
54. Doublie, S., Tabor, S., Long, A. M., Richardson, C. C., and Ellenberger, T. (1998)
Crystal structure of a bacteriophage T7 DNA replication complex at 2.2 A resolution.
Nature 391, 251-258
129
55. Kiefer, J. R., Mao, C., Braman, J. C., and Beese, L. S. (1998) Visualizing DNA
replication in a catalytically active Bacillus DNA polymerase crystal. Nature 391, 304-
307
56. Turner, R. M., Jr., Grindley, N. D., and Joyce, C. M. (2003) Interaction of DNA
polymerase I (Klenow fragment) with the single-stranded template beyond the site of
synthesis. Biochemistry 42, 2373-2385
57. Kukreti, P., Singh, K., Ketkar, A., and Modak, M. J. (2008) Identification of a new motif
required for the 3'-5' exonuclease activity of Escherichia coli DNA polymerase I
(Klenow fragment): the RRRY motif is necessary for the binding of single-stranded DNA
substrate and the template strand of the mismatched duplex. J Biol Chem 283, 17979-
17990
58. Bailey, M. F., Thompson, E. H., and Millar, D. P. (2001) Probing DNA polymerase
fidelity mechanisms using time-resolved fluorescence anisotropy. Methods 25, 62-77
59. Bailey, M. F., van der Schans, E. J., and Millar, D. P. (2004) Thermodynamic dissection
of the polymerizing and editing modes of a DNA polymerase. J Mol Biol 336, 673-693
60. Carver, T. E., Jr., Hochstrasser, R. A., and Millar, D. P. (1994) Proofreading DNA:
recognition of aberrant DNA termini by the Klenow fragment of DNA polymerase I.
Proc Natl Acad Sci U S A 91, 10670-10674
61. Datta, K., Johnson, N. P., and von Hippel, P. H. (2006) Mapping the conformation of the
nucleic acid framework of the T7 RNA polymerase elongation complex in solution using
low-energy CD and fluorescence spectroscopy. J Mol Biol 360, 800-813
62. Johnson, N. P., Baase, W. A., and Von Hippel, P. H. (2004) Low-energy circular
dichroism of 2-aminopurine dinucleotide as a probe of local conformation of DNA and
RNA. Proc Natl Acad Sci U S A 101, 3426-3431
63. Joyce, C. M. (1989) How DNA travels between the separate polymerase and 3'-5'-
exonuclease sites of DNA polymerase I (Klenow fragment). J Biol Chem 264, 10858-
10866
64. Datta, K., Johnson, N. P., Licata, V. J., and von Hippel, P. H. (2009) Local conformations
and competitive binding affinities of single- and double-stranded primer-template DNA
at the polymerization and editing active sites of DNA polymerase. J Biol Chem
65. Cowart, M., Gibson, K. J., Allen, D. J., and Benkovic, S. J. (1989) DNA substrate
structural requirements for the exonuclease and polymerase activities of procaryotic and
phage DNA polymerases. Biochemistry 28, 1975-1983
66. Shamoo, Y., and Steitz, T. A. (1999) Building a replisome from interacting pieces:
sliding clamp complexed to a peptide from DNA polymerase and a polymerase editing
complex. Cell 99, 155-166
130
67. Franklin, M. C., Wang, J., and Steitz, T. A. (2001) Structure of the replicating complex of
a pol alpha family DNA polymerase. Cell 105, 657-667
68. Gangurde, R., and Modak, M. J. (2002) Participation of active-site carboxylates of
Escherichia coli DNA polymerase I (Klenow fragment) in the formation of a
prepolymerase ternary complex. Biochemistry 41, 14552-14559
69. Joyce, C. M., Potapova, O., Delucia, A. M., Huang, X., Basu, V. P., and Grindley, N. D.
(2008) Fingers-closing and other rapid conformational changes in DNA polymerase I
(Klenow fragment) and their role in nucleotide selectivity. Biochemistry 47, 6103-6116
70. Tindall, K. R., and Kunkel, T. A. (1988) Fidelity of DNA synthesis by the Thermus
aquaticus DNA polymerase. Biochemistry 27, 6008-6013
71. Kim, Y., Eom, S. H., Wang, J., Lee, D. S., Suh, S. W., and Steitz, T. A. (1995) Crystal
structure of Thermus aquaticus DNA polymerase. Nature 376, 612-616
72. Perler, F. B., Kumar, S., and Kong, H. (1996) Thermostable DNA polymerases. Adv
Protein Chem 48, 377-435
73. Kuchta, R. D., Mizrahi, V., Benkovic, P. A., Johnson, K. A., and Benkovic, S. J. (1987)
Kinetic mechanism of DNA polymerase I (Klenow). Biochemistry 26, 8410-8417
74. Steitz, T. A. (1998) A mechanism for all polymerases. Nature 391, 231-232
75. Beese, L. S., and Steitz, T. A. (1991) Structural basis for the 3'-5' exonuclease activity of
Escherichia coli DNA polymerase I: a two metal ion mechanism. EMBO J 10, 25-33
76. Brautigam, C. A., and Steitz, T. A. (1998) Structural and functional insights provided by
crystal structures of DNA polymerases and their substrate complexes. Curr Opin Struct
Biol 8, 54-63
77. Catalano, C. E., Allen, D. J., and Benkovic, S. J. (1990) Interaction of Escherichia coli
DNA polymerase I with azidoDNA and fluorescent DNA probes: identification of
protein-DNA contacts. Biochemistry 29, 3612-3621
78. Freemont, P. S., Friedman, J. M., Beese, L. S., Sanderson, M. R., and Steitz, T. A. (1988)
Cocrystal structure of an editing complex of Klenow fragment with DNA. Proc Natl
Acad Sci U S A 85, 8924-8928
79. Derbyshire, V., Pinsonneault, J. K., and Joyce, C. M. (1995) Structure-function analysis
of 3'-->5'-exonuclease of DNA polymerases. Methods Enzymol 262, 363-385
80. Freemont, P. S., Ollis, D. L., Steitz, T. A., and Joyce, C. M. (1986) A domain of the
Klenow fragment of Escherichia coli DNA polymerase I has polymerase but no
exonuclease activity. Proteins 1, 66-73
131
81. Derbyshire, V., Grindley, N. D., and Joyce, C. M. (1991) The 3'-5' exonuclease of DNA
polymerase I of Escherichia coli: contribution of each amino acid at the active site to the
reaction. EMBO J 10, 17-24
82. Brautigam, C. A., Sun, S., Piccirilli, J. A., and Steitz, T. A. (1999) Structures of normal
single-stranded DNA and deoxyribo-3'-S-phosphorothiolates bound to the 3'-5'
exonucleolytic active site of DNA polymerase I from Escherichia coli. Biochemistry 38,
696-704
83. Brutlag, D., and Kornberg, A. (1972) Enzymatic synthesis of deoxyribonucleic acid. 36.
A proofreading function for the 3' leads to 5' exonuclease activity in deoxyribonucleic
acid polymerases. J Biol Chem 247, 241-248
84. Kunkel, T. A., and Loeb, L. A. (1980) On the fidelity of DNA replication. The accuracy
of Escherichia coli DNA polymerase I in copying natural DNA in vitro. J Biol Chem
255, 9961-9966
85. Bebenek, K., Joyce, C. M., Fitzgerald, M. P., and Kunkel, T. A. (1990) The fidelity of
DNA synthesis catalyzed by derivatives of Escherichia coli DNA polymerase I. J Biol
Chem 265, 13878-13887
86. Eger, B. T., Kuchta, R. D., Carroll, S. S., Benkovic, P. A., Dahlberg, M. E., Joyce, C. M.,
and Benkovic, S. J. (1991) Mechanism of DNA replication fidelity for three mutants of
DNA polymerase I: Klenow fragment KF(exo+), KF(polA5), and KF(exo-). Biochemistry
30, 1441-1448
87. Keohavong, P., and Thilly, W. G. (1989) Fidelity of DNA polymerases in DNA
amplification. Proc Natl Acad Sci U S A 86, 9253-9257
88. Cariello, N. F., Swenberg, J. A., De Bellis, A., and Skopek, T. R. (1991) Analysis of
mutations using PCR and denaturing gradient gel electrophoresis. Environ Mol Mutagen
18, 249-254
89. Ling, L. L., Keohavong, P., Dias, C., and Thilly, W. G. (1991) Optimization of the
polymerase chain reaction with regard to fidelity: modified T7, Taq, and vent DNA
polymerases. PCR Methods Appl 1, 63-69
90. Lundberg, K. S., Shoemaker, D. D., Adams, M. W., Short, J. M., Sorge, J. A., and
Mathur, E. J. (1991) High-fidelity amplification using a thermostable DNA polymerase
isolated from Pyrococcus furiosus. Gene 108, 1-6
91. Echols, H., and Goodman, M. F. (1991) Fidelity mechanisms in DNA replication. Annu
Rev Biochem 60, 477-511
92. Lohman, T. M., and Mascotti, D. P. (1992) Thermodynamics of ligand-nucleic acid
interactions. Methods Enzymol 212, 400-424
93. Goeddel, D. V., Yansura, D. G., and Caruthers, M. H. (1978) How lac repressor
recognizes lac operator. Proc Natl Acad Sci U S A 75, 3578-3582
132
94. Fried, M. G. (1989) Measurement of protein-DNA interaction parameters by
electrophoresis mobility shift assay. Electrophoresis 10, 366-376
95. Heyduk, T., Ma, Y., Tang, H., and Ebright, R. H. (1996) Fluorescence anisotropy: rapid,
quantitative assay for protein-DNA and protein-protein interaction. Methods Enzymol
274, 492-503
96. Hill, J. J., and Royer, C. A. (1997) Fluorescence approaches to study of protein-nucleic
acid complexation. Methods Enzymol 278, 390-416
97. Datta, K., and LiCata, V. J. (2003) Thermodynamics of the binding of Thermus aquaticus
DNA polymerase to primed-template DNA. Nucleic Acids Res 31, 5590-5597
98. Datta, K., Wowor, A. J., Richard, A. J., and LiCata, V. J. (2006) Temperature
dependence and thermodynamics of Klenow polymerase binding to primed-template
DNA. Biophys J 90, 1739-1751
99. Datta, K., and LiCata, V. J. (2003) Salt dependence of DNA binding by Thermus
aquaticus and Escherichia coli DNA polymerases. J Biol Chem 278, 5694-5701
100. Sturtevant, J. M. (1977) Heat capacity and entropy changes in processes involving
proteins. Proc Natl Acad Sci U S A 74, 2236-2240
101. Gill, S. J., Dec, S. F., Olofsson, G., and Wadso, I. (1985) Anomalous heat capacity of
hydrophobic solvation. J Phys Chem 89, 3758-3761
102. Privalov, P. L., and Gill, S. J. (1988) Stability of protein structure and hydrophobic
interaction. Adv Protein Chem 39, 191-234
103. Robertson, A. D., and Murphy, K. P. (1997) Protein Structure and the Energetics of
Protein Stability. Chem Rev 97, 1251-1268
104. Dill, K. A., and Bromberg, S. (2003) Molecular Driving Forces. New York: Garland
Science.
105. Petri, V., Hsieh, M., and Brenowitz, M. (1995) Thermodynamic and kinetic
characterization of the binding of the TATA binding protein to the adenovirus E4
promoter. Biochemistry 34, 9977-9984
106. Kozlov, A. G., and Lohman, T. M. (1999) Adenine base unstacking dominates the
observed enthalpy and heat capacity changes for the Escherichia coli SSB tetramer
binding to single-stranded oligoadenylates. Biochemistry 38, 7388-7397
107. Ladbury, J. E., Wright, J. G., Sturtevant, J. M., and Sigler, P. B. (1994) A thermodynamic
study of the trp repressor-operator interaction. J Mol Biol 238, 669-681
108. Dragan, A. I., Klass, J., Read, C., Churchill, M. E., Crane-Robinson, C., and Privalov, P.
L. (2003) DNA binding of a non-sequence-specific HMG-D protein is entropy driven
with a substantial non-electrostatic contribution. J Mol Biol 331, 795-813
133
109. Kozlov, A. G., and Lohman, T. M. (2000) Large contributions of coupled protonation
equilibria to the observed enthalpy and heat capacity changes for ssDNA binding to
Escherichia coli SSB protein. Proteins Suppl 4, 8-22
110. Lundback, T., van Den Berg, S., and Hard, T. (2000) Sequence-specific DNA binding by
the glucocorticoid receptor DNA-binding domain is linked to a salt-dependent histidine
protonation. Biochemistry 39, 8909-8916
111. Cooper, A. (2005) Heat capacity effects in protein folding and ligand binding: a re-
evaluation of the role of water in biomolecular thermodynamics. Biophys Chem 115, 89-
97
112. Ha, J. H., Spolar, R. S., and Record, M. T., Jr. (1989) Role of the hydrophobic effect in
stability of site-specific protein-DNA complexes. J Mol Biol 209, 801-816
113. Murphy, K. P., and Freire, E. (1992) Thermodynamics of structural stability and
cooperative folding behavior in proteins. Adv Protein Chem 43, 313-361
114. Spolar, R. S., Livingstone, J. R., and Record, M. T., Jr. (1992) Use of liquid hydrocarbon
and amide transfer data to estimate contributions to thermodynamic functions of protein
folding from the removal of nonpolar and polar surface from water. Biochemistry 31,
3947-3955
115. Spolar, R. S., and Record, M. T., Jr. (1994) Coupling of local folding to site-specific
binding of proteins to DNA. Science 263, 777-784
116. Makhatadze, G. I., and Privalov, P. L. (1995) Energetics of protein structure. Adv Protein
Chem 47, 307-425
117. Myers, J. K., Pace, C. N., and Scholtz, J. M. (1995) Denaturant m values and heat
capacity changes: relation to changes in accessible surface areas of protein unfolding.
Protein Sci 4, 2138-2148
118. Jen-Jacobson, L. (1997) Protein-DNA recognition complexes: conservation of structure
and binding energy in the transition state. Biopolymers 44, 153-180
119. Ferrari, M. E., and Lohman, T. M. (1994) Apparent heat capacity change accompanying a
nonspecific protein-DNA interaction. Escherichia coli SSB tetramer binding to
oligodeoxyadenylates. Biochemistry 33, 12896-12910
120. Lundback, T., Hansson, H., Knapp, S., Ladenstein, R., and Hard, T. (1998)
Thermodynamic characterization of non-sequence-specific DNA-binding by the Sso7d
protein from Sulfolobus solfataricus. J Mol Biol 276, 775-786
121. Milev, S., Gorfe, A. A., Karshikoff, A., Clubb, R. T., Bosshard, H. R., and Jelesarov, I.
(2003) Energetics of sequence-specific protein-DNA association: binding of integrase
Tn916 to its target DNA. Biochemistry 42, 3481-3491
134
122. Kozlov, A. G., and Lohman, T. M. (2006) Effects of monovalent anions on a
temperature-dependent heat capacity change for Escherichia coli SSB tetramer binding to
single-stranded DNA. Biochemistry 45, 5190-5205
123. Liu, C. C., Richard, A. J., Datta, K., and LiCata, V. J. (2008) Prevalence of temperature-
dependent heat capacity changes in protein-DNA interactions. Biophys J 94, 3258-3265
124. Poon, K., and Macgregor, R. B., Jr. (1998) Unusual behavior exhibited by multistranded
guanine-rich DNA complexes. Biopolymers 45, 427-434
125. Jezewska, M. J., Rajendran, S., and Bujalowski, W. (1998) Transition between different
binding modes in rat DNA polymerase beta-ssDNA complexes. J Mol Biol 284, 1113-
1131
126. Salinas, F., and Benkovic, S. J. (2000) Characterization of bacteriophage T4-coordinated
leading- and lagging-strand synthesis on a minicircle substrate. Proc Natl Acad Sci U S A
97, 7196-7201
127. Tsoi, P. Y., and Yang, M. (2002) Kinetic study of various binding modes between human
DNA polymerase beta and different DNA substrates by surface-plasmon-resonance
biosensor. Biochem J 361, 317-325
128. Ishmael, F. T., Trakselis, M. A., and Benkovic, S. J. (2003) Protein-protein interactions in
the bacteriophage T4 replisome. The leading strand holoenzyme is physically linked to
the lagging strand holoenzyme and the primosome. J Biol Chem 278, 3145-3152
129. Joubert, A. M., Byrd, A. S., and LiCata, V. J. (2003) Global conformations,
hydrodynamics, and X-ray scattering properties of Taq and Escherichia coli DNA
polymerases in solution. J Biol Chem 278, 25341-25347
130. Ho, D. L., Byrnes, W. M., Ma, W. P., Shi, Y., Callaway, D. J., and Bu, Z. (2004)
Structure-specific DNA-induced conformational changes in Taq polymerase revealed by
small angle neutron scattering. J Biol Chem 279, 39146-39154
131. Bailey, M. F., Van der Schans, E. J., and Millar, D. P. (2007) Dimerization of the Klenow
fragment of Escherichia coli DNA polymerase I is linked to its mode of DNA binding.
Biochemistry 46, 8085-8099
132. Tang, K. H., Niebuhr, M., Aulabaugh, A., and Tsai, M. D. (2008) Solution structures of 2
: 1 and 1 : 1 DNA polymerase-DNA complexes probed by ultracentrifugation and small-
angle X-ray scattering. Nucleic Acids Res 36, 849-860
133. Heyduk, T., and Lee, J. C. (1990) Application of fluorescence energy transfer and
polarization to monitor Escherichia coli cAMP receptor protein and lac promoter
interaction. Proc Natl Acad Sci U S A 87, 1744-1748
134. Riggs, A. D., Suzuki, H., and Bourgeois, S. (1970) Lac repressor-operator interaction. I.
Equilibrium studies. J Mol Biol 48, 67-83
135
135. Fried, M., and Crothers, D. M. (1981) Equilibria and kinetics of lac repressor-operator
interactions by polyacrylamide gel electrophoresis. Nucleic Acids Res 9, 6505-6525
136. Garner, M. M., and Revzin, A. (1981) A gel electrophoresis method for quantifying the
binding of proteins to specific DNA regions: application to components of the
Escherichia coli lactose operon regulatory system. Nucleic Acids Res 9, 3047-3060
137. Brown, M. P., and Royer, C. (1997) Fluorescence spectroscopy as a tool to investigate
protein interactions. Curr Opin Biotechnol 8, 45-49
138. Eftink, M. R. (1997) Fluorescence methods for studying equilibrium macromolecule-
ligand interactions. Methods Enzymol 278, 221-257
139. Chin, J., Langst, G., Becker, P. B., and Widom, J. (2004) Fluorescence anisotropy assays
for analysis of ISWI-DNA and ISWI-nucleosome interactions. Methods Enzymol 376, 3-
16
140. Jameson, D. M., and Sawyer, W. H. (1995) Fluorescence anisotropy applied to
biomolecular interactions. Methods Enzymol 246, 283-300
141. Lundblad, J. R., Laurance, M., and Goodman, R. H. (1996) Fluorescence polarization
analysis of protein-DNA and protein-protein interactions. Mol Endocrinol 10, 607-612
142. Royer, C. A. (1995) Quantitative detection of macromolecules with fluorescent
nucleotides, U.S. Patent # 5,445,935.
143. Royer, C. A. (1998) Quantitative detection of macromolecules with fluorescent
oligonucleotides, U.S. Patent # 5,756,292.
144. Royer, C. A. (2001) Quantitative detection of macromolecules with fluorescent
oligonucleotides, U.S. Patent # 6,326,142.
145. Shanker, N., and Bane, S. L. (2008) Basic aspects of absorption and fluorescence
spectroscopy and resonance energy transfer methods. Methods Cell Biol 84, 213-242
146. Waggoner, A. (1995) Covalent labeling of proteins and nucleic acids with fluorophores.
Methods Enzymol 246, 362-373
147. Perez-Howard, G. M., Weil, P. A., and Beechem, J. M. (1995) Yeast TATA binding
protein interaction with DNA: fluorescence determination of oligomeric state,
equilibrium binding, on-rate, and dissociation kinetics. Biochemistry 34, 8005-8017
148. Rusinova, E., Tretyachenko-Ladokhina, V., Vele, O. E., Senear, D. F., and Alexander
Ross, J. B. (2002) Alexa and Oregon Green dyes as fluorescence anisotropy probes for
measuring protein-protein and protein-nucleic acid interactions. Anal Biochem 308, 18-25
149. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram
quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72, 248-
254
136
150. Boyer, M., Poujol, N., Margeat, E., and Royer, C. A. (2000) Quantitative characterization
of the interaction between purified human estrogen receptor alpha and DNA using
fluorescence anisotropy. Nucleic Acids Res 28, 2494-2502
151. Jezewska, M. J., and Bujalowski, W. (1996) A general method of analysis of ligand
binding to competing macromolecules using the spectroscopic signal originating from a
reference macromolecule. Application to Escherichia coli replicative helicase DnaB
protein nucleic acid interactions. Biochemistry 35, 2117-2128
152. Zettner, A. (1973) Principles of competitive binding assays (saturation analysis). 1.
Equilibrium techniques. Clin Chem 19, 699-705
153. Burke, T. J., Loniello, K. R., Beebe, J. A., and Ervin, K. M. (2003) Development and
application of fluorescence polarization assays in drug discovery. Comb Chem High
Throughput Screen 6, 183-194
154. Parker, G. J., Law, T. L., Lenoch, F. J., and Bolger, R. E. (2000) Development of high
throughput screening assays using fluorescence polarization: nuclear receptor-ligand-
binding and kinase/phosphatase assays. J Biomol Screen 5, 77-88
155. Rishi, V., Potter, T., Laudeman, J., Reinhart, R., Silvers, T., Selby, M., Stevenson, T.,
Krosky, P., Stephen, A. G., Acharya, A., Moll, J., Oh, W. J., Scudiero, D., Shoemaker, R.
H., and Vinson, C. (2005) A high-throughput fluorescence-anisotropy screen that
identifies small molecule inhibitors of the DNA binding of B-ZIP transcription factors.
Anal Biochem 340, 259-271
156. Wang, S. Y., Ahn, B. S., Harris, R., Nordeen, S. K., and Shapiro, D. J. (2004)
Fluorescence anisotropy microplate assay for analysis of steroid receptor-DNA
interactions. Biotechniques 37, 807-808, 810-807
157. Millar, D. P. (2000) Time-resolved fluorescence methods for analysis of DNA-protein
interactions. Methods Enzymol 323, 442-459
158. McCauley, T. G., Hamaguchi, N., and Stanton, M. (2003) Aptamer-based biosensor
arrays for detection and quantification of biological macromolecules. Anal Biochem 319,
244-250
159. Nickoloff, J. A., and Hoekstra, M. F. (1998) DNA Damage and Repair: DNA Repair in
Prokaryotes and Lower Eukaryotes, Vol. 1. New York: Humana Press. 1st edition.
160. Ruscitti, T., Polayes, D. A., Karu, A. E., and Linn, S. (1992) Selective
immunoneutralization of the multiple activities of Escherichia coli DNA polymerase I
supports the model for separate active sites and indicates a complex 5' to 3' exonuclease.
J Biol Chem 267, 16806-16811
161. Stengel, G., Gill, J. P., Sandin, P., Wilhelmsson, L. M., Albinsson, B., Norden, B., and
Millar, D. (2007) Conformational dynamics of DNA polymerase probed with a novel
fluorescent DNA base analogue. Biochemistry 46, 12289-12297
137
162. DeLucia, A. M., Grindley, N. D., and Joyce, C. M. (2007) Conformational changes
during normal and error-prone incorporation of nucleotides by a Y-family DNA
polymerase detected by 2-aminopurine fluorescence. Biochemistry 46, 10790-10803
163. Wang, C. X., Zakharova, E., Li, J., Joyce, C. M., Wang, J., and Konigsberg, W. (2004)
Pre-steady-state kinetics of RB69 DNA polymerase and its exo domain mutants: effect of
pH and thiophosphoryl linkages on 3'-5' exonuclease activity. Biochemistry 43, 3853-
3861
164. Joyce, C. M., and Benkovic, S. J. (2004) DNA polymerase fidelity: kinetics, structure,
and checkpoints. Biochemistry 43, 14317-14324
165. Purohit, V., Grindley, N. D., and Joyce, C. M. (2003) Use of 2-aminopurine fluorescence
to examine conformational changes during nucleotide incorporation by DNA polymerase
I (Klenow fragment). Biochemistry 42, 10200-10211
166. Lam, W. C., Thompson, E. H., Potapova, O., Sun, X. C., Joyce, C. M., and Millar, D. P.
(2002) 3'-5' exonuclease of Klenow fragment: role of amino acid residues within the
single-stranded DNA binding region in exonucleolysis and duplex DNA melting.
Biochemistry 41, 3943-3951
167. LiCata, V. J., and Wowor, A. J. (2008) Applications of fluorescence anisotropy to the
study of protein-DNA interactions. Methods Cell Biol 84, 243-262
168. Minkley, E. G., Jr., Leney, A. T., Bodner, J. B., Panicker, M. M., and Brown, W. E.
(1984) Escherichia coli DNA polymerase I. Construction of a polA plasmid for
amplification and an improved purification scheme. J Biol Chem 259, 10386-10392
169. Joyce, C. M., and Derbyshire, V. (1995) Purification of Escherichia coli DNA
polymerase I and Klenow fragment. Methods Enzymol 262, 3-13
170. Inglese, J., Blatchly, R. A., and Benkovic, S. J. (1989) A multisubstrate adduct inhibitor
of a purine biosynthetic enzyme with a picomolar dissociation constant. J Med Chem 32,
937-940
171. Record, M. T., Jr., Zhang, W., and Anderson, C. F. (1998) Analysis of effects of salts and
uncharged solutes on protein and nucleic acid equilibria and processes: a practical guide
to recognizing and interpreting polyelectrolyte effects, Hofmeister effects, and osmotic
effects of salts. Adv Protein Chem 51, 281-353
172. Wyman, J., Jr. (1964) Linked Functions and Reciprocal Effects in Hemoglobin: A
Second Look. Adv Protein Chem 19, 223-286
173. Record, M. T., Jr., Ha, J. H., and Fisher, M. A. (1991) Analysis of equilibrium and
kinetic measurements to determine thermodynamic origins of stability and specificity and
mechanism of formation of site-specific complexes between proteins and helical DNA.
Methods Enzymol 208, 291-343
138
174. King, J. S., Fairley, C. F., and Morgan, W. F. (1996) DNA end joining by the Klenow
fragment of DNA polymerase I. J Biol Chem 271, 20450-20457
175. King, J., Fairley, C., and Morgan, W. (1998) The joining of blunt DNA ends to 3'-
protruding single strands in Escherichia coli. Nucleic Acids Res 26, 1749-1754
176. Baker, B. M., and Murphy, K. P. (1996) Evaluation of linked protonation effects in
protein binding reactions using isothermal titration calorimetry. Biophys J 71, 2049-2055
177. Englund, P. T., Huberman, J. A., Jovin, T. M., and Kornberg, A. (1969) Enzymatic
synthesis of deoxyribonucleic acid. XXX. Binding of triphosphates to deoxyribonucleic
acid polymerase. J Biol Chem 244, 3038-3044
178. Han, H., Rifkind, J. M., and Mildvan, A. S. (1991) Role of divalent cations in the 3',5'-
exonuclease reaction of DNA polymerase I. Biochemistry 30, 11104-11108
179. Slater, J. P., Tamir, I., Loeb, L. A., and Mildvan, A. S. (1972) The mechanism of
Escherichia coli deoxyribonucleic acid polymerase I. Magnetic resonance and kinetic
studies of the role of metals. J Biol Chem 247, 6784-6794
180. Wang, J., Sattar, A. K., Wang, C. C., Karam, J. D., Konigsberg, W. H., and Steitz, T. A.
(1997) Crystal structure of a pol alpha family replication DNA polymerase from
bacteriophage RB69. Cell 89, 1087-1099
181. Philo, J. (1994) Measuring sedimentation, diffusion, and molecular weights of small
molecules by direct fitting of sedimentation velocity concentration profiles, in Modern
Analytical Ultracentrifugation, edited by T.M. Schuster and T.M. Laue, pp. 156-170,
Birkhauser: Boston.
182. Philo, J. S. (1997) An improved function for fitting sedimentation velocity data for low-
molecular-weight solutes. Biophys J 72, 435-444
183. Millero, F. J., Ward, G. K., and Chetirkin, P. (1976) Partial specific volume,
expansibility, compressibility, and heat capacity of aqueous lysozyme solutions. J Biol
Chem 251, 4001-4004
184. Gekko, K., Kimoto, A., and Kamiyama, T. (2003) Effects of disulfide bonds on
compactness of protein molecules revealed by volume, compressibility, and expansibility
changes during reduction. Biochemistry 42, 13746-13753
185. Cantor, C., and Schimmel, P.R. (1980) Biophysical Chemistry, W.H. Freeman: San
Francisco.
186. Edelstein, S. J., and Schachman, H. K. (1967) The simultaneous determination of partial
specific volumes and molecular weights with microgram quantities. J Biol Chem 242,
306-311
187. Svedberg, T., Pederson, K.O. and Bauer, J.H. (1959) The Ultracentrifuge, Johnson
Reprint Corp.: New York.
139
188. van Holde, K. E., Johnson, W.C. and Ho, P.S. (1998) Principles of Physical
Biochemistry, Prentice Hall: Upper Saddle River, pp. 192-212.
189. Zhao, J., Wang, J., Chen, D. J., Peterson, S. R., and Trewhella, J. (1999) The solution
structure of the DNA double-stranded break repair protein Ku and its complex with
DNA: a neutron contrast variation study. Biochemistry 38, 2152-2159
190. Osterberg, R., Sjoberg, B., Rymo, L., and Lagerkvist, U. (1975) Small-angle x-ray
scattering study of the interaction between lysine transfer RNA ligase from yeast and
transfer RNA. J Mol Biol 99, 383-393
191. McAfee, J. G., Edmondson, S. P., Zegar, I., and Shriver, J. W. (1996) Equilibrium DNA
binding of Sac7d protein from the hyperthermophile Sulfolobus acidocaldarius:
fluorescence and circular dichroism studies. Biochemistry 35, 4034-4045
192. Woody, R. W. (1995) Circular dichroism. Methods Enzymol 246, 34-71
193. Heyduk, E., Baichoo, N., and Heyduk, T. (2001) Interaction of the alpha-subunit of
Escherichia coli RNA polymerase with DNA: rigid body nature of the protein-DNA
contact. J Biol Chem 276, 44598-44603
194. Selivanova, O. M., Shiryaev, V. M., Tiktopulo, E. I., Potekhin, S. A., and Spirin, A. S.
(2003) Compact globular structure of Thermus thermophilus ribosomal protein S1 in
solution: sedimentation and calorimetric study. J Biol Chem 278, 36311-36314
195. Datta, K. (2004) Thermodynamic Characterization of DNA Binding by Type I DNA
Polymerases from Thermus aquaticus and Escherichia coli. Diss. Louisiana State
University, Baton Rouge.
196. Iyer, R. R., Pluciennik, A., Burdett, V., and Modrich, P. L. (2006) DNA mismatch repair:
functions and mechanisms. Chem Rev 106, 302-323
197. Georgiou, C. D., and Papapostolou, I. (2006) Assay for the quantification of
intact/fragmented genomic DNA. Anal Biochem 358, 247-256
198. Nguyen, L. T., Wiencek, J. M., and Kirsch, L. E. (2003) Characterization methods for the
physical stability of biopharmaceuticals. PDA J Pharm Sci Technol 57, 429-445
199. Banachowicz, E. (2006) Light scattering studies of proteins under compression. Biochim
Biophys Acta 1764, 405-413
200. Domingues, M. M., Santiago, P. S., Castanho, M. A., and Santos, N. C. (2008) What can
light scattering spectroscopy do for membrane-active peptide studies? J Pept Sci 14, 394-
400
201. Hochstrasser, R. A., Carver, T. E., Sowers, L. C., and Millar, D. P. (1994) Melting of a
DNA helix terminus within the active site of a DNA polymerase. Biochemistry 33,
11971-11979
140
202. Ramreddy, T., Rao, B. J., and Krishnamoorthy, G. (2007) Site-specific dynamics of
strands in ss- and dsDNA as revealed by time-domain fluorescence of 2-aminopurine. J
Phys Chem B 111, 5757-5766
203. Tleugabulova, D., and Reha-Krantz, L. J. (2007) Probing DNA polymerase-DNA
interactions: examining the template strand in exonuclease complexes using 2-
aminopurine fluorescence and acrylamide quenching. Biochemistry 46, 6559-6569
204. Ballin, J. D., Prevas, J. P., Bharill, S., Gryczynski, I., Gryczynski, Z., and Wilson, G. M.
(2008) Local RNA conformational dynamics revealed by 2-aminopurine solvent
accessibility. Biochemistry 47, 7043-7052
205. Tang, G. Q., Paratkar, S., and Patel, S. S. (2009) Fluorescence mapping of the open
complex of yeast mitochondrial RNA polymerase. J Biol Chem 284, 5514-5522
141
APPENDIX
ELSEVIER LICENSE
TERMS AND CONDITIONS
Feb 13, 2009
This is a License Agreement between Andy J Wowor (“You”) and Elsevier (“Elsevier”) provided
by Copyright Clearance Center (“CCC”). The license consists of your order details, the terms and
conditions provided by Elsevier, and the payment terms and conditions.
All payments must be made in full to CCC. For payment instructions, please see
information listed at the bottom of this form.
Supplier
Elsevier Limited
The Boulevard,Langford Lane
Kidlington,Oxford,OX5 1GB,UK
Registered Company Number 1982084
Customer name Andy J Wowor
Customer address 107 Life Science Bldg.
Baton Rouge, LA 70803
License Number 2123171095025
License date Feb 06, 2009
Licensed Chapter Title Applications of Fluorescence Anisotropy to the Study of Protein–
DNA Interactions
Licensed Chapter ID S0091679X0784009X
Licensed content publisher Elsevier
Licensed content publication Elsevier Books
Book title Methods in Cell Biology, Volume 84
Book author Vince J. LiCata and Andy J. Wowor
Publication date 2008
Pages 20
Type of Use Thesis / Dissertation
Portion Full chapter
Format Both print and electronic
You are an author of the
Elsevier book Yes
Are you translating? No
Purchase order number
Expected publication date Mar 2009
Elsevier VAT number GB 494 6272 12
Permissions price 0.00 USD
142
Value added tax 0.0% 0.00 USD
Total 0.00 USD
Terms and Conditions
INTRODUCTION
1. The publisher for this copyrighted material is Elsevier. By clicking "accept" in connection
with completing this licensing transaction, you agree that the following terms and conditions
apply to this transaction (along with the Billing and Payment terms and conditions established by
Copyright Clearance Center, Inc. ("CCC"), at the time that you opened your Rightslink account
and that are available at any time at http://myaccount.copyright.com).
GENERAL TERMS
2. Elsevier hereby grants you permission to reproduce the aforementioned material subject to the
terms and conditions indicated.
3. Acknowledgement: If any part of the material to be used (for example, figures) has appeared
in our publication with credit or acknowledgement to another source, permission must also be
sought from that source. If such permission is not obtained then that material may not be
included in your publication/copies. Suitable acknowledgement to the source must be made,
either as a footnote or in a reference list at the end of your publication, as follows:
“Reprinted from Publication title, Vol /edition number, Author(s), Title of article / title of
chapter, Pages No., Copyright (Year), with permission from Elsevier [OR APPLICABLE
SOCIETY COPYRIGHT OWNER].” Also Lancet special credit - “Reprinted from The Lancet,
Vol. number, Author(s), Title of article, Pages No., Copyright (Year), with permission from
Elsevier.”
4. Reproduction of this material is confined to the purpose and/or media for which permission is
hereby given.
5. Altering/Modifying Material: Not Permitted. However figures and illustrations may be
altered/adapted minimally to serve your work. Any other abbreviations, additions, deletions
and/or any other alterations shall be made only with prior written authorization of Elsevier Ltd.
(Please contact Elsevier at [email protected])
6. If the permission fee for the requested use of our material is waived in this instance, please be
advised that your future requests for Elsevier materials may attract a fee.
7. Reservation of Rights: Publisher reserves all rights not specifically granted in the combination
of (i) the license details provided by you and accepted in the course of this licensing transaction,
(ii) these terms and conditions and (iii) CCC's Billing and Payment terms and conditions.
8. License Contingent Upon Payment: While you may exercise the rights licensed immediately
upon issuance of the license at the end of the licensing process for the transaction, provided that
143
you have disclosed complete and accurate details of your proposed use, no license is finally
effective unless and until full payment is received from you (either by publisher or by CCC) as
provided in CCC's Billing and Payment terms and conditions. If full payment is not received on
a timely basis, then any license preliminarily granted shall be deemed automatically revoked and
shall be void as if never granted. Further, in the event that you breach any of these terms and
conditions or any of CCC's Billing and Payment terms and conditions, the license is
automatically revoked and shall be void as if never granted. Use of materials as described in a
revoked license, as well as any use of the materials beyond the scope of an unrevoked license,
may constitute copyright infringement and publisher reserves the right to take any and all action
to protect its copyright in the materials.
9. Warranties: Publisher makes no representations or warranties with respect to the licensed
material.
10. Indemnity: You hereby indemnify and agree to hold harmless publisher and CCC, and their
respective officers, directors, employees and agents, from and against any and all claims arising
out of your use of the licensed material other than as specifically authorized pursuant to this
license.
11. No Transfer of License: This license is personal to you and may not be sublicensed, assigned,
or transferred by you to any other person without publisher's written permission.
12. No Amendment Except in Writing: This license may not be amended except in a writing
signed by both parties (or, in the case of publisher, by CCC on publisher's behalf).
13. Objection to Contrary Terms: Publisher hereby objects to any terms contained in any
purchase order, acknowledgment, check endorsement or other writing prepared by you, which
terms are inconsistent with these terms and conditions or CCC's Billing and Payment terms and
conditions. These terms and conditions, together with CCC's Billing and Payment terms and
conditions (which are incorporated herein), comprise the entire agreement between you and
publisher (and CCC) concerning this licensing transaction. In the event of any conflict between
your obligations established by these terms and conditions and those established by CCC's
Billing and Payment terms and conditions, these terms and conditions shall control.
14. Revocation: Elsevier or Copyright Clearance Center may deny the permissions described in
this License at their sole discretion, for any reason or no reason, with a full refund payable to
you. Notice of such denial will be made using the contact information provided by you. Failure
to receive such notice will not alter or invalidate the denial. In no event will Elsevier or
Copyright Clearance Center be responsible or liable for any costs, expenses or damage incurred
by you as a result of a denial of your permission request, other than a refund of the amount(s)
paid by you to Elsevier and/or Copyright Clearance Center for denied permissions.
LIMITED LICENSE
The following terms and conditions apply to specific license types:
15. Translation: This permission is granted for non-exclusive world English rights only unless
144
your license was granted for translation rights. If you licensed translation rights you may only
translate this content into the languages you requested. A professional translator must perform all
translations and reproduce the content word for word preserving the integrity of the article. If this
license is to re-use 1 or 2 figures then permission is granted for non-exclusive world rights in all
languages.
16. Website: The following terms and conditions apply to electronic reserve and author websites:
Electronic reserve: If licensed material is to be posted to website, the web site is to be
password-protected and made available only to bona fide students registered on a relevant course
if:
This license was made in connection with a course,
This permission is granted for 1 year only. You may obtain a license for future website posting,
All content posted to the web site must maintain the copyright information line on the bottom of
each image,
A hyper-text must be included to the Homepage of the journal from which you are licensing at
http://www.sciencedirect.com/science/journal/xxxxx or, for books, to the Elsevier homepage at
http://www.elsevier.com,
Central Storage: This license does not include permission for a scanned version of the material to
be stored in a central repository such as that provided by Heron/XanEdu.
17. Author website for journals with the following additional clauses:
All content posted to the web site must maintain the copyright information line on the bottom of
each image, and
The permission granted is limited to the personal version of your paper. You are not allowed to
download and post the published electronic version of your article (whether PDF or HTML,
proof or final version), nor may you scan the printed edition to create an electronic version,
A hyper-text must be included to the Homepage of the journal from which you are licensing at
http://www.sciencedirect.com/science/journal/xxxxx,
Central Storage: This license does not include permission for a scanned version of the material to
be stored in a central repository such as that provided by Heron/XanEdu.
18. Author website for books with the following additional clauses:
Authors are permitted to place a brief summary of their work online only.
A hyper-text must be included to the Elsevier homepage at http://www.elsevier.com.
All content posted to the web site must maintain the copyright information line on the bottom of
each image
You are not allowed to download and post the published electronic version of your chapter, nor
may you scan the printed edition to create an electronic version.
Central Storage: This license does not include permission for a scanned version of the material to
be stored in a central repository such as that provided by Heron/XanEdu.
19. Website (regular and for author): A hyper-text must be included to the Homepage of the
journal from which you are licensing at http://www.sciencedirect.com/science/journal/xxxxx or,
for books, to the Elsevier homepage at http://www.elsevier.com.
20. Thesis/Dissertation: If your license is for use in a thesis/dissertation your thesis may be
145
submitted to your institution in either print or electronic form. Should your thesis be published
commercially, please reapply for permission. These requirements include permission for the
Library and Archives of Canada to supply single copies, on demand, of the complete thesis and
include permission for UMI to supply single copies, on demand, of the complete thesis. Should
your thesis be published commercially, please reapply for permission.
21. Other conditions: None
v1.5
Gratis licenses (referencing $0 in the Total field) are free. Please retain this printable
license for your reference. No payment is required.
If you would like to pay for this license now, please remit this license along with your
payment made payable to "COPYRIGHT CLEARANCE CENTER" otherwise you will be
invoiced within 30 days of the license date. Payment should be in the form of a check or
money order referencing your account number and this license number 2123171095025.
If you would prefer to pay for this license by credit card, please go to
http://www.copyright.com/creditcard to download our credit card payment authorization
form.
Make Payment To:
Copyright Clearance Center
Dept 001
P.O. Box 843006
Boston, MA 02284-3006
If you find copyrighted material related to this license will not be used and wish to cancel,
please contact us referencing this license number 2123171095025 and noting the reason for
cancellation.
Questions? [email protected] or 877-622-5543 or +1-978-646-2777.
146
Dear Mr Wowor
We hereby grant you permission to reprint the material detailed below at no charge in your thesis
subject to the following conditions:
1. If any part of the material to be used (for example, figures) has appeared in our publication
with credit or acknowledgement to another source, permission must also be sought from that
source. If such permission is not obtained then that material may not be included in your
publication/copies.
2. Suitable acknowledgment to the source must be made, either as a footnote or in a reference
list at the end of your publication, as follows:
“This article was published in Publication title, Vol number, Author(s), Title of article, Page
Nos, Copyright Elsevier (or appropriate Society name) (Year).”
3. Your thesis may be submitted to your institution in either print or electronic form.
4. Reproduction of this material is confined to the purpose for which permission is hereby
given.
5. This permission is granted for non-exclusive world English rights only. For other
languages please reapply separately for each one required. Permission excludes use in an
electronic form. Should you have a specific electronic project in mind please reapply for
permission.
6. This includes permission for UMI to supply single copies, on demand, of the complete
thesis. Should your thesis be published commercially, please reapply for permission.
Yours sincerely
______________________________________________
Steph Smith :: Global Rights :: Elsevier
T: +44 (0)1865 843325 :: F: +44 (0)1865 853333
-----Original Message-----
From: [email protected] [mailto:[email protected]]
Sent: 06 February 2009 20:11
To: Rights and Permissions (ELS)
Subject: Obtain Permission
This Email was sent from the Elsevier Corporate Web Site
and is related to Obtain Permission form:
----------------------------------------------------------------
Product: Customer Support
Component: Obtain Permission
Web server: http://www.elsevier.com
IP address: 195.212.150.72
Client: Mozilla/4.0 (compatible; MSIE 7.0; Windows NT 5.1; .NET CLR 1.1.4322; .NET
CLR 2.0.50727; .NET CLR 3.0.04506.30; InfoPath.2)
147
Invoked from:
http://www.elsevier.com/wps/find/obtainpermissionform.cws_home?isSubmitted=yes&navigate
XmlFileName=/store/scstargets/prd53/act/framework_support/obtainpermission.xml
Request From:
Andy Wowor
Louisiana State University
107 Life Science Bldg.
70803
Baton Rouge
United States
Contact Details:
Telephone: 2255781589
Fax:
Email Address: [email protected]
To use the following material:
ISSN/ISBN:
Title: Biophysical Journal
Author(s): Datta, Wowor, Richard, and LiCata
Volume: 90
Issue: 5
Year: 2006
Pages: 1739 - 1751
Article title: Temperature Dependence and Thermodynamics
How much of the requested material is to be used:
Certain Pages
Are you the author: Yes
Author at institute: Yes
How/where will the requested material be used: [how_used]
Details:
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
Additional Info: The authors' names are Kausiki Datta, Andy J. Wowor, Allison J. Richard,
and Vince J. LiCata.
The full title of the article is "Temperature Dependence and Thermodynamics of Klenow
Polymerase Binding to Primed-Template DNA." Thank you.
[acronym]
- end -
Elsevier Limited. Registered Office: The Boulevard, Langford Lane, Kidlington, Oxford, OX5
1GB, United Kingdom, Registration No. 1982084 (England and Wales).
148
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.