Investigation of Noncanonical DNA Polymerases and Their Mechanisms
Dissertation
Presented in Partial Fulfillment for the Degree Doctor of Philosophy
in the Graduate School of The Ohio State University
By
Jason David Fowler
Ohio State Biochemistry Program
The Ohio State University
2009
Dissertation Committee:
Zucai Suo, Advisor
Venkat Gopalan
Mamuka Kvaratskhelia
Chenglong Li
Copyright by
Jason David Fowler
2009
Abstract
DNA polymerases have evolved complex biological functions to balance the need
for faithful DNA replication with the need for subtle genetic flexibility in the form of
random mutations that are necessary for sustaining life in an ever-changing environment.
A more fundamental understanding of the diverse mechanisms that lie at the heart of
DNA polymerases is the ultimate goal of the ongoing research presented here.
Gemcitabine, 2'-deoxy-2', 2'-difluorocytidine (dFdC), is a drug approved for use
against various solid tumors. Clinically, this moderately toxic nucleoside analog causes
side effects which closely mimic symptoms of mitochondrial dysfunction, although there
is no direct evidence to show gemcitabine interferes with mitochondrial DNA replication
catalyzed by human DNA polymerase gamma. Here we employed pre-steady state kinetic
methods to directly investigate the incorporation of the 5'-triphosphorylated form of
gemcitabine (dFdCTP), the excision of the incorporated monophosphorylated form
(dFdCMP), and the bypass of template base dFdC catalyzed by human DNA polymerase
gamma. Opposite template base dG, dFdCTP was incorporated with a 432-fold lower
efficiency than dCTP. Although dFdC is not a chain terminator, the incorporated
dFdCMP decreased the incorporation efficiency of the next 2 correct nucleotides by 214-
ii
and 7-fold, respectively. Moreover, the primer 3'-dFdCMP was excised with a 50-fold
slower rate than the matched 3'-dCMP. When dFdC was encountered as a template base,
DNA polymerase gamma paused at the lesion and one downstream position but
eventually elongated the primer to full-length product. These pauses were because of a
1,000-fold decrease in nucleotide incorporation efficiency. Interestingly, the polymerase
fidelity at these pause sites decreased by 2 orders of magnitude. Thus, our pre-steady
state kinetic studies provide direct evidence demonstrating the inhibitory effect of
gemcitabine on the activity of human mitochondrial DNA polymerase.
Crystallographic studies of the truncated C-terminal DNA polymerase-beta-like
domain of human DNA polymerase lambda (tPollambda) suggested the catalytic cycle
might not involve a large protein domain rearrangement as observed with replicative
DNA polymerases and DNA polymerase beta. To examine solution-phase protein
conformational changes in full length DNA polymerase lambda (fpollambda), which
contains two additional domains at its N-terminus, we used a mass spectrometry-based
protein footprinting approach. In parallel experiments, surface accessibility maps for Arg
residues were compared for the free fPollambda versus the binary complex of
enzyme*gapped DNA and the ternary complex of enzyme*gapped DNA*dNTP (2'deoxynucleotide triphosphate). These experiments suggested that fPollambda does not
undergo major conformational changes during the catalysis in the solution phase.
iii
Furthermore, the mass spectrometry-based protein footprinting experiments revealed that
active site residue R386 was shielded from the surface only in the presence of both a
gapped DNA substrate and an incoming nucleotide. Site-directed mutagenesis and presteady-state kinetic studies confirmed the importance of R386 for the enzyme activity and
indicated the key role for its guanidino group in stabilizing the negative charges of an
incoming nucleotide and the leaving pyrophosphate product. We suggest that such
interactions could be shared by and important for catalytic functions of other DNA
polymerases.
iv
Acknowledgments
I would first like to acknowledge my advisor, Dr. Zucai Suo. Without his tireless
guidance, I would never have been able to realize my scientific potential. Dr. Suo has
shown me the value of relentless pursuit of intelectual excellence, and what one can
accomplish at the intersection of intellegence and dicipline. I would also like to sincerely
thank my comittee member Dr. Mamuka Kvaratskhelia for his patient and selfless
assistance, without which much of the work presented herein would never have been
possible. In addition, I would like to thank Dr. Venkat Gopalan for providing me with
both a wealth of invaluable advice and a role model for what a leader in the scientific
community should and must be. Also, for his assistance as a member of my graduate
comitte and for teaching me the art and valuable science of protein crystallography, I
would like to extend my sincere thanks to Dr. Chenglong Li. Furthermore, I would like to
acknowledge Jessica Brown for her assistance with many of my projects and her
generous efforts on my behalf, which more than once made the difference between a
dataset and a scientific publication. Also, I must sincerely thank the numerous other
current and former members of our lab (too numerous to mention in their entirety but
especially: Dr. Kevin Fiala, Shanen Sherrer, Dr. Cuiling Xu, John Pryor, Lindsey Pack,
Carlo Dela Seña, Nikunj Bhatt and many others), who over these difficult years have
v
provided me with the support, assistance and friendship that in my darkest hours, made
all the difference.
Finally, I would also like to acknowledge the generous support of the Ohio State
Biochemistry Program and the American Heart Association whose financial fellowships
provided funding for my graduate career.
vi
Vita
Education
1997 – 2001…………A.S. Biology
Columbus State Community College, Columbus, Ohio
2001 – 2003…………B.S. Biochemistry
The Ohio State University, Columbus, Ohio
2003 – Present………Graduate Teaching and Research Associate
College of Biological Sciences, the Ohio State University
Awards and Honors
2001…………………Cum Laude graduation honors from Columbus State Community
College
2006…………………Nominated for the Ohio State University Graduate Teaching
Award
2006…………………Awarded the Ohio State University College of Biological Sciences
Dean’s Award for Excellence as a Teaching Assistant
2007…………………Award for Outstanding Poster Presentation at the Ohio State
University Molecular Life Sciences Interdisciplinary Graduate
Programs Symposium
Fellowships
2006…………………Awarded the American Heart Association Pre-Doctoral Fellowship
for academic years 2006-2007 and 2007-2008
Conferences and Presentations
2007…………………Attended the 9th annual Midwest DNA Repair Symposium
2007…………………Presented a Poster At: The 76th annual Gordon Research
Conference
vii
Conferences and Presentations (continued)
2007…………………Presented a Poster At: The Ohio State University Comprehensive
Cancer Center 9th Annual Scientific Meeting
2007…………………Presented a Poster At: The Ohio State University Molecular Life
Sciences Interdisciplinary Graduate Programs Symposium
2008…………………Presented a Poster At: The Ohio State University Molecular Life
Sciences Interdisciplinary Graduate Programs Symposium
Professional Membership
2005-Present
American Association for the Advancement of Science (AAAS)
Publications
1.
2.
3.
4.
5.
Fowler, J. & Suo, Z. (2006) Biochemical, Structural, and Physiological
Characterization of Terminal Deoxynucleotidyl Transferase. Chemical Reviews
106, 2092-2110.
Brown, J. A., Duym, W. W., Fowler, J. D., and Suo, Z. (2007) Single-Turnover
Kinetic Analysis of the Mutagenic Potential of 7,8-Dihydro-8-oxoguanine During
Gap-Filling Synthesis Catalyzed by Human DNA Polymerases and β, J. Mol.
Biol. 367, 1258-1269.
Fowler, J. D., Brown, J.A., Johnson, K.A. and Suo, Z. (2008) Kinetic
Investigation of the Inhibitory Effect of Gemcitabine on DNA Polymerization
Catalyzed by Human Mitochondrial DNA Polymerase. J. Biol. Chem. 283,
15339-15348.
Sherrer, S. M., Brown, J.A., Pack, L. R., Fowler, J. D., Basu, A. K. and Suo, Z.
(2009) Mechanistic studies of the bypass of a bulky single-base lesion catalyzed
by a Y-family DNA polymerase. J. Biol. Chem. 284, 6379-6388.
Fowler, J.D., Brown, J. A., Kvaratskhelia, M., Suo, Z. (2009) Probing
conformational changes of human DNA polymerase lambda using mass
spectrometry-based protein footprinting. J Mol Biol, 2009. 390(3): p. 368-79.
Fields of Study
Major Field: Ohio State Biochemistry Program
viii
Table of Contents
Abstract ............................................................................................................................. ii
Acknowledgments..............................................................................................................v
Vita.................................................................................................................................. vii
List of Tables ...................................................................................................................xv
List of Figures ................................................................................................................ xvi
Abbreviations ............................................................................................................... xviii
Chapter 1: DNA polymerases and terminal deoxynucleotidyl transferase ........................1
1.1. Introduction ...........................................................................................................1
1.2. Classification of DNA Polymerases .....................................................................2
Family A ................................................................................................................2
Family B.................................................................................................................3
Family C.................................................................................................................4
Family D ................................................................................................................4
Family Y ................................................................................................................5
Family X ................................................................................................................6
1.3. Sequence Alignment Analysis of the X-family DNA Polymerases ...................10
1.4. Isoforms of TdT ..................................................................................................12
1.5. TdT Expression and Purification ........................................................................14
1.6. Three-Dimensional Structures of DNA Polymerases .........................................15
Crystal Structures of Template-Dependant DNA Polymerases ...........................15
Crystal Structure of the Polβ-Like Domain of Murine TdT ................................16
1.7. Enzymatic Activities of TdT ...............................................................................21
Template-Independent Polymerase Activity........................................................21
dRPase-Deficiency in TdT ...................................................................................21
Primer Requirement .............................................................................................22
Metal Ion Dependence .........................................................................................23
Kinetic Mechanism of Template-Independent Polymerization ...........................24
Nucleotide Selectivity ..........................................................................................26
ix
Other Enzymatic Activities ..................................................................................27
1.8. Immune System and TdT ....................................................................................29
V(D)J Recombination ..........................................................................................32
Role of TdT in V(D)J Recombination .................................................................33
TdT Regulation ....................................................................................................37
Somatic Hypermutation .......................................................................................40
1.9. Experience Dependence Memory Processing and TdT ......................................44
1.10. Figures...............................................................................................................46
Figure 1.1 Domain organization of six X-family DNA polymerases. .................46
Figure 1.2 Ternary structure of human DNA polymerase β•single nucleotide
gapped DNA•ddCTP......................................................................................47
Figure 1.3 Proposed “two-divalent-metal-ion” mechanism for nucleotide
incorporation catalyzed by human DNA polymerase β .................................48
Figure 1.4 Binary crystal structures of the Polβ-like domain (residues 148-510) of
murine TdT complexed with a brominated 9-mer at 3.0 Å ...........................49
Figure 1.5 Binary crystal structure of the Polβ-like domain (residues 148-510) of
murine TdT complexed with a ddATP-Co2+ at 3.0 Å ..................................51
Figure 1.6 Minimal kinetic mechanism for polymerization catalyzed by DNA
polymerases....................................................................................................52
Figure 1.7 Chemical structures of nucleotide analogs .........................................53
Figure 1.8 Crystal structure of anti-lysozyme Fab and hen egg white lysozyme 54
Figure 1.9 T cell receptor encoded by tandemly arranged clusters of V, D, and J
gene segments. ...............................................................................................56
Figure 1.10 V(D)J Recombination .......................................................................58
Figure 1.11 Proposed mechanism for the “N region” formation at the junction
between a V and a D segment. .......................................................................60
1.11. Tables ................................................................................................................61
Chapter 2: Kinetic investigation of the inhibitory effect of gemcitabine on DNA
polymerization catalyzed by human mitochondrial DNA polymerase ......................62
2.1. Introduction .........................................................................................................62
2.2. Materials .............................................................................................................66
x
Optimized Reaction Buffer G ..............................................................................66
Optimized Reaction Buffer L...............................................................................66
Optimized Reaction Buffer M .............................................................................66
Purification of Human Polymerase Gamma Subunits .........................................67
Synthetic Oligodeoxyribonucleotides ..................................................................67
Synthetic Oligodeoxyribonucleotides Containing Gemcitabine ..........................67
2.3. Methods...............................................................................................................69
Single-Turnover Nucleotide Incorporation Assay ...............................................69
Excision Reactions ...............................................................................................69
Running Start Nucleotide Incorporation Assay ...................................................70
Product Analysis ..................................................................................................70
Data Analysis .......................................................................................................70
2.4. Results .................................................................................................................72
Determination of the Pre-Steady State Kinetic Parameters for dFdCTP and dCTP
Incorporation ..................................................................................................72
Measurement of the Excision Rate Constants of Matched 3’-dFdCMP and 3’dCMP .............................................................................................................74
Measurement of the Extension Efficiency of a Primer Terminated with 3’dFdCMP .........................................................................................................75
Running Start Primer Extension Assays ..............................................................76
Measurement of the Excision Rate Constant of Primer 3’-dNMP Opposite
Template Base dFdC ......................................................................................78
Measurement of Incorporation Efficiency of Nucleotides Opposite Template
dFdC ...............................................................................................................78
Measurement of Nucleotide Incorporation Fidelity at the Pause Sites ................80
2.5. Discussion ...........................................................................................................82
Inhibition of Mitochondrial DNA Synthesis by Gemcitabine as an Incoming
Nucleotide ......................................................................................................82
Incorporated dFdCMP Eludes Editing Mechanism .............................................84
Inhibition of Mitochondrial DNA Synthesis by Gemcitabine as a Template Base
........................................................................................................................85
Unfaithful Bypass of Template dFdCMP ............................................................87
xi
Pathologic Effects Associated with Gemcitabine Therapy ..................................88
2.6. Figures.................................................................................................................91
Figure 2.1 Chemical Structure of Gemcitabine and 2'-Deoxycytidine. ...............91
Figure 2.2 Gemcitabine Activation and Self Potentiation Pathways. ..................92
Figure 2.3 Pre-steady state kinetic analysis of Polγ .............................................93
Figure 2.4 Measurement of the Rate Constant of DNA Primer Degradation by the
3' 5' Exonuclease Proofreading Activity of the Wild-type Polγ ................95
Figure 2.5 Running Start Primer Elongation Catalyzed by the Wild-Type Polγ and
the Exonuclease Deficient Mutant E200A. ....................................................96
Figure 2.6 Sequencing gel image of single nucleotide incorporation catalyzed by
Polγ mutant E200A ........................................................................................98
2.7. Tables ................................................................................................................100
Chapter 3: Probing protein conformational changes of a human DNA polymerase using
mass spectrometry....................................................................................................106
3.1. Introduction .......................................................................................................106
3.2. Materials ...........................................................................................................109
Preparation of Human fPolλ, dPolλ, and tPolλ ..................................................109
Synthetic Oligodeoxyribonucleotides ................................................................109
Reaction Buffer ..................................................................................................110
3.3. Methods.............................................................................................................111
Mass Spectrometry-Based Protein Footprinting Assay .....................................111
Gap-Filling DNA Polymerase Activity Assay for HPG-Modified Enzymes ....112
Determination of kp and Kd Values ....................................................................113
3.4. Results ...............................................................................................................114
Investigating the stability of human Polλ during HPG modification ................115
MS-Based Footprinting of fPolλ, dPolλ, and tPolλ ...........................................116
Pre-steady state kinetic analysis of two R386 mutants of fPolλ ........................118
3.5. Discussion .........................................................................................................120
Structural Implications of Our MS-Based Protein Footprinting Data ...............120
Structural and Functional Roles of R386 ...........................................................122
xii
Conservation of R386 and R420 in Other DNA Polymerases ...........................123
3.6. Figures...............................................................................................................126
Figure 3.1 Domain structure of human fPolλ, dPolλ, tPolλ, and Polβ...............126
Figure 3.2 Gap-filling DNA polymerase activity of fPolλ following HPG
modification. ................................................................................................127
Figure 3.3 Representative segments of the MALDI-ToF MS spectra. ..............128
Figure 3.4 Tryptic digestion map of human fPolλ. ............................................130
Figure 3.5 Crystal structure of tPolλ detailing the interactions of R386, R275,
ddTTP, and the DNA template. ...................................................................131
Figure 3.6 Concentration dependence on the rate of dTTP incorporation into 2119/41A-mer ..................................................................................................133
Figure 3.7 Arginine residues modified by HPG in the crystal structure of the
ternary complex of tPolλ. .............................................................................135
Figure 3.8 Active site of tPolλ. ..........................................................................136
Figure 3.9 Y-family DNA polymerase sequence alignment. .............................137
3.7. Tables ................................................................................................................139
Chapter 4 - Preliminary investigation of the mechanism of Y-family DNA polymerases
using mass spectrometry ..........................................................................................144
4.1. Introduction .......................................................................................................144
4.2. Materials ...........................................................................................................146
Preparation of Dpo4 ...........................................................................................146
Synthetic oligodeoxyribonucleotides .................................................................146
Reaction buffer D ...............................................................................................146
4.3. Methods.............................................................................................................147
Mass spectrometry-based protein footprinting assay .........................................147
4.4. Preliminary results ............................................................................................149
4.5. Figures...............................................................................................................151
Figure 4.1 Crystal structure of Dpo4 showing K282 .........................................151
Figure 4.2 Crystal structure of Dpo4 in the binary complex .............................152
Figure 4.3 Crystal structure of Dpo4 in the ternary complex ............................153
xiii
4.6. Tables ................................................................................................................154
References ......................................................................................................................155
xiv
List of Tables
Table 1.1 Effect of metal ions on the incorporation rate of each dNTP catalyzed
by TdT. ...................................................................................................................61
Table 2.1 DNA Substrates ...............................................................................................100
Table 2.2 Kinetic Parameters of Single Nucleotide Incorporation Catalyzed by
Polγ E200A under Single-Turnover Conditions at 37 C ....................................102
Table 2.3 Excision Rate Constants for the 3' 5' Exonuclease Activity of the
Wild-Type Human Polγ Holoenzyme under Single-Turnover Conditions at
37 C ....................................................................................................................103
Table 2.4 Kinetic Parameters of Single Nucleotide Incorporation into DNA
Containing a Template Base dFdCMP Catalyzed by Polγ E200A under
Single-Turnover Conditions at 37 C ..................................................................104
Table 2.5 Fidelity at the Two Strong Pause Sites ............................................................105
Table 3.1 DNA substrates. ...............................................................................................139
Table 3.2 Summary of modified arginine residues ..........................................................140
Table 3.3 Kinetic parameters of dTTP incorporation into single-nucleotide gapped
21-19/41A-mer catalyzed by fPolλ variants at 37 C ..........................................141
Table 3.4 Positively-charged residues that potentially stabilize the triphosphate
moiety of an incoming nucleotide and/or pyrophosphate product.......................142
Table 4.1 Summary of modified lysine residues in Dpo4. ..............................................154
xv
List of Figures
Figure 1.1 Domain organization of six X-family DNA polymerases. ...............................46
Figure 1.2 Ternary structure of human DNA polymerase β•single nucleotide
gapped DNA•ddCTP..............................................................................................47
Figure 1.3 Proposed “two-divalent-metal-ion” mechanism for nucleotide
incorporation catalyzed by human DNA polymerase β .........................................48
Figure 1.4 Binary crystal structures of the Polβ-like domain (residues 148-510) of
murine TdT complexed with a brominated 9-mer at 3.0 Å. ..................................49
Figure 1.5 Binary crystal structure of the Polβ-like domain (residues 148-510) of
murine TdT complexed with a ddATP-Co2+ at 3.0 Å. .........................................51
Figure 1.6 Minimal kinetic mechanism for polymerization catalyzed by DNA
polymerases............................................................................................................52
Figure 1.7 Chemical structures of nucleotide analogs .......................................................53
Figure 1.8 Crystal structure of anti-lysozyme Fab and hen egg white lysozyme ..............54
Figure 1.9 T cell receptor encoded by tandemly arranged clusters of V, D, and J
gene segments. .......................................................................................................56
Figure 1.10 V(D)J Recombination.....................................................................................58
Figure 1.11 Proposed mechanism for the “N region” formation at the junction
between a V and a D segment. ...............................................................................60
Figure 2.1 Chemical Structure of Gemcitabine and 2'-Deoxycytidine. .............................91
Figure 2.2 Gemcitabine Activation and Self Potentiation Pathways. ................................92
Figure 2.3 Pre-steady state kinetic analysis of Polγ ...........................................................93
Figure 2.4 Measurement of the Rate Constant of DNA Primer Degradation by the
3' 5' Exonuclease Proofreading Activity of the Wild-type Polγ ........................95
Figure 2.5 Running Start Primer Elongation Catalyzed by the Wild-Type Polγ and
the Exonuclease Deficient Mutant E200A.............................................................96
Figure 2.6 Sequencing gel image of single nucleotide incorporation catalyzed by
Polγ mutant E200A ................................................................................................98
Figure 3.1 Domain structure of human fPolλ, dPolλ, tPolλ, and Polβ.............................126
Figure 3.2 Gap-filling DNA polymerase activity of fPolλ following HPG
modification. ........................................................................................................127
Figure 3.3 Representative segments of the MALDI-ToF MS spectra. ............................128
Figure 3.4 Tryptic digestion map of human fPolλ. ..........................................................130
Figure 3.5 Crystal structure of tPolλ detailing the interactions of R386, R275,
ddTTP, and the DNA template. ...........................................................................131
xvi
Figure 3.6 Concentration dependence on the rate of dTTP incorporation into 2119/41A-mer ..........................................................................................................133
Figure 3.7 Arginine residues modified by HPG in the crystal structure of the
ternary complex of tPolλ. .....................................................................................135
Figure 3.8 Active site of tPolλ. ........................................................................................136
Figure 3.9 Y-family DNA polymerase sequence alignment. ...........................................137
Figure 4.1 Crystal structure of Dpo4 showing K282 .......................................................151
Figure 4.2 Crystal structure of Dpo4 in the binary complex ...........................................152
Figure 4.3 Crystal structure of Dpo4 in the ternary complex ..........................................153
xvii
Abbreviations
AA
AID
ARDS
ASFV PolX
BER
BRCT
C gene
CDR
dFdC
dFdCDP
dFdCMP
dFdCTP
DNA
DNA-PK
dNTP
dPolλ
dRPase
dRPase
DSBs
EDTA
FIAU
fPolλ
HhH
HPG
Ig
MALDI-ToF MS
mtDNA
NHEJ
Sulfo-NHS-Biotin
NLS
PCNA
Pol IV
Pol
Pol
Amino Acid
Activation-Induced Cytidine Deaminase
Acute Respiratory Distress Syndrome
African Swine Fever Virus DNA Polymerase X
Base Excision Repair
Breast Cancer Susceptibility Protein BRCA1 C-Terminus
Constant Gene Segment
Complementarity Determining Region
2’-deoxy-2’,2’-difluorocytidine
Gemcitabine Diphosphate
Gemcitabine Monophosphate
Gemcitabine 5’-Triphosphate
Deoxyribonucleic Acid
DNA Dependent Protein Kinase
2′-Deoxynucleotide Triphosphate
DNA Polymerase Lambda Deletion Construct (AA 132-575)
5’-Deoxyribose-5-Phosphate Lyase
5′-Deoxyribose-5-Phosphate Lyase
Double-Stranded Breaks
Ethylendiaminetetraacetic Acid
1-(2-Deoxy-2-Fluoro-β-d-Arabinofuranosyl)-5-Iodouracil
Full-Length DNA Polymerase Lambda
Helix-Hairpin-Helix
p-Hydroxyphenylglyoxal
Immunoglobulin
Matrix Assisted Laser Desorption Ionization Time of Flight Mass
Spectrometry
Mitochondrial DNA
Non-Homologous End Joining
N–Hydroxysulfosuccinimido Biotin
Nuclear Localization Signal
Proliferating Cell Nuclear Antigen
DNA Polymerase IV
DNA Polymerase Beta
DNA Polymerase Eta
xviii
Pol
Pol
Pol
Pol
Pol
Pol
Polβ
Polγ
Polλ
RAG
RSS
TCR
TdiF
TdT
TdTL
TdTS
tPolλ
XRCC4
DNA Polymerase Iota
DNA Polymerase Kappa
DNA Polymerase Lambda
DNA Polymerase Mu
DNA Polymerase Sigma
DNA Polymerase Zeta
DNA Polymerase Beta
Human DNA Polymerase Gamma Holoenzyme
DNA Polymerase Lambda
Recombinase Activating Gene Product
Recombination Signal Sequences
T Cell Antigen Receptor
TdT Interacting Factor
Terminal Deoxynucleotidyltransferase
TdT Long Isoform
TdT Short Isoform
Truncated DNA Polymerase Lambda (AA 245-575)
X-ray cross complementing group 4
xix
Chapter 1: DNA polymerases and terminal deoxynucleotidyl transferase
1.1. Introduction
It is of paramount importance to biological organisms that their genetic
information be preserved in an intact, replicable state in order to maintain and perpetuate
their existence. However, perfect conservation of the genome is neither possible nor
desirable, because those infrequent and tiny changes in the molecules of life provide the
basis for evolution and adaptation to an ever-changing and frequently hostile
environment. Fortunately, there has come to exist an ensemble of machinery to allow for
both the faithful maintenance and the subtle, random change that has laid the foundation
for life itself. At the very core of this machine are the DNA polymerases, the caretakers
of the genome. These polymerases are responsible for DNA replication and
recombination, repair of DNA lesions, and even tolerance of potentially lethal DNA
damage through unique mechanisms of lesion bypass. Most polymerases are highly
accurate when performing the tasks of genomic replication and repair. However, in those
circumstances when “making a mistake is the only way to get ahead”,[1] a lesser known
group of low-fidelity polymerases can be brought to bear. Members of this group have
1
greatly enhanced flexibility with respect to what substrates they can utilize. This reduced
degree of fidelity possessed by these enzymes is what allows them to replicate patches of
DNA that are severely damaged or even completely non-informative.
1.2. Classification of DNA Polymerases
Beginning with Arthur Kornberg’s discovery of Escherichia coli (E. coli) DNA
polymerase I in the 1950s,[2, 3] many DNA polymerases performing a diverse repertoire
of biological functions have been identified. These DNA polymerases have been grouped
into six families: A, B, C, D, X, and Y based on their phylogenetic relationships [4, 5].
With the exception of the highly conserved carboxylate residues found within the
polymerase active sites (Section 6.1), little sequence similarity is shared between
members of different families. Indeed, within each polymerase family, many distinct
biological functions can be found. Except for the Y-family, no DNA polymerase family
has yet been found that is universally conserved among the three domains of life
(Archaea, Bacteria, and Eukaryota). Not surprisingly, evolution of the DNA polymerase
families is very complex and is likely to involve multiple gene exchanges between
cellular and viral proteins [6].
Family A
Family A DNA polymerases can be found in bacteria, metazoa, plants,
mitochondria and viruses [6]. In addition to their template-dependant polymerase activity,
2
members of the A family possess 3’ 5’ exonuclease activity and possibly 5’
3’
exonuclease activity. The representative member of family A is E. coli DNA polymerase
I, which possesses all three of the activities mentioned above and is involved in DNA
repair and recombination [7]. Mitochondrial DNA polymerase , another member of
family A, is a heterodimeric protein that exhibits both polymerase and 3’
5’
exonuclease activities [8]. While primarily functioning in the replication of mitochondrial
DNA, mitochondrial DNA polymerase
also takes part in the repair of mitochondrial
DNA through its 3’ 5’ exonuclease activity [9]. Eukaryotic DNA polymerase [1] is
also a member of family A and helps to replicate specific templates containing abasic
lesions, via its 3’ 5’ exonuclease functionality [10]. Viral replicative DNA polymerases
in family A, such as vaccinia virus DNA polymerase,[11, 12] catalyze templatedependant viral genome replication.
Family B
Family B is mainly composed of the eukaryotic replicative polymerases [5] which
are homologous to E. coli polymerase II,[1] the prototype of family B. Family B
members can be also found in Archaea, proteobacteria, phages, and viruses [6]. DNA
polymerases , , , and
[13, 14] are typical Family B members. Aside from template-
dependant polymerase activity, most family B members such as DNA polymerases
also possess 3’ 5’ exonuclease activity. Although lacking associated 3’
exonuclease activity, DNA polymerase
and
5’
contains both polymerase and primase activities
3
and plays a significant role in eukaryotic replication [15]. DNA polymerase
possesses
93% conservation from mouse to human [16] and functions in elongation of the leading
and lagging strands during DNA replication. DNA polymerase
is involved in DNA
repair [17]. Analysis of the N-terminal and C-terminal regions of polymerase
that this enzyme serves as a way of quality control in the cell while
polymerizes extended DNA chains [5]. DNA polymerase
indicate
exclusively
(Pol ), a recently discovered
enzyme, is likely involved in DNA lesion bypass [13, 18, 19] and somatic hypermuation
(Section 1.8).
Family C
Family C DNA polymerases are found exclusively in bacteria [6]. Family C is a
high-fidelity family and each member possesses both template-dependant polymerase and
3’ 5’ exonuclease activities. The prototype of family C is E. coli DNA polymerase III
which replicates the genomic DNA of E. coli [1].
Family D
Family D polymerases are found in the Euryarchaeota subdomain of Archea,[1921] not in bacteria or Eukaryota [6]. Each family D DNA polymerase exhibits both
template-dependant polymerase activity and 3’-5’ exonuclease activity [20, 22-24]. The
high-fidelity Family D polymerases catalyze DNA replication in Euryarchaea.[21] One of
the best known members of Family D is from a hyperthermophilic archaeon Pyrococcus
furiosus (Pfu) [21]. Two Pfu proteins DP1 and DP2 encoded by tandem genes form a
4
polymerase complex: the former is a small accessory subunit while the latter is the large
catalytic subunit. The two Pfu proteins are highly conserved in the Euryarchaeota
subdomain. The homologs of DP2 share more than 50% amino acid conservation while
the DP1 homologs possess more than 30% identity. However, Family D polymerases
generally share little sequence homology to polymerases from other families.
Family Y
Family Y DNA polymerases have been found in Archaea, Bacteria, and
Eukaryota, but not in viruses. In humans, four Y-family members including DNA
polymerases eta (Pol ), iota (Pol ), kappa (Pol ) and REV1 have been identified [19]. Yfamily members known as translesion polymerases have the ability to bypass DNA lesions
which stop replicative DNA polymerases [1, 5]. For example, Pol has demonstrated an
ability to perform translesion synthesis on several aberrant primer-templates including
those substrates containing abasic sites, N-2-acetyl aminofluorene (AAF)-adducts, 8oxoguanine lesions, and (-)-trans-anti-benzo(a)pyrene-N2-dG adducts [25]. All the Yfamily polymerases that have thus far been biochemically characterized are devoid of
intrinsic proof-reading exonuclease activities and catalyze template-dependant DNA
synthesis with low fidelity and poor processivity [19, 25-30]. The fidelity of
polymerization catalyzed by Pol differs with respect to the template base, with an error
rate of 10-2 to 10-4 opposite a template “A”, “G”, or “C” [31-33]. Interestingly, Pol
preferentially selects misincorporation of “G” opposite a template “T”,[29, 31, 32]
possibly by Hoogsteen basepairing [34]. When a Y-family polymerase encounters a DNA
5
lesion, it can bypass the lesion either in an error-free or in an error-prone manner [1]. For
example, human DNA polymerase
has been shown to faithfully replicate through cis-
syn thymine dimers [19]. Mutational inactivation of human Pol
leads to cancer-prone
syndrome, a variant form of xeroderma pigmentosum (XPV) [19, 25, 26]. In contrast,
human Pol bypasses an 8-oxoguanine lesion by incorporating either base “A” or “C”, an
abasic site by inserting base “A” and less frequently base “G”, a (+)-trans-antibenzo[a]pyrene-N2-dG adduct by incorporating base “A” and less frequently base “T”, a
1,N6-Ethenodeoxyadenosine lesion by inserting base “T” and less frequently base “A”, an
O6-methylguanine lesion by incorporating base “C” or “T” [27, 29, 32].
Family X
The X Family of DNA polymerases is a subdivision of a larger superfamily of
nucleotidyltransferases [6]. Members of this family can be found in Achaea, Bacteria,
Eukaryota, and in viruses. In addition to TdT, DNA polymerase
polymerase
(Pol ), DNA
(Pol ), [35-37] DNA polymerase µ (Pol ), [35, 38] African swine fever
virus DNA polymerase X (ASFV PolX),[39] yeast DNA polymerase IV (Pol IV), [40]
and yeast DNA polymerase
(Pol ) [41-43] are also members of Family X [44]. TdT is
known to catalyze non-templated, random nucleotide addition at the V(D)J junctions
thereby increasing antigen receptor diversity (Section 1.8). In vivo, TdT expression is
thought to be restricted to primary lymphoid tissues (thymus and bone marrow); [45-49]
although other theories do exist (Section 1.9). Pol
removes the 5’-deoxyribose
phosphate moiety [50, 51] and catalyzes gap-filling synthesis [50] during base excision
6
repair (BER). ASFV PolX plays a role in BER analogous to the function of its
mammalian counterpart, Pol [39]. Pol couples DNA replication to the establishment of
sister chromatid cohesion [41-43].
Non-homologous end joining (NHEJ), a major pathway for repair of DNA
double-strand breaks introduced by exogenous sources including oxidation and ionizing
radiation, exists in all cell types. Yeast Pol IV functions in NHEJ of double strand breaks
[52] and possibly in BER [53].
So far, the biological role(s) of the recently discovered Pol
have not been
established. It is plausible that Pol contributes to BER since it is related to Polβ and
possesses two key enzymatic activities (gap-filling polymerase and a 5’-2-deoxyribose-5phosphate lyase) required by BER. The gene encoding Pol
is mapped to mouse
chromosome 19. Like Polβ, [54] Pol is expressed at high levels in the developing mouse
testes, suggesting a possible function of Pol in DNA repair pathways, especially BER,
associated with meiotic recombination [37]. In an in vitro BER reconstitution reaction,
recombinant human Pol and Polβ can replace each other to efficiently repair uracilcontaining DNA in the presence of human uracil-DNA glycosylase, human AP
endonuclease, and human DNA ligase I [55]. The role of Pol in DNA repair is further
supported by the following observations: i) mouse embryonic fibroblast Polβ-/- cell
extract contains substantial amounts of active Pol
which can replace Polβ in
reconstituted and uracil-initiated short-patch BER, and monoclonal antibodies against
7
Pol in this cell extract strongly reduce in vitro BER; [56] ii) Pol is the only X-family
DNA polymerase found in higher plants and its expression is induced by DNA-damaging
treatments; [57] iii) Pol protects mouse fibroblasts against oxidative DNA damage and
is recruited to oxidative DNA damage sites [58]. Thus, Pol may complement or support
the function of Polβ in BER in vivo. On the basis of the current biochemical data, the
second proposed biological role of Pol
is to repair double-stranded breaks (DSBs)
through NHEJ pathways [59, 60]. This hypothesis is supported by the results from
immunodepletion studies suggesting that Pol , rather than other X-family polymerases, is
primarily responsible for the gap-filling synthesis associated with NHEJ in human
nuclear extracts [59]. The last proposed role of Pol in vivo is to bypass DNA lesions.
This hypothesis is based purly on its ability to bypass an abasic site in the presence of
Mn2+ in vitro [4, 61]. So far, the generation of knock-out mice through deletion of exons
5-7 of the Pol gene has not yet confirmed the involvement of Pol in this or any other
biological process [62]. These Pol knock-out experiments are likely complicated by the
existence of other DNA polymerases, especially Polβ, [63] which could fill in and
compensate for the loss of functions of Pol .
Similarly, the biological role(s) of another novel X-family member, Pol , have
yet to be identified. Preferential expression in secondary lymphoid tissues as well as the
observed low fidelity of Pol have led to the hypothesis that this enzyme is an errorprone mutase active in somatic hypermutation [64]. The presence of Pol
8
and the
absence of TdT in germinal center B cells, the low levels of Pol expression in thymus
and bone marrow, and the intrinsic terminal transferase activity possessed by Pol in the
presence of Mn2+ all suggest that this enzyme may play a role in V(D)J recombination,
thereby complementing the biological functions of TdT [64]. Moreover, the basal
expression of Pol in most tissues suggests a potential role in NHEJ for general repair of
DNA double-strand breaks. The proposed role of Pol in NHEJ and V(D)J recombination
is substantiated by the following two in vitro observations: Pol and TdT form essentially
identical complexes with the end-joining factors Ku and the XRCC4-ligase IV complex
[65] and Pol promotes microhomology searching and pairing to realign primers with
terminal mismatches by looping out any mismatched template nucleotides [66]. Recently,
Pol , like TdT, has been shown to incorporate both rNTPs and dNTPs using either DNA
or RNA primers [67-69]. Additionally, Pol can bypass several DNA lesions through a
deletion mechanism [70, 71]. The lesion bypass ability of Pol indirectly supports the
proposed role of this polymerase as a mutase during somatic hypermutation.
Other than Pol , none of the Family X polymerases contain proof-reading
exonuclease activity. Recently, recombinant Pol
purified from E. coli was shown to
display a Mg2+-dependant 3’ 5’ exonuclease activity in vitro,[43] although it seems
more experiments would need to be performed to exclude the possibility that this
observed exonuclease activity was due to a contaminating E. coli enzyme.
9
1.3. Sequence Alignment Analysis of the X-family DNA Polymerases
Sequence alignment and three dimensional structural modeling studies predict
that the C-termini of all Family X polymerases possess Pol -like domains (Figure 1.1).
Each Pol -like domain is further divided into the following subdomains: 8-kDa, fingers,
palm, and thumb (Figure 1.2). Notably, we prefer the subdomain nomenclature of Pol
(Figure 1.2), rather than the nomenclature initially proposed for E. coli DNA polymerase
I,[72] to describe the domain structures of X-family polymerases in this article. The
difference between these two nomenclatures is simply a reversal of the names “thumb”
and “fingers” for the subdomains on either side of the palm domain. Interestingly, the
full-length ASFV Pol X, the smallest known nucleotide polymerase (174 residues, 20
kDa), possesses only the palm and thumb subdomains as revealed by nuclear magnetic
resonance spectoscopy (NMR) [73, 74]. In addition to the C-terminal Pol -like domain,
TdT, Pol , Pol , and Yeast Pol IV all have nuclear localization signal (NLS) motifs and
breast cancer susceptibility protein BRCA1 C-terminal (BRCT) domains on their Ntermini. BRCT domains are known to mediate protein/protein and protein/DNA
interactions in DNA repair pathways and cell cycle check point regulation upon DNA
damage [75]. For example, the BRCT domain of TdT is thought to interact with Ku70/86
[76], a protein heterodimer involved in recognizing and binding free DNA ends during
V(D)J recombination and double strand break repair [77]. In Pol , a proline-rich domain
can be found located between the BRCT and Pol -like domains, (Figure 1.1). Analysis of
deletion mutants has suggested that the proline-rich domain may functionally suppress
10
the polymerase activity of Pol
while the BRCT domain does not affect polymerase
activity [78]. In contrast, the Pol -like domain of Pol is not active as a DNA polymerase
in the absence of the BRCT domain [69].
Sequence alignment analysis indicates that Pol is most similar to Polβ sharing
32% amino acid identity [79]. The C-terminal Polβ-like domain of Pol is predicted to
fold in a manner similar to Polβ. Analysis of the NMR structure of the 8-kDa domain of
Pol reveals a high degree of similarity to the corresponding domain in Polβ [80]. The Xray crystal structure of the Polβ-like domain complexed with single-nucleotide gapped
DNA and an incoming nucleotide ddTTP [81] is also similar in many respects to the
ternary structure of Polβ shown in Figure 1.2.
Among all of the X family polymerases, Pol has been found to be the closest
relative of TdT, [1, 5, 70, 82] sharing approximately 42% amino acid identity (Figure
1.1) [5, 38]. These two polymerases are predicted to possess an organization of domains
as shown in Figure 1.1 Unfortunately, the crystal structure of Pol
solved and thus, its predicted domain organization cannot be confirmed.
11
has not yet been
1.4. Isoforms of TdT
TdT itself is highly conserved across the vertebrate phyla, from cartilaginous fish
to birds, and to humans [83-89]. For example, the TdT sequences of skate and shark share
70% identity at the amino acid level and over 50% nucleotide identity with the mouse
TdT [89]. So far, two mRNA splice variants have been reported in mice, and three each
in bovines and humans. The murine mRNA splice variants are translated into mature TdT
isoforms designated TdT short isoform (TdTS) and TdT long isoform (TdTL). Murine
TdTL (529 residues) differs from murine TdTS in that it contains an additional 20 amino
acid residue insertion between the two β-sheets in the TdTS thumb subdomain (509
residues) [90]. This addition is the result of the expression of an additional exon in the
murine TdT gene [91]. Intriguingly, the effect of this insertion on the polymerase activity
of murine TdT is somewhat controversial. Papanicolaou et al. have shown that this
insertion decreases the thermostability of TdTS, but does not affect its catalytic activity
[92]. In contrast, murine TdTL is found by Kearney et al. to possess 3’ 5’ exonuclease
activity, rather than the template-independent polymerase activity associated with TdTS
[93]. Although both murine TdTS and TdTL localize to the nucleus, it is believed that the
long isoform may down regulate the polymerase activity of the short isoform in vivo [94].
However, this hypothesis has yet to be confirmed by experimentation. In transgenic mice
deficient in TdT, the short isoform is sufficient to rescue N-addition activity [94] (Section
1.8) suggesting that the template-independent polymerase activity of TdTS, not TdTL, is
required in vivo.
12
In vivo, the three mRNA splice transcripts in cattle [95] and humans [96] are also
translated into three mature protein isoforms designated TdTS, TdTL1, and TdTL2 [97].
In humans, the normal B and T lymphocytes express exclusively hTdTS and hTdTL2,
whereas hTdTL1 expression appears to be restricted to transformed lymphoid cell lines.
In in vitro recombination and primer extension/digestion assays, both human TdTL
isoforms are shown to possess 3’ 5’ exonuclease activity while human TdTS acts as a
template-independent DNA polymerase [96]. Overexpression of hTdTS or hTdTL2
greatly reduced the efficiency of recombination, which was reverted to normal levels by
the simultaneous expression of both enzymes. These data suggest that alternative mRNA
splicing may prevent the adverse effects of unchecked elongation or diminution of coding
ends during V(D)J recombination, thus affecting the survival of a B or T cell precursor
during receptor gene rearrangements [96].
13
1.5. TdT Expression and Purification
Chang and Bollum were among the first to attempt to purify TdT from calf
thymus cell lysate [98]. Due to proteolysis, they incorrectly suggested that TdT was a
heterodimer of
and β subunits. It was later discovered that purified full-length TdT is
actually a single polypeptide with a molecular weight of approximately 60 kDa [99].
Because efforts to purify TdT from calf thymus were hindered by proteolysis, researchers
started to investigate alternate means of obtaining homogenous TdT [100, 101].
Unfortunately, although several research groups have attempted to express recombinant
human TdT in bacteria, none have succeeded in obtaining soluble and active protein
[102, 103]. It was in 1988 that TdT was first successfully expressed and purified using
the baculovirus expression system [103]. A decade later, active murine TdTS and TdTL
were successfully expressed in E. coli by lowering the bacterial growth temperature to 15
C and overexpressing a rare arginyl tRNA. Those two isoforms obtained in this manner
were successfully purified to apparent homogeneity through column chromatography [92,
104]. This E. coli method allows for large scale production of those full-length murine
TdTS and TdTL for enzymatic and structure-function relationship analysis [92, 104].
Thanks to a high degree of sequence homology, this E. coli method may be applicable in
the production of TdTs from other species as well.
14
1.6. Three-Dimensional Structures of DNA Polymerases
Crystal Structures of Template-Dependant DNA Polymerases
All template-dependant polymerases with known structures (both those
crystalized and those in solution) [34, 72, 73, 88, 105-120] share a similar architecture at
their polymerase catalytic domain. Intriguingly, these structures resemble the right hand
of a human being, with domains that resemble the palm, fingers, and thumb (and were so
named). Domain nomenclature based on this observation was first proposed for E. coli
DNA polymerase I [72] and has since been adopted for other polymerases as well. As an
example, these domains can be seen in Figure 1.2 which showcases the crystal structure
of human DNA polymerase β complexed with a single-nucleotide gapped DNA substrate
and an incoming nucleotide ddCTP [121]. In the polymerase active site, three aspartic
acid residues, one water molecule, the 3’-OH of the upstream primer strand and the
triphosphate moiety of ddCTP act as ligands to bind two Mg2+ ions (Figure 1.3). During
polymerization, the first Mg2+ ion (B) promotes the deprotonation of the 3’-OH of the
primer strand, facilitating the 3’ oxyanion’s nucleophillic attack on the
–phosphate of
the incoming nucleotide. The second Mg2+ ion (A) then stabilizes the pentacovalent
transition state of the –phosphate and assists the leaving of the pyrophosphate (Figure
1.3) [121]. In fact, two metal ions have been found in the active sites of all DNA
polymerases with known crystal structures [34, 72, 73, 88, 105-120]. Based on this
observation, T. A. Steitz proposed that perhaps all DNA polymerases might use a “twodivalent-metal-ion” mechanism to catalyze nucleotide incorporation [122]. These
15
essential metal ions, which are likely to be Mg2+ in vivo, are bound by three carboxylates
(aspartate and/or glutamate). Notably, these carboxylates are conserved across each of the
six DNA polymerase families [123].
Crystal Structure of the Polβ-Like Domain of Murine TdT
The crystal structures of: i) the Polβ-like domain of murine TdTS (residues 130510, resolution: 2.35 Å), ii) the binary complex of the Polβ-like domain and a brominated
DNA primer 9-mer (3 Å, Figure 1.4), and iii) the binary complex of the Polβ-like domain
and an incoming nucleotide ddATP-Co2+ (3 Å, Figure 1.5) have been solved [88]. The
three dimensional structure of the Polβ-like domain resembles a torus (Figure 1.4), with
an
-helical N-terminal 8-kDa subdomain (residues 163–243), an
-helical fingers
subdomain (residues 243–302), a central palm subdomain with a large antiparallel β-sheet
(residues 302–450) and a C-terminal thumb subdomain (residues 450–510) containing a
small antiparallel β-sheet. The thumb subdomain makes extensive contact with the 8-kDa
domain to close the protein ring. Curiously, despite a low shared sequence identity of
only 22–24%, the four subdomains in TdT are structurally homologous to the
corresponding subdomains in Polβ (Figure 1.2). In comparison to the protein sequence of
Polβ, TdT has two insertions of 10–15 residues between β3 and β4 and between β4 and
β5, which form Loop 1 and Loop 2 in the palm subdomain, respectively. The antiparallel
beta sheets in the palm subdomain contain the three aspartate residues (Asp343, Asp345,
and Asp434, Figure 1.5) which are highly conserved within the nucleotidyltransferase
family. These residues have been demonstrated by site-directed mutagenesis
16
experimentation to be essential for the binding of two divalent metal ions and for the
catalytic activity of TdT [102, 124].
Upon analysis of the crystal structure in Figure 1.4, the primer strand is observed
to lie on the palm subdomain, perpendicular to the axis of the protein ring. Notably, only
four nucleotides from the 3’ end of the 9-mer primer are ordered, suggesting tight
association of these four nucleotides with the polymerase. It should be noted that these
residues are in the B-type DNA conformation. The disordered five nucleotides from the
5’ terminus of the 9-mer are not in contact with the protein. Interestingly, the 3’-terminal
nucleotide is located at the position of an incoming nucleotide found in the ternary
complex of Polβ shown in Figure 1.2 [121] Thus, the structure in Figure 1.4 is considered
by Delarue et al. to mimic the ternary structure of TdT, DNA, and a nucleotide [88].
Alternatively, the binary structure in Figure 1.4 may represent the complex of TdT and a
DNA product after TdT has incorporated the incoming nucleotide but before TdT has
repositioned itself for the binding of next incoming nucleotide. Nevertheless, the position
of the primer in the active site of TdT shown in Figure 1.4 must change in the presence of
an incoming nucleotide. Interestingly, the small number of ordered nucleotides in Figure
1.4 is consistent with the finding that TdT requires at least a 5’ phosphorylated trimer as a
primer in order to act as an efficient polymerase [125]. However, as discussed below, in
the presence of Mn2+, TdT has been observed to catalyze DNA synthesis de novo [82]. In
addition, no atoms can be identified between the amino acid residues of TdT and the
primer nucleotides which reside in sufficient proximity to one another to be considered a
17
polar interaction, thus indicating that binding must rely entirely upon interaction with the
sugar phosphate backbone. This observation could further explain the in vitro results
indicating that TdT displays a low degree of specificity with respect to nucleotide
selection [88]. TdT does, however, strongly prefer single stranded DNA. Upon
examination of the location of Loop 1, it is likely that the presence of this lariat-shaped
loop might preclude the accommodation of a template strand, thus making TdT an
inefficient DNA polymerase in the presence of double-stranded DNA [88].
Like several DNA binding proteins including Polβ and Pol , [126] TdT possesses
two DNA-binding helix-hairpin-helix (HhH) motifs (residues 208-231 and 244-267). The
second HhH motif (residues 244-267), which interacts with the primer strand, uses the
carbonyl groups of residues Thr253, Val255, and Val258 as ligands to chelate a Na+ ion
(Figure 1.4). Similarly, these HhH motifs in Polβ [105, 121] and Pol [53, 127] are found
to coordinate K+ or Na+ ions and participate in sequence-independent interactions with
the backbones of the template and primer strands. The HhH motifs of Polβ are further
shown to have a preference for cations in the order K+>Na+> Mg2+>Ca2+ [105].
The binary complex of an efficient template-dependant DNA polymerase and
DNA usually binds to a correct incoming nucleotide tightly with a dissociation
equilibrium constant (Kd) in the low micromolar range. This high ground-state binding
affinity is partly achieved through base pairing interactions between the incoming
nucleotide and the opposite template base. Since there is no template with which to base
18
pair and anchor an incoming nucleotide in the active site of TdT, nucleotide binding must
result exclusively from interaction with the polymerase active site. While it is true that
the actual nucleotide binding site can only be revealed by observing the ternary structure
of TdT, a primer, and a nucleotide (not yet reported), the nucleotide binding site of TdT
can be estimated through analysis of the binary structure of the Polβ-like domain and
ddATP-Co2+ (Figure 1.5) [88]. In this structure, it can be seen that the aromatic ring of
Trp450 is parallel to and partially stacked with the adenine ring of ddATP, with the CZ2
atom of Trp450 located 3.6 Å from the C8 atom of the adenine ring. The side chain of
Lys403 is observed to point towards the adenine ring of the ddATP, with the
amino
group located 4 Å above the base. Additionally, one will notice that the anionic
triphosphate moiety of ddATP is neutralized and stabilized by three positively charged
residues (Arg336, Lys338, and Arg454). The sugar ring of the incoming ddATP is
observed to bind Trp450 on one side and reside in close proximity to the cis-peptide bond
between Gly452 and Ser453 on the other side. Notably, none of the amino acid residue
side chains seem to reside in sufficiently close proximity to the 2’ or 3’ position of the
ddATP ribose ring to sterically prohibit the accomodation of a ribonucleotide in the
proposed active site. Therefore, one might hypothesize that TdT should be able to
incorporate both deoxynucleotides and ribonucleotides with similar efficiency. This
prediction is substantiated by the observed sugar selectivity values of TdT, which lie in
the range of 2-5 [124] or 2.6-8.9 [67]. However, DNA polymerases such as E. coli DNA
polymerase I[128] and T7 DNA polymerase, [129] do not specifically bind a nucleotide
in the free state, prior to the binding of DNA. This arouses the suspicion that the observed
19
interactions between the active site residues of TdT and ddATP in Figure 1.5 might be
altered in the presence of a single-stranded DNA primer. Furthermore, TdT’s N-terminal
BRCT domain, which in this structure is absent, may also affect the nucleotide binding
site through domain-domain interactions.
20
1.7. Enzymatic Activities of TdT
Template-Independent Polymerase Activity
Most DNA polymerases require a DNA template during replication of genomic
DNA, while repairing DNA damage or bypassing DNA lesions. However, exceptions can
be found in certain members of family X. Within this family, DNA polymerases possess
either template-dependant or template-independent activity. Pol , for example, catalyzes
template-dependant gap-filling DNA synthesis durning the process of base excision
repair [51]. However, TdT will add random nucleotides to single-stranded DNA in a
completely template-independent manner. In fact, TdT actually prefers single-stranded
DNA over double-stranded DNA (recessed and blunt) and completely lacks the ability to
copy a template [130].
dRPase-Deficiency in TdT
Cellular DNA is subject to a continuous assault by exogenous and endogenous
DNA damaging agents. Under these conditions, DNA will accumulate a number of
harmful and potentially lethal lesions. One of the major mechanisms by which these
aberrations are corrected is the base excision repair pathway. A major player in this
process, Polβ, catalyzes the following crucial enzymatic steps: it removes the 5’deoxyribose phosphate moiety via its 5’-deoxyribose-5-phosphate lyase (dRPase) activity
[131, 132] after it has catalyzed gap-filling synthesis to replace the previously excized
nucleobases [132]. The active site residues of the dRPase activity are located in the 821
kDa domain of Polβ. This domain will bind to the downstream primer of a gapped DNA
substrate and increase the processivity and efficiency of polymerization [121]. The
terminal 5’-phosphate on the downstream primer is buried in a positively charged pocket
of Polβ consisting of His34, Lys35, Tyr39, Lys60, Lys68 while Lys72 acts as the
nucleophile for the dRPase reaction [121, 133]. Although structurally TdT contains a
similar 8-kDa subdomain (Figures 1.1 and 1.2), it lacks dRPase activity for the following
two reasons: i) the above residues essential for the lyase activity in Polβ are not
conserved in TdT; ii) the 8-kDa domain is highly basic in Polβ (net charge +10), but is
acidic in murine TdT (net charge –6) and all other known TdTs (net charge –4 to –6) [88].
Logically, a basic 8-kDa domain will bind more tightly to single-stranded DNA than an
acidic 8-kDa domain simply due to charge-charge interactions. Lack of interaction
between the 8-kDa and the 9-mer oligo in Figure 1.4 confirms this prediction.
Primer Requirement
In order to catalyze template-independent polymerase activity, TdT requires a
primer at least as large as a trinucleotide, a free 3’-OH moiety for extension of that
primer, and a free primer 5’-phosphate [125]. For example, pdApdApdA is an active
primer but not dApdApdA. The minimum size of the primer is similar to the number of
ordered nucleotides in Figure 1.4 (Section 1.6). Interestingly, when a primer contains a
3’-terminal β-L-nucleotide, TdT can only add one or two dNTPs to the primer [134].
22
Metal Ion Dependence
As mentioned previously, it was T. A. Steitz who first proposed that perhaps all
polymerases might use a “two-divalent-metal-ion” mechanism to catalyze nucleotide
incorporation [122]. These essential metal ions, which are likely to be Mg2+ in vivo, are
bound to DNA polymerases via three conserved carboxylates (aspartate and/or glutamate)
found in the palm domain of the enzyme. TdT is no exception. It too requires the
presence of divalent metal ions as cofactors. However, there is more than one species of
ion including Mg2+, Mn2+, Zn2+, and Co2+ that can be incorporated into the TdT active
site in vitro. The binary structure of TdT complexed with ddATP-Co2+ (Figure 1.5)
reveals that two bound Co2+ ions are next to ddATP at the active site, [88] indicating that,
TdT too likely utilizes a “two-divalent-metal-ion” mechanism to catalyze nucleotide
incorporation [122]. One should be cautioned however, that biochemical studies have not
yet revealed unequivocally the number or identity of the metal ions bound by TdT in
vivo. Curiously, the efficiency of polymerization and the bias towards purines and
pyrimidines can be significantly affected by the identity of the metal ion that TdT has
chelated. For example, in the presence of Mg2+, purine incorporation occurs at about a
10-fold faster rate than pyrimidine incorporation (Table 1.1), [135] however this substrate
specificity is opposite in the presence of Co2+ with pyrimidines favored over purines by
10-fold (Table 1.1). In addition, micromolar amounts of Zn2+ added to the reaction along
with Mg2+ increase the efficiency of polymerization of all nucleotides [136].
23
Kinetic Mechanism of Template-Independent Polymerization
As yet, the kinetic mechanism for template-independent polymerization catalyzed
by TdT has not been reported. Although a handful of template-dependant DNA
polymerases that share the same minimal kinetic mechanism shown in Figure 1.6, have
been kinetically characterized [137]. In this mechanism, DNA first binds to a polymerase
to form E DNAn (Step 1). This binary complex then binds an incoming nucleotide dNTP
to form a ground-state ternary complex E DNAn dNTP (Step 2) in which the polymerase
is in an open conformation. In the following step, nucleotide binding energy is used to
induce a change in protein conformation to form a tightly bound ternary complex
E* DNAn dNTP in which the polymerase is in a closed conformation (Step 3). This
open closed conformational change is then followed by the incorporation of the dNTP
into the growing DNA polymer and formation of pyrophosphate (Step 4). After the
second protein conformational change (Step 5), pyrophosphate is released. At this point,
if the polymerase is not processive, it will dissociate from the DNA product (Step 7).
However, if the polymerase is processive, it will translocate to the next template base
(Step 8) and start the incorporation cycle again. For DNA polymerases including the
Klenow fragment of E. coli DNA polymerase I, [138] T7 phage DNA polymerase, [139]
HIV-1 reverse transcriptase, [140, 141] human mitochondrial DNA polymerase, [142]
yeast DNA polymerase , [143] and Sulfolobus solfataricus DNA polymerase IV, [30] a
noncovalent step preceding phosphodiester bond formation (Step 3) limits correct
nucleotide incorporation, while phosphodiester bond formation itself (Step 4) is ratelimiting for incorrect nucleotide incorporation. Thus, Step 3 is considered as a critical
24
fidelity check point [137]. Previous X-ray structural studies have suggested that Step 3
may involve the swing and closure of the finger domain of these polymerases once a
correct nucleotide is bound [107, 144, 145]. However, recent stopped-flow fluorescence
studies suggest that local structural rearrangement at the active site, rather than the finger
domain closing, may limit single nucleotide incorporation [137, 146, 147]. In contrast,
the phosphodiester bond formation (Step 4) is found to kinetically limit both correct and
incorrect nucleotide incorporation catalyzed by Polβ [121, 148]. Although, a significant
protein conformational change in Step 3 is observed when the binary and ternary
structures of Polβ are compared [121]. In the case of TdT, our preliminary kinetic studies
have indicated that it likely follows the same kinetic mechanism shown in Figure 1.6 to
incorporate a single nucleotide, but with different microscopic rate constants (J. Fowler
and Z. Suo, unpublished results). Whether the protein conformational change step (Step
3) or the chemistry step (Step 4) limits single nucleotide incorporation catalyzed by TdT
is not clear at present. The binary structure of TdT shown in Figure 1.4 superimposes best
with the ternary structure of Polβ*•DNA•dNTP (the closed conformation), [88]
suggesting that TdT may not undergo significant conformational change in Step 3. This
seems to be reasonable because TdT, unlike template-dependant DNA polymerases, does
not have to consider fidelity. Thus, a slow conformational change step (Step 3), which is
utilized by replicative DNA polymerases to perform fidelity check, is not necessary. If
this structural prediction is correct, Step 4 rather than Step 3, likely limits nucleotide
incorporation catalyzed by TdT. However, if Step 3 is limiting, local structural
rearrangements at the active site of TdT must be involved [137]. Finally, because TdT
25
catalyzes DNA synthesis in a strictly distributive mode, [124] TdT most likely dissociates
from a DNA product (Step 7), rather than progressing through Step 8, to complete the
incorporation cycle (Figure 1.6). This prediction is reasonable because there is no
template involved. We suspect that the primer strand alone does not form a stable DNA
-helix as double-stranded DNA, leading to difficult translocation of TdT along the DNA
after TdT incorporates a nucleotide.
Nucleotide Selectivity
The physiological role of TdT is to catalyze the addition of random dNTPs onto
the 3’ hydroxyl terminus of single-stranded DNA. Although, in vitro it has also been
shown that TdT can accept ribonucleotide triphosphates (rNTPs). Generally, sugar
selectivity for DNA polymerases is very high. For example, Polβ has a sugar selectivity
in the range of 2,000-6,000 [67]. In comparison, TdT has only a vanishingly small
preference (2-9 fold) for dNTPs over rNTPs [124, 125, 149, 150]. Strikingly, TdT can
incorporate a wide variety of nucleotide analogs such as p-nitrophenylethyl triphosphate,
[151] p-nitrophenyl triphosphate, [134] d4TTP, [134] cordycepin 5’-triphosphate, [152]
2’,3’-dideoxynucleotides (ddNTPs), [153]
-D-dNTPs, [154] and dinucleoside 5’,5’-
tetraphosphates (Figure 1.7) [155]. TdT can also catalyze the transfer of phosphate ester
groups and phosphonate residues from their corresponding triphosphate derivatives onto
the primer 3’-terminus (Figure 1.7) [155]. The broad nucleotide substrate specificity of
TdT suggests that the specific interaction between an incoming nucleotide and the TdT
active site most likely occurs at the triphosphate moiety of the nucleotide, whereas the
26
role of the base and sugar may be of lesser importance. If the incorporated analog lacks a
3’-OH moiety, it will terminate primer extension, leading to drug-induced apoptosis
[156]. For example, cordycepin has been used recently to target TdT-positive leukemia
cells [155] as it is known that the level of TdT in the leukocytes of leukemia patients is
very high [157]. Moreover, taking advantage of its broad nucleotide substrate
skpecificity, TdT has been used to synthesize copolymers like (pdT)6(dG)10(dA)313, [158]
label primer 3’-termini with highly radioactive nucleotides, [152] and attach biotinylated
or fluoresceinated probes to synthetic oligonucleotides [159].
Other Enzymatic Activities
In addition to template-independent polymerase activity, TdT can also catalyze
pyrophosphorolytic dismutation of oligodeoxy-nucleotides by removing a 3’-nucleotide
from one oligonucleotide and adding it to the 3’-end of another [160]. TdT further
distinguishes itself from classical DNA polymerases through its ability to catalyze the
creation of polynucleotides of 2-mer, 7-mer, 15-mer, and a 21-mer de novo in the
presence of Mn2+, given only dNTPs. This ability has only been observed in two other
polymerases to date, Polλ and Pol
[82]. It has been speculated that these DNA
fragments might act as a recognition signal for DNA repair or recombination machinery
[82]. However, the relevant metal ions for DNA polymerase in vivo are likely Mg2+,
rather than carcinogenic Mn2+, [161] therefore these observations may not be
physiologically relevant. Interestingly, as mentioned earlier, it has recently been reported
27
by Kearney et al. that the long isoforms of murine and human TdT’s possess 3’
exonuclease activities, rather than template-indepadant polymerase activities [93, 96].
28
5’
1.8. Immune System and TdT
The main goal of the vertebrate adaptive immune system is to defend the
organism from harmful foreign agents or “antigens”. This objective is accomplished
primarily through the recognition of antigens by antigen-binding proteins that the
immune system produces. These proteins are divided into two major classes: the
immunoglobulins (Igs, Figure 1.8) which are either free glycoproteins present in the
serum and tissue fluids (antibodies) or attached to the membrane of certain cells such as
memory B cells and the T cell antigen receptors (TCRs, Figure 1.9) which are very
similar glycoproteins present on the surface of T cells [162]. In general, these proteins are
remarkably specific for particular antigens that they recognize. Therefore, the number of
these antigen-binding proteins must be tremendously large so that the immune system can
respond to the maximum number of antigens. In fact, some have estimated that in
humans, there are approximately 1014 unique Igs and around 1018 unique TCRs [163,
164]. Given that there are somewhere between 30,000 and 150,000 genes in the entire
human genome, [165] it would seem impossible that each of these antigen-binding
proteins could be encoded by a unique gene. Even when one takes into account
alternative gene processing pathways, the number of different gene products derived from
a single gene cannot exceed a small number [164]. Therefore, in order to mount a
maximally effective defense, the cells of the immune system have developed an
interesting method to “shuffle” and “randomize” small elements of the germline
immunoglobulin and T cell receptor genes. This process makes it possible for a relatively
29
small number of genes to establish an astonishingly large body of unique proteins
(Section 1.8).
A basic Ig protein consists of two large polypeptides and two small polypeptides
(Figure 1.8). The two larger “heavy” chains are joined by several disulfide bonds at what
is referred to as the “hinge” region. The two smaller “light” chains are each joined to one
of the heavy chains by a disulfide bond. Each chain, heavy and light, has a conserved
“constant” region and a “variable” region. The “constant” region of a heavy chain
consists of equal thirds (CH1, CH2, and CH3) that are similar in sequence. The “constant”
region of the light chain (CL) closely resembles CH1, CH2, and CH3. A T cell antigen
receptor, in contrast, consists of a 43-kDa α chain joined to a 43-kDa β chain by a
disulfide bond (Figure 1.9 part (B)). Each chain of a T cell antigen receptor also
possesses both “constant” and “variable” regions [162]. Although the overall structures of
Igs are conserved, the N-terminal variable regions of both the heavy and light chains each
contain three hypervariable loops called complementarity determining regions (CDRs)
which make specific contact with antigens and are responsible for the high antigen
affinity of Igs (Figure 1.8). The binding affinity between an antibody and an antigen
(protein, oligosaccharide, etc.) is generally in the picomolar to 0.1 micromolar range
[166, 167]. For example, anti-lysozyme antibody D1.3 binds to the antigen, hen egg
white lysozyme, with an affinity of 2 nM (Figure 1.8) [168]. It is believed that induced fit
mechanistics play a role in the formation of many tight antibody-antigen complexes.
Variable regions of Ig heavy chains and TCR β chains are assembled from three gene
30
segments called variable (V), dependent (D), and joining (J) segments. In comparison,
each corresponding variable region in the Ig light chain and TCR α chain is assembled
from V and J gene segments only. In both mice and humans, germline V, D, and J gene
segments are inherited as tandem clusters (Figure 1.9 part (A) and Figure 1.10 part (A)).
Low-affinity Igs such as IgM circulate in the blood before encountering antigens,
while high-affinity Igs are necessary to disable viruses, toxins, and other foreign
microorganisms. In the absence of the four classes of high-affinity Igs including IgA,
IgD, IgE, and IgG, an individual is unable to fight off infection and thus dies at a
premature age [169, 170]. Therefore, it becomes clear that a diverse pool of antibodies is
of crucial importance to the ability of an organism to defend itself against the wide
variety of antigens with which it may be presented. Currently, three mechanisms
including V(D)J recombination, somatic hypermutation, and “Class” or “Isototpe”
switching, are known to diversify Igs in vivo. In comparison, only V(D)J recombination
is involved to diversify TCRs in vivo. Here, we address only the first two mechanisms
because of their relevance to TdT and other DNA polymerases. “Class” or “Isototpe”
switching (not reviewed here) is a region-targeted recombination pathway to translocate a
VDJ gene from a site near one “constant” gene segment (C gene), to a site near another C
gene. Many recent reviews covering the “Class” or “Isototpe” switching mechanism can
be found in the literature [171-175].
31
V(D)J Recombination
In order to achieve greater diversity in the variable regions of both Ig heavy
chains and TCR β chains and maximize antigen-binding affinity, germline V, D, and J
gene segments are recombined in a combinatorial manner to generate specific antigenbinding proteins (Figure 1.10). In germline DNA, V, D, and J genes are flanked by
conserved DNA motifs called recombination signal sequences (RSS). An RSS consists of
two conserved motifs: a heptamer (CACAGTG) and a nonamer (ACAAAAACC). One
should note however, that these motifs can and frequently do vary, thus affecting the
efficiency of recombination [176]. These elements must be separated by a stretch of DNA
that is either 12 or 23 base pairs in length, making the final size of an RSS either 28 (12RSS) or 39 (23-RSS) base pairs long (Figure 1.10 part (B)) [177]. In immunoglobulin
genes and some T cell receptor genes, the V and J segments are associated with a 23RSSs while the D segments have 12-RSSs flanking them on either side. Recombination
events always occur between gene segments bordered by RSSs of different sizes, thus
promoting the recombination of gene segments from different regions and making it
unlikely that two segments of the same region will recombine [177]. This is known as the
“12/23 rule” [178]. Upon recognition of the RSS elements by a mixed tetramer
containing two monomers of recombinase activating gene products 1 and 2 (RAG1 and
2), a nick is created 5’ to the heptamer element of the RSS (Figure 1.10 part (C)). The
nicked strand 3’-OH then initiates a trans-esterification (mediated by RAG) on the
phosphate of the complementary strand thereby forming a covalent hairpin structure on
the coding end, leaving the other end of the double stranded break (the RSS) blunt [177,
32
179]. Interestingly, although the presence of the RAG proteins (RAG1 and RAG2) is
critical for the initiation of V(D)J recombination, the exact mechanism of their activity is
still unclear. It has been noted, however, that the active site of RAG1 is similar in many
ways to the active sites of retroviral integrases and transposases [180-183]. After
cleavage, the DNA ends remain in association with the RAG proteins which most likely
aids in the subsequent joining steps [183]. These newly created DNA ends are processed
and joined through the NHEJ pathway in which the ends are recognized by the Ku
proteins and then processed and ligated through the actions of “Artemis”, DNA-PKcs,
TdT, X-ray cross complementing group 4 (XRCC4), and DNA ligase IV [179, 184-186]
and possibly other factors not yet identified [176]. These NHEJ proteins will be described
further in the following section.
Role of TdT in V(D)J Recombination
While, the combinatorial selection of V, D, and J gene segments serves as a major
source of diversity for Ig and TCR genes, there is yet another strategy employed by the
adaptive immune system to enhance the antigen-binding repertoire. During V(D)J
recombination, the joining of V, D, and J segments is highly imprecise and the coding
joints between these segments are observed to lose or gain nucleotides before they are
finally ligated (Figure 1.10 part (C)). This extra genetic information may arise via our
proposed mechanism shown in Figure 1.11. This mechanism is based on the enzymatic
properties of “Artemis”, TdT, exonucleases, template-dependant DNA polymerases, and
ligases and on the statistical analysis of the sequences of 543 mouse Ig heavy chains
33
[187]. In the first step, the covalently closed coding ends are opened via scission of the
phosphodiester backbone at an imprecise location very near to the apex of the hairpin,
most likely by the endonuclease “Artemis” [188]. This endonucleolytic cleavage can
leave the DNA either blunt ended or with either a 3’ or 5’ overhang. Following cleavage,
a polynucleotide palindromic sequence is likely to exist due to the imprecision of the
cleavage site. For example, Figure 1.11 shows that the terminal two nucleotides from the
5’ end of one of the gene segments are reversed and joined to the 3’ end of the other. The
DNA ends created in this way are favorable substrates for TdT based on our preliminary
finding that TdT catalyzes template-independent polymerization on this type of DNA as
efficiently as on exclusively single-stranded DNA substrates (J. Fowler and Z. Suo,
unpublished data). Interestingly, the statistical analysis of the sequences of 543 mouse Ig
heavy chains indicates that the “N regions” are formed predominantly from DNA plus
strand or from DNA minus strand polymerizations, rather than from both simultaneously
[187]. Thus in Step 2, TdT is shown to only extend the protruding plus strand at the V
segment. The same statistical analysis has also suggested that homologous overlaps of as
few as one nucleotide between gene segments may cause significant skewing of
recombination sites [187]. Thus in Step 3, the microhomology alignment between the two
protruding strands is shown to occur through base pairing between two nucleotides. In
Step 4, the mismatched 3’-nucleotides on both strands are excised by an unknown 3’
5’
exonuclease. In Step 5, an unknown template-dependent DNA polymerase catalyzes gapfilling DNA synthesis. It should be noted however that the fidelity of this gap-filling
polymerase must not be too low because the sequence bias in “N regions”, which are rich
34
in Gs and Cs, generated by TdT would be lost in the presence of an error-prone DNA
polymerase. Also, double-stranded DNA containing any number of mismatched base
pairs would not be stable enough for ligation in the next step. In addition, this gap-filling
polymerase must be relatively processive in order to catalyze efficient V(D)J
recombination. Additionally, to increase coordination of the V(D)J recombination
reaction, the 3’ 5’ exonuclease activity, and the template-dependent DNA polymerase
activity should reside on the same enzyme, as occurs in some repair DNA polymerases
like eukaryotic DNA polymerase . Finally, the nicked strands are ligated by DNA ligase
IV in the presence of XRCC4. Notably, many steps in Figure 1.11 may require the
addition of other NHEJ proteins, e.g. stabilizing of the base pairs in Step 3 by the Ku
proteins. Moreover, exonucleolytic activity may shorten the 3’ palindromic sequence by
one or more nucleotides in Step 4. However if any nucleotides from the palindromic
sequence that were encoded by the germline DNA remain in the final coding joint, these
bases are designated “P” nucleotides (for palindromic) which in Figure 1.11 are shaded
[189, 190].
It has been demonstrated that TdT can only accomplish “N” nucleotide addition
after recruitment to the site of recombination [76, 191]. In addition, it has been shown
that the factor responsible for the recruitment of TdT to the site of its activity is Ku.
Perhaps one of the best understood functions of Ku is as a regulatory component of the
DNA dependent Protein Kinase (DNA-PK) which is a core component involved in the
mammalian NHEJ pathway. However, Ku is also critical to the production of N-region
35
diversity during the process of V(D)J recombination [38, 76, 88]. DNA-PKcs, the
catalytic subunit of DNA-PK, is a nuclear serine/threonine kinase which is recruited to
DSBs by the Ku heterodimer and initiates repair of this potentially lethal damage via the
phosphorylation of many downstream targets [192]. Ku is believed to regulate DNA-PK
by acting as the primary recognition element of DSBs. Ku functions as a heterodimer
composed of 70 and 80 kDa subunits having a high affinity for the ends of DNA duplexes
[76]. After Ku binds to the ends of a DSB, the catalytic subunit of DNA-PK (DNA-PKcs)
is recruited to initiate NHEJ, thereby repairing the damage. Therefore, it is reasonable to
hypothesize that Ku is functioning in a similar capacity to recruit TdT to the ends of gene
segments during the process of V(D)J recombination. The first convincing evidence to
confirm that Ku and TdT are functional partners came in 1999 when these proteins were
co-immunopreciptated from human Molt-4 lymphoid cell extracts [76]. This interaction
was shown through mutational analysis to occur via the N-terminal BRCT domain of
TdT. This revelation is not surprising considering that BRCT domains are commonly
involved in mediation of protein-protein interactions between DNA repair components
[38, 76, 82, 88]. In addition, although TdT is stable in Ku80 deficient fibroblasts, [191]
V(D)J junctions from these cells lack N regions, suggesting that Ku80 may play a crucial
role in their formation. Ku70 is believed to stabilize Ku80, as neither of these proteins is
stable by itself. It has also been demonstrated that Ku70 makes more intimate contact
with DNA than does Ku80 [193]. It should be noted however, that only the Ku70/80
dimer can act to recruit TdT.
36
TdT Regulation
As stated previously, it is of great importance that V(D)J recombination take
place in order for the creation of a competent immune system. However, it is just as
important that the process be tightly regulated. V(D)J recombination is observed to occur
only very early in lymphocyte development and only at very well defined loci [179].
While it has been demonstrated that TdT is regulated at the level of transcription by
proteins such as AP-1, [194] regulation is thought to be achieved principally through the
expression of the RAG genes [179]. However, the rate of random nucleotide addition
catalyzed by TdT also seems to be under a complex system of both negative and positive
control.
In addition to regulation at the level of expression, TdT is also regulated by
proteins called TdT interacting factors (TdiFs). One of these proteins, TdiF1 is found to
bind to TdT and enhance its polymerase activity by 1.5-4 fold [195]. Interestingly, TdiF1
interacts only with the C-terminus of TdT. If any portion of TdT’s 360 C-terminal amino
acid residues is deleted, TdiF1 and the deletion mutant of TdT cannot bind to each other.
Additionally, TdiF1 is expected to reside in the nucleus (where TdT is located) due to the
presence of a nuclear localization sequence. The presence of TdiF1 in the nucleus is
confirmed by both immunofluorescence microscopy and immunostaining [195].
A second TdT interacting factor (TdiF2) has recently been identified and been
shown to downregulate the polymerase activity of TdT in vitro [185]. Through
37
immunoprecipitation, TdiF2 is demonstrated to bind TdT. However, in order for efficient
binding, the entire C-terminus of TdT must be intact, including the proline-rich and Polβlike domains. TdiF2 has also been shown to bind to single-stranded DNA. In the presence
of increasing amounts of TdiF2, the polymerase activity of TdT is observed to drop by as
much as 54% in vitro. Physically, TdiF2 is an acidic 82 kDa protein that is a member of a
family of chromatin remodeling proteins.
Recently, TdT has been found to directly bind to TReP-132, which is involved in
P450scc gene expression in steroid-hormone-producing cells or lymphoid cells.[196] The
co-expression of TdT and TReP-132 in COS7 cells showed that these proteins are colocalized within the nucleus. TReP-132 reduces the N-addition activity of TdT to 2.5% of
its maximum value in an in vitro polymerase assay in the presence of double-stranded
DNA with a 3' protrusion as a primer [196]. Thus, these results suggest that TReP-132
also downregulates the polymerase activity of TdT.
Proliferating cell nuclear antigen (PCNA) is another protein known to interact
with TdT. Like TdiF2, PCNA downregulates TdT polymerase activity by as much as
83% [195]. It has been speculated that PCNA and TdiF1 may compete for the C-terminal
region of TdT, representing a means of both positive and negative control for this enzyme
[195]. Interestingly, upon binding to DNA, TdT has been demonstrated to lose its ability
to bind both PCNA and TdiF2 [185, 197].
38
Protein phosphorylation is another key regulatory mechanism of many cellular
events. Experiments on labeling of human lymphoblastoid cells with [32P]-phosphate has
shown that TdT is phosphorylated in lymphoblastoid cells.[198] In vitro, recombinant
human TdT has also been shown to be phosphorylated by Protein Kinase C (PKC) [199].
PKC is found to localize to the nucleus in KM-3 cells [199]. In addition, fragments of
calf thymus TdT are found to be independently phosphorylated by beef heart cAMPdependant protein kinase, suggesting that calf thymus TdT may be phosphorylated at
multiple sites [101]. These phosphorylation sites were later resolved to the TdT Nterminus, for example, Ser7 and Thr19 in human TdT [103, 198]. Although it is clear that
TdTs are phosphorylated in vivo, how phosphorylation regulates the activity of TdT is not
yet clear.
TdT activity at DNA ends may also be regulated indirectly by the highly
homologous Polμ. Mahajan et al have demonstrated that, at least in vivo, TdT and Polμ
can efficiently compete for the same DNA substrate. If these proteins were both present
during the process of V(D)J recombination, it is reasonable to assume that competition
from Polμ may affect the activity of TdT during “N” region synthesis [65].
The Ku proteins may also play a role in the regulation of TdT activity. TdT has
been shown in vivo to add “N” nucleotides to double stranded DNA breaks generated
through exonuclease activity. However, the “N” regions generated in the absence of
Ku80 are unusually long [200]. This observation suggests that Ku80 may not only recruit
39
TdT to the site of V(D)J recombination but also play a role in the regulation of its
catalytic activity.
Somatic Hypermutation
As a B cell enters a germinal center of peripheral lymphoid tissue, it undergoes a
second round of antibody diversification in a process called somatic hypermutation. In
mice and humans, somatic hypermutation occurs at rates of 10-5 to 10-3 mutations per
basepair per generation which is about 106-fold higher than the spontaneous mutation
rate in most other genes [201]. The somatic mutations are mainly single base substitions,
with infrequent insertions and deletions. This process preferetially targets and mutates
WRCY (W = A or T, R =A or G, Y = T or C) and WA motifs in the rearranged
“variable” regions and its immediate flanking sequences, resulting in the generation of
high-affinity antigen binding sites, [202] consequently developing the extensive and
diverse immunoglobulin repertoire needed for survival. Variable regions that are not
rearranged are rarely seen to undergo mutations [203, 204]. The mature B cells resulting
from this process are then selected for and become memory B cells which produce
antibodies for the recognition of pathogens.
So far, the mechanism of somatic hypermutation has not been established.
Recently, somatic hypermutation was hypothesized to be initiated by a protein called
activation-induced cytidine deaminase (AID) [205, 206] based on the following facts: i)
AID, encoded by the AICDA gene, is expressed only in B lymphocytes; [207] ii) mice
40
deficient in AID are compeletely defective in somatic hypermutation and class switch
recombination; [208] iii) AID, which deaminates cytosine to uracil in DNA, peferentially
targets WRC motifs in single-stranded DNA [209]. Single-stranded DNA may arise
transiently during gene transcription; [210] iv) somatic hypermutation depends on the
active transcription of antibody genes to create the target for cytosine deamination by
AID [211]. Following initiation, the U:G mispairs in Ig DNA are either directly copied by
a DNA polymerase to form C:G to T:A transition mutations, excised by a uracil-DNA
glycosylase and then repaired through BER, or recognized by MSH2-MSH6 mismatchrecognition complex and then repired [210]. The latter two possiblities like the first also
involve DNA polymerase(s), which may exhibit low fidelity and produce mutations.
At present, it is not clear how many or which DNA polymerase(s) catalyze
somatic hypermutation. We speculate that TdT can be excluded as a candidate because it
is not a template-dependent DNA polymerase nor is it expressed in the germinal centers
of peripheral lymphoid tissues. This speculation is substantiated by the observation that
somatic hypermutation can occur in the B cells of TdT-deficient mutant mice [212].
Preferential expression in secondary lymphoid tissues [64] as well as the low templatedependant polymerization fidelity (10-3-10-5) of Pol [69] have led to the hypothesis that
this most homologous X-family DNA polymerase to TdT is an error-prone mutase, active
in somatic hypermutation [64]. However, no alterations in the somatic hypermutation
process have been found in Pol knockout mice [62].
41
Interestingly, four error-prone DNA polymerases including Pol (A-family), Polδ (Bfamily), Polε (Y-family), and Polι (Y-family) have been implicated directly or indirectly
in somatic hypermutation [5, 213] based on the following evidence: i) Pol is highly
expressed in lymphoid tissues including the spleen and germinal centers [214, 215]. A
complete deletion of the gene encoding Pol in mice leads to a reduction of overall
somatic hypermutation frequency by 60–80% and the mutation spectrum is moderately
shifted towards more transitions at both A:T and C:G basepairs; [216] ii) inhibition of the
catalytic subunit of Polδ in human B cells by specific phosphorothioate-modified
oligonucleotides impaires Ig and bcl-6 hypermutation by ~70% [217]. Expressing
antisense RNA to a portion of mouse REV3, the gene encoding Polδ, in transgenic mice
has been found to delay the generation of high affinity antibodies and to decrease the
accumulation of somatic mutations in the VH gene segments of memory B cells; [218] iii)
human Pol has an average misincorporation frequency of 10-2-10-3 [31-33, 219].
Strikingly, the fidelity of DNA incorporation by Pol
is asymmetric, with a
misincorporation rate of about 1x10-4 at a template base “A” while the incorporation of
“G” is favored by 3-11 times over “A” opposite a template base “T” [31, 32]. This
observed asymmetric fidelity is surprisingly similar to the strand bias found in Ig V
regions, [220, 221] where there are more mutations from “A” (due to misincorporations
opposite template “T”) and fewer mutations from “T” (due to accurate incorporations
opposite template “A”); iv) steady-state kinetic analyses have shown that both human
Pol [222-226] and yeast Pol [226-228] incorporate nucleotides opposite both normal
and UV-damaged DNA with a similarly low fidelity of about 10-2-10-3. When human
42
Pol
is mutated among xeroderma pigmentosum-variant patients, they have normal
immune systems and undergo somatic hypermutation, but they have an altered mutation
spectra;[229] v) expression of Polδ, Polε, and Polι in cultured Burkitt's lymphoma cells
leads to a 5-10-fold increases in heavy chain V-region mutations if co-stimulated with T
cells and IgM crosslinking, the presumed in vivo requirements for somatic
hypermutation.[213] Together, these data suggest that more than one DNA polymerase is
likely to be involved in somatic hypermutation.
43
1.9. Experience Dependence Memory Processing and TdT
In addition to V(D)J recombination, TdT has been hypothesized to play a role in
the storage of memory. As early as 1965 it was suggested that long-term memory could
be stored in the form of structural modification of synaptic connections within the brain
[230]. In addition, these structural modifications are known to require protein synthesis
[231-235]. It has been noted that the immune system and the nervous system are similar
in many ways. Most notably, both systems have the unique task of storing environmental
information that is not genetically inherited [235]. If TdT were to play a role in memory
storage, it would almost certainly have to be expressed in brain cells. Unfortunately,
reports of TdT expression in the nervous system have been mixed. For example, in 1976
Viola et. al. [236] reported that a cell lysate prepared from the cerebral cortex of the
occipital lobe of a human with no cerebral pathology was demonstrated to exhibit TdT
polymerase activity. But, in 1997 when analyzing rainbow trout cell lysates for the
presence of TdT and RAG1, reverse-transcriptase polymerase chain reaction analysis
failed to detect the presence of TdT cDNA in brain tissue [83]. However, in 2003 TdT
mRNA was in fact detected in the neurons of mouse brain tissue using an in situ
hybridization screening [235]. Specifically, TdT mRNA was found in neuronal cells of
the hippocampal formation, cerebellum, amygdala, and neocortex. These areas of the
brain have all been implicated in the storage of memory [235, 237]. In addition, the level
of TdT mRNA in mice is found to differ as a function of the environment in which they
are raised. Mice raised in enriched and highly stimulatory environments demonstrate
44
enhanced spatial discrimination learning and memory. However, transgenic mice that
have no TdT genes are found not to benefit in this way from an enriched environment
during their development [235]. It may be noted that many of the components responsible
for V(D)J recombination are also critical to neurogenesis [235, 238-242]. Therefore, one
might theorize that the TdT knockout mice used in this study may in fact be poor
“learners” not because the TdT activity is missing, but because of inhibited neurogenesis.
However, the enrichment induced improvement of learning has been shown not to be
dependent upon neurogenesis [235, 243].
45
1.10. Figures
Figure 1.1 Domain organization of six X-family DNA polymerases. The protein sequence
of each DNA polymerase is indicated by a bar, with domains differentiated using
different colors [64]. NLS denotes a nuclear localization signal motif. BRCT: BRCA1
Carboxy Terminus Domain.
46
Figure 1.2 Ternary structure of human DNA polymerase β•single nucleotide gapped
DNA•ddCTP [121]. The 8-kDa (purple), fingers (blue), palm (mixed colors), and thumb
(green) domains are shown in solid ribbon. The template 16-mer (yellow), upstream
primer 10-mer (white), and downstream primer 5-mer (white) are depicted by arrows.
The incoming ddCTP (mixed colors, ball and stick model), two Mg2+ ions (green CPK
sphere), and two Na+ ions (yellow CPK sphere) are also shown.
47
Figure 1.3 Proposed “two-divalent-metal-ion” mechanism for nucleotide incorporation
catalyzed by human DNA polymerase β [121].
48
Figure 1.4 Binary crystal structures of the Polβ-like domain (residues 148-510) of murine
TdT complexed with a brominated 9-mer at 3.0 Å [88]. The 8-kDa (purple), fingers
(blue), palm (mixed colors), and thumb (green) subdomains are shown in solid ribbon.
Three β-sheets β3, β4, and β5 are labeled as 3, 4, and 5, respectively. Loop 1 and Loop 2
are shown in yellow. Four 3’-nucleotides of the 9-mer primer are ordered and drawn in
the ball and stick model. Mg2+ (green) and Na+ (purple) ions are shown as CPK spheres.
49
Figure 1.4
50
Figure 1.5 Binary crystal structure of the Polβ-like domain (residues 148-510) of murine
TdT complexed with a ddATP-Co2+ at 3.0 Å [88]. All subdomains are depicted in the
same manner as those in Figure 1.4. Three active site residues Asp343, Asp 345, and
Asp434 (white color, stick and ball model) as well as Lys403 (yellow color, stick model),
Trp450 (yellow color, stick model), Co2+ (yellow CPK sphere), Na+ (purple CPK sphere),
and an incoming ddATP (mixed colors, stick and ball model) are also shown.
51
Figure 1.6 Minimal kinetic mechanism for polymerization catalyzed by DNA
polymerases. E and E* denote a polymerase before and after conformational change,
respectively. PPi represents pyrophosphate.
52
Base
O
O
O
Base
O
O
HO P O P O P O
OH
OH
OH
O
O
O
HO P O P O P O
OH
HO
OH
dNTP
OH
HO
OH
rNTP
H2 N
O
O
O
O
O
N
O
O
HO P O P O P O
OH
OH
N
N
N
O
O
O
N
N
HO P O P O P O
OH
OH
d4TTP
OH
OH
OH
cordycepin 5'-triphosphate
Base
O
O
O
O
O
OH
OH
OH
OH
ddNTP
O
O
O
O
OH
OH
O
OH
OH
Base'
Base
O
O
O
O P O P O P O P
OH
HO
OH
OH
O
O
OH
HO
dinucleoside 5',5'-tetraphosphate
Figure 1.7 Chemical structures of nucleotide analogs.
53
Base
O
OH
p-nitrophenylethyl triphosphate
OH
O
OH
HO P O P O P O CH2CH2
NO2
p-nitrophenyl triphosphate
O
O
-D-dNTP
HO P O P O P O
OH
O
HO P O P O P O
HO P O P O P O
OH
O
NO2
Figure 1.8 Crystal structure of anti-lysozyme Fab and hen egg white lysozyme [244]. In
the domain structure of an Ig molecule, the variable region in the heavy chain is
composed of each of V, D, and J gene segments while the variable region in the light
chain possesses a V and a J gene segment. The intra and inter chain disulfide bonds are
denoted as -S-S-.
54
Figure 1.8
55
A
(continued)
Figure 1.9 T cell receptor encoded by tandemly arranged clusters of V, D, and J gene
segments. (A) The constant region (C) gene segments follow the joining (J) gene
segments. The TCR
chains do not possess the dependant (D) gene segments. (B)
Schematic diagram of a T cell receptor. Each TCR chain is composed of a variable and a
constant region.
56
Figure 1.9 (continued)
B
57
A
(continued)
Figure 1.10 V(D)J Recombination. (A) A germline Ig heavy chain becomes a functional
Ig heavy chain after V(D)J recombination. The colored boxes denote clustered coding
segments. One of each of the V, D, and J segments are joined to form the variable region
of a functional Ig. Random nucleotides (not shown) are added to the junctions between V,
D and J segments. The constant regions are not involved in V(D)J recombination. (B)
Coding segments V and D are associated with recombination signal sequences 23-RSS
and 12-RSS, respectively. The 23-RSS and 12-RSS are enlarged for clarity. (C) A V
segment is joined by a D segment through cleavage by RAG proteins and processing by
NHEJ proteins. TdT adds random nucleotides to the junction between V and D segments.
58
Figure 1.10 (continued)
B
C
59
Figure 1.11 Proposed mechanism for the “N region” formation at the junction between a
V and a D segment. “P” nucleotides are shaded in grey.
60
1.11. Tables
Relative Incorporation Ratea
dNTP
dATP
dGTP
dTTP
dCTP
Mg2+
1.00
1.63
0.10
0.13
Co2+
1.13
2.30
16.46
14.39
Table 1.1 Effect of metal ions on the incorporation rate of each dNTP catalyzed by TdT
[135]. aIncorporation rates are relative to the rate of dATP (245 nmol/mg TdT/hour).
61
Chapter 2: Kinetic investigation of the inhibitory effect of gemcitabine on DNA
polymerization catalyzed by human mitochondrial DNA polymerase
2.1. Introduction
Many nucleoside analogs are potent anticancer and antiviral small molecules.
Among fifteen Food and Drug Administration-approved nucleoside analogs, gemcitabine
or 2’-deoxy-2’,2’-difluorocytidine (dFdC, Figure 2.1) is an anti-cancer drug which is
clinically used for the treatment of non-small cell lung cancer [245], pancreatic cancer
[246], metastatic breast cancer [247], and ovarian cancer [248]. It has also shown
promising efficacy for the treatment of other solid tumors and hematological
malignancies [249-256] suggesting more widespread use in the future. In addition to its
use as a monotherapy, gemcitabine is often most effective when used as part of a
combination therapy, frequently with platinum-based and topoisomerase-targeted
chemotherapeutic agents [257-259].
Gemcitabine is administered in the form of a biologically, inactive prodrug that
first permeates the cellular membrane by facilitated diffusion [260, 261] almost
exclusively via the human equilibrative nucleoside transporter number 1 [261, 262].
62
Following transport, dFdC is metabolized to the biologically active monophosphorylated
form (dFdCMP) by deoxycytidine kinase, the rate limiting step in the activation of
gemcitabine [263]. Subsequently, dFdCMP is further phosphorylated to form the
cytotoxic metabolites gemcitabine diphosphate (dFdCDP) and gemcitabine triphosphate
(dFdCTP) by cellular kinases. It has been shown that dFdCTP competes effectively
against endogenous dCTP for incorporation into genomic DNA [264], against CTP into
RNA [265], and that the proofreading exonuclease activity of human DNA polymerase
is essentially unable to remove dFdCMP once incorporated into DNA [266].
Interestingly, dFdCTP incorporation by human DNA polymerase α results in “masked
termination” of DNA synthesis where, following a single dFdCTP incorporation into
DNA, the primer is extended by only one additional dNTP before polymerization is
inhibited [264, 266]. However, in addition to being incorporated into DNA and RNA,
dFdCDP and dFdCTP are known to inhibit ribonucleotide reductase, thereby significantly
decreasing cellular dCTP concentrations and leading to increased phosphorylation of
dFdCDP [267, 268]. Furthermore, high concentrations of dFdCTP inhibit CTP
synthetase, thereby reducing dCTP and CTP pools yet further [269, 270]. Reduced
competition from smaller dCTP pools makes dFdCTP incorporation into DNA and RNA
more probable, promotes cell cycle arrest and apoptosis, and inhibits DNA repair [267].
Furthermore, dFdCMP and dFdCTP also inhibit dCMP deaminase, the major pathway by
which dFdCMP is eliminated [271]. The combined synergistic effect of these inhibitory
activities is termed “Self-Potentiation” and is illustrated in Figure 2.2 [272, 273].
63
Moderate toxicity of gemcitabine has been observed in cancer patients with
peripheral
neuropathy
[274,
275]
and
hematological
dysfunction
in
which
myelosuppresion frequently emerges as the dose-limiting factor [248]. Nonhematological toxicities are also common and include lethargy, mild flu-like symptoms,
pruritic skin rash, nausea, edema and vomiting [276]. Pulmonary toxicity resulting from
gemcitabine therapy is observed in nearly 10% of patients and ranges in severity from
dyspnea, which occurs shortly after administration of gemcitabine and is short lived
[277], to Acute Respiratory Distress Syndrome (ARDS) which is frequently fatal [278].
Most of these observations are not surprising considering the toxicities of other antiviral
nucleoside analogs, many of which mimic the symptoms of mitochondrial diseases
caused by genetic defects [279]. Moreover, loss of mitochondrial DNA (mtDNA) and
changes in mitochondrial ultrastructure have been observed in cell culture studies after
treatment with nucleoside analogs [280-284]. The mitochondrial toxicity of an antihepatitis B nucleoside analog fialuridine (FIAU), developed by Lilly in the early 1990’s,
is a good example: FIAU killed five patients in the clinical trials [285, 286]. The
interference of mtDNA synthesis by gemcitabine could contribute to the observed
toxicities of this anticancer drug in cancer patients. Previously, we have used pre-steady
state kinetic methods to evaluate the mitochondrial toxicity of several anti-HIV
nucleoside analogs including (R)-9-(2)Phosphonyl(methoxypropyl)adenine, 3'-azido-3'deoxythymidine, 2',3-dideoxycytosine, 2'-3'-dideoxy-3'-thiacytidine, 2',3'-dideoxyinosine,
2'3'-didehydro-3'-deoxythymidine,
and
hydroxymethyl)-2-cyclopenten-1-yl)-6H-purine-6-one,
64
(-)-cis-2-amino-1,9-dihydro-9-(4and
the
anti-hepatitis
B
nucleoside analog FIAU with recombinant human DNA polymerase holoenzyme (Pol )
[8], and our kinetic data correlate well with the observed toxicities of these drugs in vivo
[287]. Therefore, to evaluate the potential mitochondrial toxicity of gemcitabine, we
again employed pre-steady state kinetic methods to evaluate the incorporation, extension,
and excision of gemcitabine catalyzed by Pol . In addition, we examined whether or not
an incorporated gemcitabine as a template base could lead to mutations in the next round
of mitochondrial DNA synthesis. Our data provide direct evidence demonstrating the
mitochondrial toxicity of gemcitabine.
65
2.2. Materials
What follows is a list of the reagents used for these experiments and their sources.
[γ-32P]ATP, GE Healthcare (Piscataway, NJ); Bio-Spin columns, Bio-Rad Laboratories
(Hercules, CA); dNTPs, Gibco-BRL (Rockville, MD); dFdCTP, donated by Trilink
Biotechnologies INC. (San Diego, CA); OptiKinase, USB (Cleveland, OH).
Optimized Reaction Buffer G
50 mM Tris-Cl, pH 7.5 at 37 °C, 100 mM NaCl, and 2.5 mM MgCl2. Note that all
concentrations listed in this paper refer to the final concentration after mixing unless
otherwise noted.
Optimized Reaction Buffer L
50 mM Tris-Cl, pH 8.4 at 37 °C, 100 mM NaCl, 5 mM MgCl2, 0.1 mM EDTA, 5
mM DTT, 0.1 mg/ml BSA, and 10% glycerol.
Optimized Reaction Buffer M
50 mM HEPES, pH 8.0 at 25 °C, 12 mM NaCl, 8.75 mM MgCl2, 0.2 mM EDTA,
5 mM DTT, 0.1 mg/mL BSA, and 10% glycerol.
66
Purification of Human Polymerase Gamma Subunits
Expression and purification of wild-type human DNA polymerase γ, its
exonuclease-deficient mutant E200A, and the small accessory subunit were carried out as
described previously [8, 9].
Synthetic Oligodeoxyribonucleotides
All DNA substrates not containing gemcitabine were purchased from Integrated
DNA Technologies (Coralville, IA) and purified by denaturing polyacrylamide gel
electrophoresis
(17%
acrylamide,
8M
urea).
Concentrations
of
synthetic
oligodeoxyribonucleotides were determined from their UV absorbance at 260 nm.
Primers were 5’-[32P]-labeled by incubation with [γ-32P]ATP and OptiKinase at 37 °C for
one hour. Remaining [γ-32P]ATP was subsequently removed by size exclusion
chromatography in a Bio-Spin 6 column. All primers were annealed to their respective
templates in a 1:1.15 (primer:template) molar ratio by heating the mixture to 95 °C for 10
minutes and then slowly cooling to room temperature over approximately 6 hours.
Synthetic Oligodeoxyribonucleotides Containing Gemcitabine
To create two DNA primers and a template that contain gemcitabine in their
sequence context, a primer extension and ligation strategy was employed. Primer 23Fmer (Table 2.1), which is a 23-mer containing a 3’-dFdCMP, was synthesized by mixing
DNA 22/41-mer (Table 2.1) with 5 μM dFdCTP and human DNA polymerase
[69], a
template-directed DNA polymerase capable of efficiently incorporating dFdCTP, in
67
reaction buffer M. The reaction was conducted for 2 hours at 25 °C yielding maximum
conversion of the 22-mer primer to 23F-mer. A similar reaction was performed to
synthesize primer 24FG-mer (Table 2.1), except following the 2 hour incubation with
dFdCTP, 20 μM dGTP was added to the reaction mixture and the reaction was allowed to
continue for an additional 5 minutes. Products of these reactions were purified using
denaturing PAGE.
To synthesize template 41F-mer (Table 2.1), the DNA substrate 21-19/35-mer
(Table 2.1) was incubated with 8 μM dFdCTP and human DNA polymerase
[288], a
template directed, gap-filling DNA polymerase capable of efficiently incorporating
dFdCTP, for 5 minutes at 37 °C in reaction buffer L, to form 21F-19/35-mer. Unreacted
dFdCTP was then removed using gel filtration (Bio-Spin 6, Bio-Rad). The DNA solution
was heated to 95 °C for 10 minutes and then slowly cooled to room temperature over six
hours to re-anneal 21F-19/35-mer (a nicked DNA substrate). A solution of 10 mM
MgCl2, 1 mM ATP, and T4 DNA ligase (11 units/μl) was added to the annealed DNA
solution to ligate the nicked DNA for 7 minutes at 37 °C. The resulting 41F-mer was
purified from the mixture using denaturing PAGE.
68
2.3. Methods
Single-Turnover Nucleotide Incorporation Assay
All assays using Polγ were carried out at 37 °C in buffer G containing 2.5 mM
MgCl2. For single nucleotide incorporation assays, Polγ (90 nM) and its cofactor SSU
(450 nM) were combined (1:5 molar ratio) and preincubated on ice in buffer G for 20
minutes to form human Pol holoenzyme. Next, 30 nM of a DNA substrate containing a
5’-[32P]-labeled DNA primer was added to the reconstituted holoenzyme (3:1 molar ratio,
holoenzyme:DNA) and incubated on ice for an additional 20 minutes. The single
nucleotide incorporation reaction was initiated by the addition of dNTP and 2.5 mM
MgCl2 in buffer G using a rapid chemical quench apparatus (KinTek, Clarence, PA).
After varying reaction times at 37°C, the reactions were quenched by the addition of 0.37
M EDTA (Figure 2.3).
Excision Reactions
For the 3’5’ exonuclease assay, wild-type Polγ (100 nM) and SSU (500 nM) in
buffer G were first preincubated on ice for 20 min to form human Pol holoenzyme and
then mixed with 5’-[32P]-labeled DNA substrate (75 nM) in the absence of Mg2+. The
3’5’ exonuclease reaction was initiated by the addition of 2.5 mM MgCl2 in buffer G
using a rapid chemical quench apparatus. After varying reaction times at 37 °C, the
reactions were quenched by the addition of 0.37 M EDTA. The concentration of
69
remaining full-length primer as a function of time was quantitated and the exonuclease
reaction time course (Figure 2.4) was fit to Eq. 3 to yield an excision rate constant (kexo).
Running Start Nucleotide Incorporation Assay
For the running start nucleotide incorporation assay, a DNA substrate (30 nM)
was first preincubated with a solution of Polγ (90 nM) and SSU (450 nM) in buffer G as
described above. This solution was rapidly mixed with MgCl2 (2.5 mM) and dNTPs (100
μM each). The primer elongation at various times was stopped by the addition of 0.37 M
EDTA.
Product Analysis
Products of the polymerase and exonuclease reactions were separated by
sequencing gel electrophoresis (17% acrylamide, 8 M urea, 1X TBE running buffer) and
quantitated using a Phosphorimager 445 SI (Molecular Dynamics).
Data Analysis
Kinetic data were fit via non-linear regression using KaleidaGraph (Synergy
Software). Data from single-turnover nucleotide incorporation assays were fit to a single
exponential (Eq. 1) to obtain an observed incorporation rate constant (kobs). The dNTP
concentration dependence of kobs was fit to a hyperbolic equation (Eq. 2) to yield both the
equilibrium dissociation constant (Kd) and the maximum nucleotide incorporation rate
constant (kp). Single-phase exonuclease reaction time courses were fit to a single
70
exponential equation (Eq. 3) to yield the exonuclease rate constant (kexo). Biphasic
exonuclease reaction time courses were fit to a double exponential (Eq. 4) to yield kexo,1
and reaction amplitude A1 in the fast phase, and kexo,2 and reaction amplitude A2 in the
slow phase.
[Product] = A[1 – exp(- kobst)]
Eq. 1
kobs = kp[dNTP]/{[dNTP] + Kd}
Eq. 2
[Product] = A[exp(- kexot)]
Eq. 3
[Product] = A1[exp(- kexo,1t)] + A2[exp(- kexo,2t)]
Eq. 4
71
2.4. Results
Determination of the Pre-Steady State Kinetic Parameters for dFdCTP and dCTP
Incorporation
The kinetic mechanism of DNA polymerization catalyzed by human DNA
polymerase γ holoenzyme has been established by our pre-steady state kinetic analysis [8,
9, 142, 289]. This mechanism has shown that an incoming dNTP binds to the Pol •DNA
binary complex to establish a rapid equilibrium prior to nucleotide incorporation [9, 142].
Therefore, the ground-state equilibrium dissociation constant of an incoming dNTP (Kd)
and its maximum incorporation rate constant (kp) can be measured by observing the
nucleotide concentration dependence of the observed single-turnover rate constant (kobs)
[289]. To examine the toxicity of gemcitabine toward human mitochondria at a molecular
level, we first determined the substrate specificity (kp/Kd) of dFdCTP catalyzed by human
Pol . Because a nucleotide analog is usually incorporated slowly, and its incorporation
rate constant is comparable or smaller than the dissociation rate constant of DNA from
the enzyme•DNA binary complex, the burst phase either is insignificant or does not exist.
Thus, the experiments to measure the kp/Kd value of dFdCTP were performed with Pol
in molar excess over DNA to allow the direct observation of nucleotide incorporation in a
single pass of the reactants through the catalytic cycle without complications resulting
from the steady-state formation of products [290]. On the other hand, because the wildtype Pol has highly efficient 3’5’ exonuclease activity [142] which excises a primer
from its 3’-terminus and thereby complicates the slow incorporation of dFdCTP, we
72
decided to use Polγ E200A, a well characterized single point mutant at the 3’5’
exonuclease active site of human Pol , to determine the kinetics of dFdCTP
incorporation. This mutant incorporates normal nucleotides with similar kinetics as the
wild-type Pol but is exonuclease deficient [287].
To measure the pre-steady state kinetic parameters for dFdCTP incorporation, a
preincubated solution of Polγ E200A (90 nM large subunit, 450 nM accessory subunit)
and 5’-[32P]-labeled 22/41-mer (30 nM, Table 2.1) was mixed and reacted with increasing
concentrations of dFdCTP at 37 ˚C for various times. An autoradiograph gel image in
Figure 2.3 part (A) shows that E200A gradually incorporated dFdCTP and thereby
elongated the primer 22-mer to 23-mer. Each time course of product formation in Figure
2.3 part (B) was fit to Eq.1 (Section 2.3) to yield an observed single turnover rate
constant, kobs. The kobs values were then plotted as a function of dFdCTP concentration
(Figure 2.3 part (C)). The data were fit to Eq. 2 (Section 2.3) to yield a kp of 2.0 ± 0.3 s-1
and a Kd of 21 ± 7 μM (Table 2.2). The substrate specificity of dFdCTP was calculated to
be 0.095 μM-1s-1. Similarly, we measured the kp (37 ± 2 s-1) and Kd (0.9 ± 0.2 μM) for the
incorporation of dCTP into the 22/41-mer (Table 2.1) under single-turnover reaction
conditions (data not shown). These kinetic parameters agree well with those that we
measured under burst reaction conditions [289], thus validating our single-turnover
approach. The incorporation efficiency of dCTP was then calculated to be 41 μM-1s-1.
Thus, the discrimination, defined as the efficiency ratio of (kp/Kd)dCTP/(kp/Kd)dFdCTP,
73
exhibited by the polymerase activity of human Pol against dFdCTP is 432-fold (Table
2.2).
Measurement of the Excision Rate Constants of Matched 3’-dFdCMP and 3’-dCMP
The 3’5’ exonuclease activity of human Polγ recognizes mismatched 3’-base(s)
in a DNA primer and rapidly excises 1-7 mismatched bases at a rate of 1-9 nucleotides
per second but slowly excises the matched 3’-terminal base of a primer [142]. Although
paired with template base dG, the incorporated 3’-dFdCMP in a primer may be subject to
this 3’5’ exonucleolytic proofreading mechanism and thereby excised. We first
synthesized and purified primer 23F-mer (Section 2.3, Table 2.1) and then measured the
excision rate constant, kexo, of 3’-dFdCMP from substrate 23F/41-mer (Table 2.1) by the
wild-type Polγ under single-turnover reaction conditions. A preincubated solution of the
wild-type Polγ (100 nM large subunit, 500 nM accessory subunit) and 75 nM 5’-[32P]labeled DNA (23/41-mer or 23F/41-mer) in buffer G was rapidly mixed and reacted with
2.5 mM MgCl2 for various time intervals prior to being quenched by 0.37 M EDTA. The
concentration of the remaining full-length primer versus reaction time was plotted and fit
to Eq. 3 (Section 2.3), yielding the kexo of 0.06 ± 0.02 s-1 and 0.0011 ± 0.0001 s-1 for the
23/41-mer and 23F/41-mer, respectively (Figure 2.4, Table 2.3). The kexo of 23/41-mer
DNA was similar to 0.05 ± 0.01 s-1 measured previously with a perfectly matched
substrate 25/45-mer (Table 2.1) [142]. Interestingly, the excision of the matched 3’dFdCMP moiety by the wild-type Pol is 50-fold slower than the excision of matched 3’-
74
dCMP. This suggests that an incorporated dFdCMP can escape the editing process and be
embedded into mtDNA.
Measurement of the Extension Efficiency of a Primer Terminated with 3’-dFdCMP
It is possible that an incorporated dFdCMP moiety on the 3’-terminus of a DNA
primer could significantly alter the ability of Polγ to extend that primer. To examine this
possibility, we measured the kinetic parameters for the incorporation of correct dGTP
into 23F/41-mer (Table 2.1) catalyzed by E200A under single-turnover conditions as
described above (data not shown). dGTP was incorporated with a kp of 1.5 ± 0.1 s-1, a Kd
of 7.2 ± 0.9 μM, and a kp/Kd of 0.21 μM-1s-1 (Table 2.2). In comparison, a matched dGTP
is incorporated into normal 25/45-mer (Table 2.1) with a kp of 37 ± 2 s-1, a Kd of 0.8 ± 0.1
μM, and a kp/Kd of 45 μM-1s-1 [289]. Thus, the 3’-dFdCMP decreased the incorporation
efficiency of the first downstream nucleotide by 214-fold, suggesting that an incorporated
gemcitabine inhibits primer extension. Such an inhibitory effect may persist beyond one
nucleotide. To examine if an embedded gemcitabine inhibits the incorporation of the
second downstream nucleotide, we prepared primer 24FG-mer (Section 2.3). We then
measured the kinetic parameters of correct dTTP incorporation into 24FG/41-mer (Table
2.1) catalyzed by E200A (data not shown). Under single-turnover reaction conditions as
described above, dTTP was incorporated with a kp of 2.8 ± 0.1 s-1, a Kd of 0.5 ± 0.1 μM,
and a kp/Kd of 5.6 μM-1s-1 (Table 2.2). In comparison, given a canonical DNA substrate
25/45-mer (Table 2.1), correct dTTP is incorporated with a kp of 25 s-1, a Kd of 0.6 μM,
and a kp/Kd of 39 μM-1s-1 [289]. Thus, primer 3’-dFdCMP lowered the second
75
downstream nucleotide incorporation efficiency by 7-fold. Interestingly, the third
nucleotide dCTP downstream from the embedded dFdCMP was incorporated into
25FGT/41-mer with similar efficiency as it was incorporated into normal 22/41-mer (data
not shown). Therefore, an incorporated dFdCMP in the DNA primer only inhibits two
downstream nucleotide incorporations, especially the first one. However, it does not
terminate primer elongation.
Running Start Primer Extension Assays
To investigate whether or not a dFdC lesion embedded in a DNA template affects
DNA synthesis catalyzed by human Polγ, we first synthesized template 41F-mer (Table
2.1) as described in section 2.3. With a primer 15-mer, a running start primer elongation
assay was performed to evaluate the ability of the wild-type Polγ (90 nM) to bypass the
template dFdCMP lesion in 15/41F-mer (30 nM, Table 2.1) in the presence of four
dNTPs (100 M each). In comparison, a similar running start assay was also performed
with an undamaged control substrate 15/41-mer (30 nM, Table 2.1). With the 15/41-mer,
the wild-type Polγ holoenzyme was able to extend the 15-mer primer to the full length
41-mer in 1 sec (Figure 2.5 part (A)). This rate is consistent with the average maximum
rate constant (38 s-1) of single nucleotide incorporation into normal 25/45-mer (Table 2.4)
[289]. In contrast, the wild-type Polγ paused opposite the dFdC lesion and one base
downstream of the dFdC lesion in the 15/41F-mer (Figure 2.5 part (B)). Consequently,
the full-length product 41-mer was formed in 3 seconds, rather than just 1 second
observed with control 15/41-mer (Figure 2.5 part (A)). Although the synthesis of the 4176
mer was delayed, Figure 2.5 part (B) shows that the wild-type Polγ eventually bypassed
the dFdC lesion.
To examine whether or not the 3’5’ exonuclease activity of human Polγ plays
any role in the bypass of the dFdC lesion, we performed the same running start primer
elongation assays with Polγ E200A. Figure 2.5 parts (C) and (D) show that the product
formation patterns with both 15/41-mer and 15/41F-mer are almost identical to the
patterns seen with the wild type enzyme seen in parts (A) and (B). Thus, the 3’5’
exonuclease activity appears not to recognize template base dFdC as a lesion and is
dispensable for its bypass.
77
Measurement of the Excision Rate Constant of Primer 3’-dNMP Opposite Template Base
dFdC
To quantitatively interrogate the effect of an embedded dFdCMP moiety on the
3’5’ exonuclease activity of Polγ, we synthesized two DNA substrates (Section 2.3):
20/41F-mer and 20T/41F-mer (Table 2.1) which contain a 3’ terminal correct base pair
and a mispair, respectively. Under single-turnover conditions, the wild-type Polγ
holoenzyme was found to excise the primer 3’-bases of the 20/41F-mer with a kexo of
0.028 ± 0.006 s-1 (data not shown). This kexo value is about 2-fold smaller than the kexo of
0.06 ± 0.02 s-1 observed with a normal substrate 23/41-mer (Table 2.3). In comparison,
the time course of the cleavage of the 20T/41F-mer by the wild-type Pol under singleturnover reaction conditions is biphasic and was fit to Eq. 4 (Section 2.3) to yield a kexo,1
of 0.2 ± 0.2 s-1 and an amplitude of (17 ± 8)% in the fast phase and a kexo,2 of 0.008 ±
0.002 s-1 and an amplitude of (83 ± 8)% in the slow phase (data not shown). Similar
biphasic kinetics are also observed previously with the cleavage of normal DNA
containing a single 3’-mismatched base but with larger kexo values in the fast phase (1.1 s1
) and slow phase (0.04 s-1) [289]. These data confirm that template base dFdC slows
down the proofreading activity of human Pol .
Measurement of Incorporation Efficiency of Nucleotides Opposite Template dFdC
The reason for Polγ's strong pause seen in Figure 2.5 parts (B) and (D), is
probably because the dFdC lesion altered local template structure and significantly
decreased the incorporation efficiencies of adjacent nucleotides. To evaluate this
78
hypothesis, we measured the incorporation efficiencies of correct nucleotides into
18/41F-mer, 19/41F-mer, 20/41F-mer, 21/41F-mer, and 22/41F-mer (Table 2.1)
catalyzed by E200A under single-turnover reaction conditions (data not shown). The
measured kinetic parameters in Table 2.4 indicate that, relative to the corresponding
values with normal 25/45-mer (Table 2.1) [289], the ground-state binding affinity of a
correct incoming nucleotide is up to two orders of magnitude lower at the two strong
pause sites but is within 5-fold at the non-pause sites. In comparison, the maximum
nucleotide incorporation rate constant drops by four orders of magnitude at the second
strong pause site while the kp values at other positions are 5-30 fold lower than the
corresponding parameters with the 25/45-mer (Table 2.4). As expected, the substrate
specificity (kp/Kd) values are more informative in demonstrating why Polγ paused at the
two positions in Figure 2.5 parts (B) and (D). Before encountering the template base
dFdCMP, Polγ E200A incorporated correct dGTP into 18/41F-mer with only 24-fold
lower substrate specificity than it did with normal 25/45-mer (Table 2.4). The efficiency
ratio increases from 24 to 1,047 when Polγ E200A incorporated dGTP opposite dFdCMP
and to 2,053 when Polγ E200A incorporated dTTP into 20/41F-mer to extend the primer
and bypass dFdCMP. These significant decreases in the efficiency ratio indicate that Polγ
was inefficient at incorporating nucleotides at these two positions and paused in Figure
2.5 parts (B) and (D). After Polγ bypassed the dFdCMP, the efficiency ratios for the next
two downstream nucleotide incorporations (Table 2.4) decrease to 132 and 4.8,
respectively. Although we did not measure the substrate specificity of dGTP
incorporation into 23/41F-mer, we expect the inhibitory effect of an embedded
79
gemcitabine in a template will disappear at this position and downstream. Notably, based
on the low nucleotide incorporation efficiency with 21/41F-mer (Table 2.4), E200A was
expected to pause at this position. However, the pause is not obvious in Figure 2.5 part
(D). This discrepancy is because the kp of 1.1 s-1 is relatively fast and the reaction times
are relatively long.
Measurement of Nucleotide Incorporation Fidelity at the Pause Sites
Opposite dFdCMP, Polγ may choose mismatched dNTPs over dGTP. To examine
this possibility, each dNTP (200 μM) was reacted individually with a preincubated
solution of E200A and 19/41F-mer (Table 2.1) for 15 sec or 1 hour. Figure 2.6 part (A)
shows that E200A preferred to incorporate dGTP opposite dFdCMP at both time
intervals. Interestingly, dTTP was misincorporated multiple times. This suggests that
E200A may have a high tendency to misincorporate dTTP. To check if this was the case,
we measured the kp/Kd for dTTP incorporation into 19/41F-mer under single-turnover
conditions (data not shown). The calculated fidelity in Table 2.5 indicates that E200A
only favors dGTP over dTTP by 3,071 fold, which is much lower than the 6.4x105 fold
observed with normal 25/45-mer [291]. Thus, the fidelity of nucleotide incorporation
opposite dFdCMP was lowered by 200-fold.
To further examine the effect of template dFdCMP on polymerase fidelity, we
tested whether or not Polγ was error prone when extending primer 20-mer to 21-mer.
Opposite template base dAMP, E200A incorporated correct dTTP more efficiently than
80
incorrect nucleotides (Figure 2.6 part (B)), and dATP was the most favored incorrect
nucleotide. We further measured the kinetic parameters of dATP misincorporation into
20/41F-mer under single-turnover conditions (data not shown). E200A favored correct
dTTP over incorrect dATP by only 396-fold (Table 2.5), which is 707-fold lower than the
corresponding fidelity (280,000) observed with canonical 25/45-mer [291].
81
2.5. Discussion
Kondering et. al. [292] have solved the solution-phase structure of an Okazaki
fragment (12-basepairs) with an internally embedded dFdCMP using NMR. This
structure reveals the following perturbations caused by gemcitabine: i) the ribose ring of
the dFdCMP moiety forms a 3’-endo pucker as opposed to the canonical 2’-endo pucker
of a dCMP moiety; ii) the highly electronegative geminal difluoro group increases the
electron density in its vicinity; iii) the two fluorine atoms in dFdCMP are physically
larger than the corresponding hydrogen atoms in dCMP. These factors are predicted to
affect DNA polymerization catalyzed by DNA polymerases including human DNA
polymerase holoenzyme as studied in this paper.
Inhibition of Mitochondrial DNA Synthesis by Gemcitabine as an Incoming Nucleotide
The pre-steady state kinetic data in Table 2.2 reveal that relative to dCTP,
dFdCTP was incorporated into normal 22/41-mer by Pol with an 18.5-fold lower kp
while the Kd was 23-fold higher, leading to a 432-fold lower kp/Kd. These kinetic
differences can be rationalized as follows. Although dFdCTP is not the same as
embedded dFdCMP in DNA, we assume that dFdCTP initially adopts the 3’-endo pucker
once it is bound to form the ground-state ternary complex E DNA dNTP. In order for the
phosphodiester bond formation to occur, the conformation of the bound dFdCTP has to
be converted to the canonical 2’-endo pucker to allow proper alignment of the primer 3’OH and the
–phosphate of dFdCTP. The energy penalty for this conformational
82
conversion should slow the incorporation of dFdCTP. Moreover, the electronwithdrawing difluoro group decreases the electron density of the triphosphate of dFdCTP
and thereby reduces the reactivity of the –phosphate moiety during phosphodiester bond
formation. This difluoro group also increases the size of dFdCTP which may cause steric
clashes with the polymerase active site residues. Relative to dCTP, the altered
conformation, electrostatics, and physical size likely weaken the interactions between
dFdCTP, template base dGMP, and polymerase active site residues, hence the lower
binding affinity of dFdCTP.
Although the efficiency ratio in Table 2.2 defines dFdCTP as a 432-fold less
efficient substrate than dCTP, the incorporation probability of dFdCTP relative to dCTP
in vivo, defined as {[dFdCTP]/[dCTP]}x[(kp/Kd)dFdCTP/[(kp/Kd)dCTP)], is likely to be high
for the following reasons. “Self Potentiation” activities of gemcitabine (Figure 2.2) [272,
273] are known to result in a dramatic cytoplasmic accumulation of dFdCTP, thereby
enhancing the cellular concentration ratio of [dFdCTP]/[dCTP]. For example, Heinemann
et. al. found that mammalian cells exposed to 100 μM gemcitabine for 4 hours showed
that cellular pools of dCTP were reduced by 50% and that the cytoplasmic concentration
of dFdCTP had risen to over 1 mM [270]. Since relative sizes of individual dNTP pools
in the cytosol and mitochondria are similar [293], we expect that in mitochondria,
dFdCTP also has a much higher concentration than dCTP. Assuming the mitochondrial
concentration ratio [dFdCTP]/[dCTP] to be 20, dFdCTP will be incorporated once every
22 incorporations of dCTP into mitochondrial DNA. Based on the size of the human
83
mitochondrial genome (16.6 kilobases) and assuming 1 in 4 template bases is dG,
dFdCTP is projected to be incorporated by Pol at a rate of 188 times per cycle of
mitochondrial genome replication.
Once incorporated, the primer 3’-dFdCMP is likely to adopt the 3’-endo pucker
conformation as observed for the embedded gemcitabine [292]. The electron density of
its 3’-hydroxyl group should be lowered by the electron-withdrawing fluorine group.
These factors are expected to lower the activity of dFdCMP-terminated primer and
inefficient incorporation of the next coming nucleotide. This hypothesis is confirmed by
the fact that correct dGTP was incorporated into 23F/41-mer (Table 2.1) with a 25-fold
smaller kp and a 214-fold lower catalytic efficiency than it was added into normal 25/45mer (Table 2.2) [289]. Thus, the incorporated gemcitabine on a primer 3’-terminus
significantly inhibits the next nucleotide incorporation. This inhibitory effect was reduced
to 7-fold for the second downstream nucleotide incorporation (Table 2.2) and was not
observed for the third and further downstream nucleotide incorporation (data not shown).
Interestingly, human DNA polymerases
and
have also been found to inefficiently
incorporate dFdCTP and then extend it by only one nucleotide [266].
Incorporated dFdCMP Eludes Editing Mechanism
The 3’5’ exonuclease activity of Polγ may recognize and excise incorporated
gemcitabine as a lesion due to its different conformation and electrostatics in comparison
to dCMP. However, Table 2.3 shows that the excision of primer 3’-dFdCMP in 23F/4184
mer (0.0011 s-1) was 51-fold slower than the excision of a corresponding 3’-dCMP in the
control substrate 23/41-mer. Interestingly, the exonuclease activity of human DNA
polymerase ε is also unable to remove an incorporated 3’-dFdCMP [266]. At an
intracellular nucleotide concentration of 100 M, E200A incorporated dGTP into 23F/41mer with a kp of 1.5 s-1 (Table 2.2). Based on the principle of kinetic partitioning, the
probability of exonuclease editing of dFdGMP, kexo/(kexo + kp) = 0.0011/(0.0011 + 1.5), is
calculated to be 0.07%. This probability is much lower than 80% observed with the
correction of a mismatch [142]. Furthermore, although the extension of a gemcitabineterminated primer is 25-fold slower than the extension of normal DNA, the reduced
excision rate constant (kexo) and extremely low editing possibility will increase the
likelihood of an incorporated dFdCMP to be embedded into mtDNA if they are not
removed by other DNA repair mechanisms. Given that mtDNA repair is limited and
inefficient [294], persistence of dFdCMP within mtDNA is predicted to be very likely.
This hypothesis is supported by the observation that extracted cellular DNA contains
dFdCMP in internucleotide linkages after cells are treated with gemcitabine [266].
Inhibition of Mitochondrial DNA Synthesis by Gemcitabine as a Template Base
Figure 2.5 parts (B) and (D) show that human Polγ holoenzyme, both the wildtype and E200A, paused strongly at the position opposite template base dFdCMP and at
the next template position. The kinetic data in Table 2.4 reveal that the efficiency ratio
correlates well with the pausing pattern. The efficiency ratio (2,053) is the poorest when
Polγ attempted to extend the 20/41F-mer, the strongest pause site (Figure 2.5 parts (B)
85
and (D)). The 2nd strongest pause site, where Polγ attempted to incorporate dGTP
opposite dFdCMP, is associated with the second worst efficiency ratio of 1,047 (Table
2.4). Our kinetic analysis also revealed that, in comparison to the extension of normal
25/45-mer, nucleotide incorporation efficiency of Polγ drops to between 5 and 132-fold
at one nucleotide preceding dFdCMP and at two to three nucleotides past the gemcitabine
lesion (Table 2.4). However, Figure 2.5 parts (B) and (D) did not show obvious pausing
by Polγ at these positions. This lack of pausing is due to the relatively high kp values (2-8
s-1) and long reaction times which allowed Polγ to rapidly elongate the corresponding
intermediate products. Taken together, one embedded gemcitabine in a template
unfavorably affected five nucleotide incorporations. This inhibitory effect can be
attributed to local DNA structural perturbations caused by dFdCMP [292] as described
above. The 3’-endo pucker conformation of template base dFdCMP likely causes
unproductive basepairing between an incoming dGTP and dFdCMP, leading to 183-fold
higher Kd for the binding of dGTP to Polγ 19/41F-mer (Table 2.4). Such a noncanonical
conformation of dFdCMP also decreased the binding affinity of the next two downstream
nucleotides by 6- to 10-fold before recovering to normal. The predicted negative impact
on the basepairing by the 3’-endo pucker is supported by the fact that the melting
temperature of the 12-basepair Okazaki fragment is lowered by 4.3 C in the presence of
an internal dFdCMP [292]. Such a noncanonical conformation of dFdCMP in a template
should also affect the positioning of an incoming dNTP for in-line attack by the primer
3’-OH moiety during phosphodiester bond formation, leading to a low kp. The negative
impact is the largest (225-fold) for dTTP incorporation into 20/41F-mer. Interestingly,
86
the kp for dGTP incorporation opposite dFdCMP is only lowered by 6-fold relative to its
incorporation opposite dCMP. Another interesting observation is that the inhibitory effect
of gemcitabine is larger as a template base than as an incoming nucleotide when we
compared the efficiency ratios in Tables 2.2 and 2.4. This indicates that the puckering
conversion is probably more difficult when gemcitabine is a template base than when it is
either an incoming nucleotide or a primer-terminal base. The reason for these interesting
observations is unclear at present.
Unfaithful Bypass of Template dFdCMP
When a template lesion causes a DNA polymerase to pause, this enzyme tends to
catalyze polymerization in an error prone manner. This general trend is also the case
when Polγ bypassed template dFdCMP. Table 2.5 shows that the polymerase activity of
Polγ has a fidelity of 396-3,071 at the two strong pause sites (Figure 2.5 parts (A) and
(D)), which is significantly smaller than the corresponding fidelity of 280,000-640,000
determined with normal 25/45-mer [291]. Thus, Polγ was 200-400 fold less faithful when
this enzyme paused. To compound these mutagenic events, the misincorporated
nucleotide, like 3’-dTMP in 20T/41F-mer, was excised more slowly than a single 3’mismatched primer base in normal DNA [289]. In addition, the matched primer 3’-dGMP
in 20/41F-mer was removed at half the speed of a normal 23/41-mer (Table 2.3). These
excision rate constants further suggest that the 3’-endo pucker conformation of template
base dFdCMP actually facilitates nucleotide misincorporations by slowing the 3’5’
exonuclease activity of Polγ. For editing, the currently established mechanism is that the
87
primer 3’-mismatched base is transferred from the polymerase active site to the 3’5’
exonuclease active site for excision while the template strand remains at the polymerase
active site of Polγ [142]. The template base dFdCMP may slow either the transfer
process, the excision step, or both. More studies in our laboratory are under way to
examine these possibilities. Taken together, a single dFdCMP template base reduces the
polymerization fidelity of Polγ by two orders of magnitude. Since mtDNA repair is
limited and inefficient [294], mitochondrial DNA replication is predicated to be error
prone if each template strand contains 188 dFdCMPs (see above estimation).
Pathologic Effects Associated with Gemcitabine Therapy
Our kinetic analysis directly confirms that gemcitabine, as both an incoming
nucleotide and as a template base, inhibits DNA synthesis catalyzed by human DNA
polymerase
holoenzyme. Moreover, our studies also showed that each template
dFdCMP is a mutagenic “hotspot” during replication. Additional mutagenic effects can
be exerted by gemcitabine metabolites in mitochondria. It is known that gemcitabine
metabolites cause cellular dNTP pool imbalances by inhibiting ribonucleotide reductase
[267, 268], CTP synthetase [269, 270], and dCMP deaminase [271]. This drug is also
likely to cause an imbalance of mitochondrial nucleotide pools. Such imbalances are
found to be mutagenic to the mitochondrial genome [295] and cause human diseases like
mitochondrial
neurogastrointestinal
encephalomyopathy
[296].
Considering
that
mammalian cells have 1,000-5,000 copies of the mitochondrial genome [297, 298] and
that mtDNA replication occurs continuously throughout the entire cell cycle [293, 299],
88
these inhibitory and mutagenic effects of gemcitabine should be further amplified in vivo.
The inhibitory effect of gemcitabine on mitochondrial genome replication either directly
leads to apoptosis or facilitates the killing of mammalian cells. The mutagenic effect of
embedded gemcitabine will result in genomic instability within mitochondria. In vivo, the
inhibitory and mutagenic effects of gemcitabine will synergistically cause mitochondrial
dysfunction and facilitate the killing of both cancer and normal cells, leading to clinical
efficacy and toxicity. For example, one toxic side effect observed in gemcitabine therapy
is peripheral neuropathy [274, 275]. It is known that mitochondria play a central role in
the propagation of neuronal cell death [300], and mitochondrial dysfunction is beginning
to emerge as a commonality linking many of the most prevalent neurodegenerative
disorders [301]. Interestingly, peripheral neuropathy has been associated with many
antiviral nucleoside analog drugs and has been shown to directly correlate to the
inhibitory effects of these drugs on human Polγ [302]. Based on our evidence in this
paper, it is reasonable to assume that gemcitabine may provoke neurological
complications by a similar mechanism.
Another side effect resulting from gemcitabine therapy is severe pulmonary
toxicity, including acute respiratory distress syndromes. Although ARDS can be triggered
in many ways, a common feature among ARDS cases is dysfunction of the pulmonary
surfactant, a unique lipid-protein mixture which lines the alveoli and is essential for
proper lung function [303, 304]. Although the exact mechanism by which the pulmonary
surfactant is generated is the subject of considerable debate, there is evidence that the
89
mitochondria may play a pivotal role. As early as 1962, it was observed that alveolar
mitochondria appear to undergo a transformation which coincides with the first
production of alveolar surfactant in the fetal murine lung and that this transformation may
result in the production of the pulmonary surfactant or its precursor [305]. Since then, it
has been hypothesized that formation of surfactant molecules may involve the
mitochondria in alveolar epithelial cells which secrete the pulmonary surfactant [306,
307]. Clearly, our data demonstrate that gemcitabine metabolites inhibit human Polγ and
most likely interfere with mitochondrial functions. However, whether gemcitabine
associated mitochondrial dysfunction contributes to pulmonary toxicity remains an
intriguing
possibility
that
warrants
90
further
investigation.
2.6. Figures
Figure 2.1 Chemical Structure of Gemcitabine and 2'-Deoxycytidine.
91
Figure 2.2 Gemcitabine Activation and Self Potentiation Pathways. Dashed lines lead to
enzyme inhibition symbolized by an “X”. Gemcitabine is denoted as dFdC.
92
Figure 2.3 Pre-steady state kinetic analysis of Polγ.
(A) An autoradiograph gel image shows the incorporation of dFdCTP (16 μM) catalyzed
by a Polγ mutant E200A. (B) DNA Incorporation of dFdCTP by Polγ. A preincubated
solution of E200A (90 nM), Polγ accessory subunit (450 nM), and 5'-[32P]-labeled
22/41-mer (30 nM) was rapidly mixed with increasing concentrations of dFdCTP•Mg2+
(2.4 μM, ; 4.9 μM, ; 9.7 μM, ; 19.5 μM, ; 39 μM, ; 78 μM, ) and reacted at
37 ˚C for increasing times. The solid lines were fit to Eq. 1 using non-linear regression
which determined the observed rate constants, kobs. (C) Pre-steady state kinetic analysis
of DNA Incorporation of dFdCTP by Polγ. The kobs values were plotted as a function of
dFdCTP concentration. The data () were fit to Eq. 2 using non-linear regression, thus
yielding a kp of 2.0 ± 0.3 s-1 and a Kd of 21 ± 7 μM.
93
Figure 2.3
94
Figure 2.4 Measurement of the Rate Constant of DNA Primer Degradation by the 3' 5'
Exonuclease Proofreading Activity of the Wild-type Polγ. A preincubated solution of the
wild-type Polγ (100 nM), Polγ accessory subunit (500 nM) and 23/41-mer or 23F/41-mer
(75 nM) in buffer G was rapidly mixed with 2.5 mM MgCl2 and reacted for various time
intervals. The excision reaction was quenched by the addition of 0.37 M EDTA. The
concentration of the remaining full length primer versus time was plotted and fit to Eq. 3
(23/41-mer, ; 23F/41-mer, ) yielding a kexo of 0.06 ± 0.02 s-1 for 23/41-mer and
0.0011 ± 0.0001 s-1 for 23F/41-mer.
95
Figure 2.5 Running Start Primer Elongation Catalyzed by the Wild-Type Polγ and the
Exonuclease Deficient Mutant E200A. A solution of control substrate 15/41-mer (Figures
2.5.3.A and 2.5.3.C, 30 nM each) or DNA substrate 15/41F-mer (Figures 3B and 3D, 30
nM each), preincubated with the wild-type Polγ (Figure 3A and 3B, 90 nM) or E200A
(Figure 2.5.3.C and 2.5.3.D, 90 nM) and Polγ accessory subunit (450 nM) in buffer G
was rapidly mixed with 2.5 mM MgCl2 and 100 μM each of dATP, dCTP, dGTP and
dTTP. Reactions were allowed to continue for various time intervals before being
quenched by the addition of 0.37 M EDTA. Product lengths and the position of the
embedded dFdC moiety in the template are indicated.
96
Figure 2.5
97
A
(continued)
Figure 2.6 Sequencing gel image of single nucleotide incorporation catalyzed by Polγ
mutant E200A . A preincubated solution of 30 nM DNA substrate (19/41F-mer (A) or
20/41F-mer (B)), 90 nM E200A, and 450 nM Polγ accessory subunit in buffer G was
reacted with a single dNTP (100 μM) for indicated time intervals before being quenched
by 0.37 M EDTA.
98
Figure 2.6 (continued)
B
99
2.7. Tables
15/41-mer
5’-GGACGGCATTGGATC
3’-CCTGCCGTAACCTAGCTGCCACTCAACCAACCTGCCGACGC-5’
22/41-mer
5’-CGCAGCCGTCCAACCAACTCAC
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
23/41-mer
5’-CGCAGCCGTCCAACCAACTCACC
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
23F/41-merb
5’-CGCAGCCGTCCAACCAACTCACF
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
24FG/41merb
5’-CGCAGCCGTCCAACCAACTCACFG
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
25FGT/41merb
5’-CGCAGCCGTCCAACCAACTCACFGT
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
15/41F-merb
5’-GGACGGCATTGGATC
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
18/41F-merb
5’-GGACGGCATTGGATCGAC
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
19/41F-merb
5’-GGACGGCATTGGATCGACG
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
20/41F-merb
5’-GGACGGCATTGGATCGACGG
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
20T/41Fmerb
5’-GGACGGCATTGGATCGACGT
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
Table 2.1 DNA Substrates. aThe downstream strand 19-mer of substrates 21-19/35-mer
and 21F-19/35-mer were 5'-phosphorylated. b“F” denotes gemcitabine. cThe template
base opposite primer 26th base varied to allow correct base pairing of incoming dNTP.
(continued)
100
Table 2.1 (continued)
21/41F-merb
5’-GGACGGCATTGGATCGACGGT
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
22/41F-merb
5’-GGACGGCATTGGATCGACGGTG
3’-CCTGCCGTAACCTAGCTGCFACTCAACCAACCTGCCGACGC-5’
25/45-merc
5’-GCCTCGCAGCCGTCCAACCAACTCA
3’-CGGAGCGTCGGCAGGTTGGTTGAGTTGGAGCTAGGTTACGGCAGG-5’
21-19/35mera
5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTAC-5’
21F-19/35mera
5’-CGCAGCCGTCCAACCAACTCAF CGTCGATCCAATGCCGTCC-3’
3’-GCGTCGGCAGGTTGGTTGAGTG-GCAGCTAGGTTAC-5’
101
DNA
dNTP
22/41-mer
dCTP
22/41-mer
dFdCTP
23F/41-mer
dGTP
24FG/41-mer dTTP
25/45-mer FIAUTPc
Kd
(μM)
kp
(s-1)
kp/Kd
(μM-1s-1)
0.9 ± 0.2
37 ± 2 41
21 ± 7 2.0 ± 0.3 0.095
7.2 ± 0.9 1.5 ± 0.1 0.21
0.5 ± 0.1 2.8 ± 0.1 5.6
2.9 ± 0.7c 24 ± 2c
8.3c
Efficiency
Ratioa
1
432
214b
7b
5
Table 2.2 Kinetic Parameters of Single Nucleotide Incorporation Catalyzed by Polγ
E200A under Single-Turnover Conditions at 37 C. aCalculated as (kp/Kd)correct dNTP into
normal DNA/(kp/Kd)correct dNTP into DNA containing dFdCMP.
reference [289]. cFrom reference [289].
102
b
(kp/Kd)correct dNTP into normal DNA is from
DNA
Fast Phase
kexo, 1 (s-1)
23/41-mer
0.06 ± 0.02
23F/41-mer
0.0011 ± 0.0001
20/41F-mer
0.028 ± 0.006
20T/41F-mer
0.2 ± 0.2
Fast Phase
Amplitude (%)
Slow Phase
kexo, 2 (s-1)
Slow Phase
Amplitude (%)
NA
NA
NA
17 ± 8.0
NA
NA
NA
0.008 ± 0.002
NA
NA
NA
83 ± 8.0
Table 2.3 Excision Rate Constants for the 3' 5' Exonuclease Activity of the Wild-Type
Human Polγ Holoenzyme under Single-Turnover Conditions at 37 C.
103
DNA
dNTP
Kd
(μM)
kp
(s-1)
kp/Kd
Efficiency
a
-1 -1
(μM s ) Ratio
18/41F-mer
19/41F-mer
20/41F-mer
21/41F-mer
22/41F-mer
dGTP
dGTP
dTTP
dGTP
dATP
1.1 ± 0.2
150 ± 10
6.0 ± 0.5
5.0 ± 2
0.7 ± 0.1
2.1 ± 0.1
6.3 ± 0.2
0.11 ± 0.002
1.7 ± 0.4
8.3 ± 0.3
1.9
0.042
0.018
0.34
12
25/45-merb
25/45-merb
25/45-merb
25/45-merb
dATP
dTTP
dGTP
dCTP
0.8 ± 0.1
0.6 ± 0.2
0.8 ± 0.1
0.9 ± 0.2
45 ± 1
25 ± 2
37 ± 2
43 ± 2
57
39
45
47
2.4 x 101
1.1 x 103
2.2 x 103
1.3 x 102
4.8
Table 2.4 Kinetic Parameters of Single Nucleotide Incorporation into DNA Containing a
Template Base dFdCMP Catalyzed by Polγ E200A under Single-Turnover Conditions at
37 C. aCalculated as (kp/Kd)correct dNTP into 25/45-mer/(kp/Kd)dNTP into DNA containing a template base
dFdCMP.
b
Kinetic parameters are from reference [289].
104
DNA
dNTP
Kd
(μM)
kp
(s-1)
19/41F-mer
19/41F-mer
20/41F-mer
20/41F-mer
dGTP
dTTPb
dTTP
dATPb
150 ± 10
110 ± 30
6.0 ± 0.5
120 ± 20
6.3 ± 0.2
0.0015 ± 0.00015
0.11 ± 0.002
0.0059 ± 0.0003
kp/Kd
(μM-1s-1)
4.2 x 10-2
1
-5
1.4 x 10 3.0 x 103
1.8 x 10-2
1
-5
4.9 x 10 3.7 x 102
Table 2.5 Fidelity at the Two Strong Pause Sites. aCalculated as (kp/Kd)correct
dNTP/(kp/Kd)incorrect dNTP.
b
Bold-type indicates incorrect nucleotide.
105
Fidelitya
Chapter 3: Probing protein conformational changes of a human DNA polymerase
using mass spectrometry
3.1. Introduction
Based upon phylogenetic analyses, DNA polymerases have been arranged into six
families which are designated as A, B, C, D, X and Y. Members of the novel X-family of
DNA polymerases belong to a larger superfamily of nucleotidyl transferases and are
found in all three domains of life [6, 308]. Thus far, at least five X-family members have
been identified in humans: DNA polymerases lambda (Polλ), beta (Polβ), Mu, sigma, and
terminal deoxynucleotidyl transferase (TdT). Polβ is known to function in base excision
repair (BER) in vivo [309, 310]. Physiologically, Polλ has been proposed to function in
BER, non-homologous end joining, and V(D)J recombination [55, 56, 58-60, 311-313].
Polβ and Polλ share 54% sequence homology and 32% sequence identity [37]. In
addition, both possess gap-filling DNA polymerase activity at their C-terminal
polymerase domain, 5′-deoxyribose-5-phosphate lyase (dRPase) activity at their 8-kD
dRPase domain, and lack 3′5′ exonuclease activity (Figure 3.1) [35, 36, 55]. However,
the full-length Polλ (fPolλ) possesses a breast cancer susceptibility gene 1 C-terminal
(BRCT) domain, a Proline-rich domain, and a nuclear localization signal motif on its Nterminus which are absent in the full-length Polβ (Figure 3.1). Interestingly, the Proline106
rich domain has been shown to functionally suppress the polymerase activity of fPolλ
[78] and to increase its fidelity by up to 100-fold [288].
In general, DNA polymerases are structurally and functionally quite diverse,
although, commonalities can be found. Firstly, crystal structures for all known templatedependent DNA polymerases reveal that they share a common three-dimensional shape
which resembles a human right hand. This feature of E. coli DNA polymerase I [72] led
to the “fingers, palm, and thumb” domain nomenclature system. Secondly, all DNA
polymerases perform the chemistry of nucleotidyl transfer using the same two divalent
metal ion mechanism as first proposed by T.A. Steitz [122]. This mechanism, believed to
be one of the earliest enzymatic activities to evolve [314], involves the coordination of
two divalent metal ions with three conserved carboxylate residues within a polymerase
active site, the primer 3′-OH moiety, and the triphosphate moiety of an incoming
nucleotide (dNTP). Finally, a dramatic protein conformational change is observed in the
conversion from the binary complex of enzyme•DNA to the ternary complex of
enzyme•DNA•dNTP. During this change, the fingers domain moves toward the palm
domain forming the hydrophobic polymerase active site. This swing of the fingers
domain changes the polymerase from the catalytically inactive “open” to the active
“closed” conformation [5]. To catalyze nucleotidyl transfer, formation of the closed
conformation aligns the polymerase active site by properly orienting the conserved
catalytic carboxylates, divalent metal ions, DNA, and dNTP [315].
107
Recent crystallographic studies with human truncated Polλ (tPolλ, Figure 3.1)
revealed that unlike other DNA polymerases [81], including Polβ [121, 316] for which
atomic structures are available, the catalytic cycle of Polλ might not involve a large,
protein domain rearrangement. Rather, when comparing three crystal structures of human
tPolλ
(tPolλ•gapped
DNA,
tPolλ•gapped
DNA•dNTP,
and
tPolλ•nicked
DNA•pyrophosphate), dNTP binding induces a repositioning of only four side chains (i.e.
Y505, F506, R514, and R517) within the active site and a minor shift in the position of
two β-strands [81]. Together, these movements shift the DNA template strand. However,
the crystallographic studies that demonstrate this unique mechanism were performed with
tPolλ, rather than fPolλ. The Proline-rich domain absent in tPolλ has recently been shown
to increase the fidelity of Polλ by up to 100-fold, although, the reason for this is not yet
known [288]. It is conceivable that upon dNTP binding, the Proline-rich domain induces
either the large swing of the finger domain [317] or relatively modest active site
rearrangements in the solution phase. Either of these protein conformational transitions
may provide a thermodynamic basis for selecting matched over mismatched incoming
dNTPs [318]. In the present study, we investigated the extent of protein conformational
changes in the solution phase using mass spectrometry (MS)-based protein footprinting
methods.
108
3.2. Materials
Preparation of Human fPolλ, dPolλ, and tPolλ
Cloning, expression, and purification of human fPolλ [288], dPolλ [288], and
tPolλ [319] were described previously.
Synthetic Oligodeoxyribonucleotides
The oligodeoxyribonucleotides in Table 3.1 were purchased from Integrated DNA
Technologies (Coralville, IA) and purified by denaturing polyacrylamide gel
electrophoresis (17% acrylamide, 8 M urea, Tris-borate-EDTA running buffer). Their
concentrations were determined by UV absorbance at 260 nm with calculated extinction
coefficients. Each single-nucleotide gapped DNA substrate was prepared by heating a
mixture of 21-mer (or 22-mer), 19-mer (or 18-mer), and 41-mer in a 1:1.25:1.15 molar
ratio, respectively, for 8 min at 95 °C and then cooling the mixture slowly to room
temperature over 3 h as described previously [319]. For polymerization assays, a DNA
primer was 5′-[32P]-labeled by incubating [γ-32P]ATP (GE Healthcare) and T4
polynucleotide kinase (New England BioLabs, Inc) for 1 hour at 37 °C. The unreacted [γ32
P]ATP was subsequently removed by centrifugation via a Bio-Spin-6 column (Bio-Rad
Laboratories). Lastly, the radiolabeled primer was annealed to form the appropriate
gapped DNA substrate as described above.
109
Reaction Buffer
Reaction buffer L contained 50 mM Tris-Cl (pH 8.4 at 37 °C), 5 mM MgCl2, 100
mM NaCl, and 0.1mM EDTA and 10% glycerol. For the kinetic assays, reaction buffer L
was supplemented with 0.1 mg/mL BSA and 5 mM DTT. This reaction buffer was
optimized previously for transient state kinetic analysis of fPolλ and its deletion
constructs [288, 319]. All reactions reported herein were carried out in the appropriate
reaction buffer at 37 °C, and all concentrations refer to the final concentration of the
components after mixing.
110
3.3. Methods
Mass Spectrometry-Based Protein Footprinting Assay
In parallel experiments using fPolλ, dPolλ, and tPolλ: enzyme (10 µM), the
enzyme (10 μM)•22-18/41G-mer (60 μM) binary complex, and the enzyme (10
μM)•22ddC-18/41G-mer (60 μM)•dCTP (100 μM) ternary complex were subjected to
chemical modification by HPG. HPG reacts specifically with the guanidine group in an
arginine residue resulting in 132 Da mass increase [320, 321]. Previous optimization
experiments indicated the Arg/HPG ratio in the range of 1:40 to 1:20 was optimal for
achieving very mild modification conditions, under which the integrity of the functional
complexes was preserved[322, 323]. These conditions were adopted for footprinting
purified free Polλ and its binary and ternary complexes. The HPG treatments were carried
out at 37 ˚C in the dark for 60 minutes and terminated by addition of 160 mM final
concentration of arginine in its free form. Pol was then separated from DNA and dNTP
by SDS-PAGE. The protein bands were excised, destained, dehydrated, and digested with
1 g of trypsin in 50 mM NH4HCO3 at 25 C overnight.
Small molecular weight peptides were analyzed by MALDI-ToF MS using
AXIMA-CFR instrument (Shimadzu Scientific Instruments). The samples were analyzed
with an α-cyano-4-hydroxycinnamic acid matrix as described previously [324]. Sequence
data and Protein Prospector v4.0.6 (http://prospector.ucsf.edu) were used to identify Pol
peptide peaks. Modified arginine residues were assigned by identifying mass peaks that
111
appear only in the spectra of HPG-modified Pol
and that have a molecular weight
corresponding to the sum of the predicted peptide fragment plus the 132 Da HPG adduct.
For accurate quantitative analysis of the modified peptide peaks, at least two unmodified
proteolytic peptide peaks were used as internal controls. A protection was considered to
be significant when the intensity of the given modified peptide peak derived from HPG
treated free protein was reduced at least 10-fold in the context of the nucleoprotein
complexes. A modified peptide peak was considered unprotected when the intensities of
the given peptide obtained from free protein and nucleoprotein complexes were within ±
20% of each other. The data were reproducibly compiled and analyzed from at least three
independent experimental groups.
Gap-Filling DNA Polymerase Activity Assay for HPG-Modified Enzymes
Following chemical modification with 10 mM HPG at 37 ˚C for 60 minutes (see
above) and quenching of the reaction with arginine (160 mM), the gap-filling DNA
polymerase activity of HPG-modified Polλ was tested by pre-incubating the modified
enzyme (10 μM) with 5′-[32P]-labeled 22-18/41G-mer (60 nM) for 5 min at 37 ˚C. The
polymerization reaction was then initiated by the addition of dCTP (100 μM), and the
reaction was terminated with EDTA (0.37 M) after 1 minute. Reaction products were
separated using denaturing polyacrylamide gel electrophoresis and visualized using
autoradiography.
112
Determination of kp and Kd Values
Both the maximum rate of nucleotide incorporation kp and the equilibrium
dissociation constant Kd of an incoming nucleotide were determined under singleturnover conditions: a pre-incubated solution of Polλ (300 nM) and 5′-[32P]-labeled 2119/41A-mer (30 nM) was mixed with increasing concentrations of dTTP (GE Healthcare)
in reaction buffer L at 37 °C. Reactions were terminated by adding 0.37 M EDTA at
various times. For rapid nucleotide incorporations, experiments were performed using a
rapid chemical-quench flow apparatus (KinTek). Reaction aliquots, sampled at discrete
time points, were analyzed by sequencing gel analysis and quantitated using a Typhoon
TRIO PhosphorImager (GE Healthcare). Each time course of product formation was fit to
a single-exponential equation (Equation 1) to yield an observed rate constant of
nucleotide incorporation kobs and reaction amplitude (A). The kobs values were
subsequently plotted against the corresponding dTTP concentration, and these data were
fit to a hyperbolic equation (Equation 2) using non-linear regression to yield the kp and Kd
values.
[Product] = A[1 - exp(-kobst)]
Eq. 1
kobs = kp[dNTP]/{[dNTP] + Kd}
Eq. 2
113
3.4. Results
In order to investigate whether or not human fPolλ undergoes a large protein
conformational change during catalysis in the solution phase, we employed a novel mass
spectrometry-based protein footprinting method [323-325]. Briefly, a gentle, covalent
modification
of
a
protein
at
solvent
accessible
arginine
residues
by
p-
hydroxyphenylglyoxal (HPG) is conducted under near physiological conditions. After
modification, the protein is purified and digested by trypsin, a serine protease, which
predominantly cleaves peptide chains at the carboxyl side of the amino acids lysine and
arginine, except when either residue is followed by proline or is chemically modified.
Thus, trypsin cannot cleave after HPG-modified Arg residues in peptide chains.
Following the trypsin digestion, the pool of low molecular weight peptides is analyzed by
Matrix Assisted Laser Desorption Ionization Time of Flight (MALDI-ToF) instrument to
obtain MS data. Notably, each HPG modification on the side chain of an Arg residue
leads to the addition of 131 Daltons to the molecular weight of a peptide. Analysis of a
peptide MS spectrum will reveal the precise location of the chemical modifications and
the extent to which they are modified (Section 3.3). The solvent accessibility of Arg
residues is sensitive to protein conformation. Therefore, comparative analysis of
modification patterns in free protein versus its complexes with cognate nucleic acids
could reveal protein conformational change(s). A side chain, which becomes less
amenable to modification (i.e. a decrease in peak intensity within the MS spectrum
compared to control peaks) is deemed “protected” and is less solvent accessible.
114
Conversely, if chemical modification becomes more facile (i.e. an increase in peak
intensity within the MS spectrum compared to control peaks), the side chain is said to be
“hyper-reactive” and more solvent accessible [323, 325]. Here, this MS foot-printing
method was employed to analyze the nature of conformational changes that may occur in
fPolλ through a single turnover of the catalytic cycle. In addition, the above set of
conditions was repeated for dPolλ and tPolλ (Figure 3.1) to assess any inherent
differences between the solution-phase structures of these proteins in the presence or
absence of DNA and dNTP.
Investigating the stability of human Polλ during HPG modification
For meaningful protein footprinting experiments it is essential to establish mild
modification conditions under which the integrity of the functional nucleoprotein
complexes is preserved. Previous studies indicated that the treatment with 10 mM HPG to
be optimal [323, 325]. To make sure that these conditions were applicable to fPolλ, we
conducted kinetic analysis of the gap-filling DNA polymerase activities following HPG
treatments (Figure 3.2). The solutions of apo-fPolλ and the binary complex of fPolλ and
22-18/41G-mer (Table 3.1) were first reacted with 10 mM HPG and then quenched by a
molar excess of arginine. The modified fPolλ in these two reaction solutions (lanes
“HPG-E” and “HPG-ED” in Figure 3.2) and unmodified fPolλ (lane “+ Control”) were
examined for their ability to incorporate correct dCTP into single-nucleotide gapped 2218/41G-mer (Section 3.3). Similar intensities of both product 23-mer and remaining
primer 22-mer on the three right lanes in Figure 3.2 suggested that the gap-filling DNA
115
polymerase activity of fPolλ, in both the apo form and the binary complex with DNA,
was almost preserved following HPG modification. Thus, HPG modification conditions
were sufficiently mild so that the solution-phase structure of fPolλ remained intact for
both its apo form and binary complex with DNA. We did not examine if the HPG
modification step altered the integrity of the ternary complex of fPolλ, 22ddC-18/41Gmer (Table 3.1), and dCTP due to the difficulty to completely replace the fPolλ-bound,
dideoxy-terminated DNA substrate with 22-18/41G-mer in the gap-filling DNA
polymerase activity assay (Section 3.3). However, such an alteration was unlikely to
occur considering that the binary complex was not affected by HPG modification. Similar
gap-filling DNA polymerase activity assays were performed with dPolλ and tPolλ and
demonstrated that HPG modification did not disturb the conformations of these truncated
mutants of fPolλ (data not shown).
MS-Based Footprinting of fPolλ, dPolλ, and tPolλ
Representative MS results comparing surface accessible Arg residues of tPolλ in
its apo form with the binary and ternary complexes are depicted in Figure 3.3. Figure 3.3
part (D) is a portion of the mass spectrum for unmodified tPolλ and shows a peak
corresponding to tryptic peptide 524-538 which is present in all spectra at the same
intensity and is used as an internal control. Figure 3.3 parts (A), (B), and (C) show the
same portion of the MS spectrum of HPG-modified apo-tPolλ, the binary complex of
tPolλ•22-18/41G-mer, and the ternary complex of tPolλ•22ddC-18/41G-mer•dCTP,
respectively. In the ternary complex, dCTP was not incorporated because 22ddC-18/41G116
mer contains a dideoxy-terminated primer. In Figure 3.3 parts (A), (B) and (C), peaks
resulting from tryptic peptides 562-573, 313-324, and 379-389 correspond to HPG
modification of R568, R323, and R386, respectively, thereby shifting their m/z ratios.
The spectra in Figure 3.3 parts (A) and (B) are almost superimposable, suggesting that
the structure of tPolλ is relatively unchanged from apo-tPolλ to the binary complex
tPolλ•22-18/41G-mer. In contrast, the intensity of the modified 379-389 (R386+HPG)
peak was significantly diminished in the tPolλ•22ddC-18/41G-mer•dCTP complex
(Figure 3.3 part (C)) indicating that R386 was shielded by the bound nucleotide.
The purified, recombinant fPolλ, dPolλ, and tPolλ possess 41, 28, and 25 Arg
residues, respectively (Figure 3.1). In the absence of DNA and dNTP, HPG modified 13,
10, and 10 Arg residues of the apo forms of fPolλ, dPolλ, and tPolλ, respectively (Table
3.2). The HPG-modified Arg residues of apo-dPolλ and apo-tPolλ are identical,
indicating that the Proline-rich domain did not shield any of the exposed Arg residues in
the Polβ–like domain (Figure 3.1). At the same time it is important to note that not all the
surface exposed Arg residues could be detected by our approach as certain large tryptic
peptide fragments may not be readily amenable for MALDI-ToF analysis. For example,
the single Arg residue (R174) in the Proline-rich domain (Figure 3.1) is followed
immediately by a Pro residue thereby making trypsin cleavage at R174 impossible
regardless of HPG modification. Therefore, the shortest peptide resulting from trypsin
digestion to contain R174 is 147-181, which likely escaped detection due to its large size.
Thus, modification at R174 is beyond the limit of detection for this assay and may or may
117
not be modified. Finally, R50, R55, and R57 which are located in the BRCT domain
(Figure 3.1), were modified by HPG in apo-fPolλ, but not in apo-dPolλ or apo-tPolλ,
because these two Polλ fragments do not contain the BRCT domain.
In the presence of 22-18/41G-mer (60 M, Table 3.1), the HPG-modified Arg
residues in fPolλ, dPolλ, and tPolλ were identical to those detected in the corresponding
apo forms of these Polλ constructs (Table 3.2). This suggested that the solution phase
structures of fPolλ, dPolλ, and tPolλ were not significantly altered in the presence of
DNA. In contrast, upon formation of the ternary complex of Polλ•gapped DNA•dNTP,
two residues R275 and R386 were shielded from HPG modification in all three Polλ
constructs (Table 3.2). Interestingly, the X-ray crystal structure of the ternary complex
tPolλ•gapped DNA•ddTTP shows that R386 forms a salt bridge with the -phosphate
moiety of the incoming ddTTP (Figure 3.5). Thus, the binding of a dNTP likely shielded
R386 from HPG modification. In addition, the X-ray crystal structure [81] reveals that
R275 forms a salt bridge with the 5’ terminal phosphate moiety of the downstream DNA
primer (Figure 3.5 part (C)).
Pre-steady state kinetic analysis of two R386 mutants of fPolλ
To investigate the significance of the salt bridge formed between R386 and a
dNTP, we created two point mutants of fPolλ, R386A and R386E. The purpose of the
alanine substitution was to generate a small, neutral side chain at residue 386, while the
glutamate substitution was to create a nearly isosteric side chain with a negative charge
118
which could repel a dNTP through charge-charge interactions. To kinetically characterize
these two mutants, we determined their gap-filling DNA polymerase activity separately
under single-turnover reaction conditions (Section 3.3). For example, a pre-incubated
solution of R386A (300 nM) and 5′-[32P]-labeled 21-19/41A-mer (30 nM, Table 3.1) was
mixed with increasing concentrations of dTTP (4-64 µM) for varying times before being
quenched by 0.37 M EDTA. Each time course of product formation was fit to Equation 1
(Section 3.3) to yield an observed rate constant (kobs) (data not shown). The kobs values
were then plotted against the corresponding dTTP concentration (Figure 3.6), and the
data were fit to Equation 2 (Section 3.3) to determine a maximum rate of nucleotide
incorporation (kp) of 1.3 ± 0.1 s-1 and an equilibrium dissociation constant (Kd) of 15 ± 5
μM (Table 3.3). Under the same reaction conditions, R386E catalyzed the incorporation
of dTTP into 21-19/41A-mer with a kp of 0.005 ± 0.001 s-1 and a Kd of 1000 ± 410 μM
(Table 3.3 and Figure 3.6 (B)). The substrate specificity, kp/Kd, was calculated to be 0.087
and 5x10-6 μM-1s-1 for R386A and R386E, respectively (Table 3.3). These values are
much lower than 1.5 μM-1s-1 observed with the wild-type fPolλ [288]. Thus, both the
charge and size of the side chain of R386 are important to the catalytic activity of fPolλ.
119
3.5. Discussion
Structural Implications of Our MS-Based Protein Footprinting Data
For the MS-based footprinting method, a small molecule like HPG is a proven
chemical to readily modify the most solvent accessible Arg residues of a protein in the
solution phase [324, 326-328]. The reagent modified 13, 10, and 10 Arg residues in the
apo forms of fPolλ, dPolλ, and tPolλ, respectively (Table 3.2). The affected amino acids,
projected onto the crystal structure of tPolλ, are depicted in Figure 3.7. Of these, only two
residues R275 and R386 were selectively protected in the ternary complex
(Polλ DNA dNTP), not in the binary complex (Polλ DNA) and apo-Polλ.
The shielding of R275 in the ternary complex is consistent with the X-ray crystal
structure of tPolλ gapped DNA dNTP [81] implicating this residue in charge-charge
interactions with the 5′-phosphate of the downstream strand. Since R275 was modified in
the tPolλ gapped DNA complex, it is logical to suggest that these interactions in the
context of the binary complex are less stable or highly transient. In contrast, in the
presence of an incoming dNTP, R275 was protected from modification. This may have
been due to dNTP-induced stabilization of the ternary complex. Such stabilization may
have arisen by equalizing unsatisfied positive charges (for example R386, R420) in the
polymerase active site. Thus, by introducing the dNTP’s negatively charged triphosphate
moiety, positively charged surfaces on the interior of the protein may have experienced
less charge-charge repulsion and thus settled into greater proximity. In addition, a
120
resulting reduction of protein dynamics may have stabilized R275 by immobilizing the
thumb domain in the tPolλ DNA dNTP structure, thus leading to greater interaction of
R275 with 5’-phosphate of the downstream strand.
MS-based protein footprinting also revealed selective protection of R386 in the
ternary complex (Polλ DNA dNTP) and not in the binary complex (Polλ DNA) and
apo-Polλ. These results are in excellent agreement with the X-ray crystal structure of the
ternary complex tPolλ•gapped DNA•ddTTP showing that R386 forms a salt bridge with
the -phosphate moiety of the incoming ddTTP (Figure 3.5 part (B)). The fact that we did
not observe additional protections or hyper-reactive Arg residues in the ternary
complexes suggests that dNTP binding does not induce significant protein
conformational changes. Consistently, crystallographic studies of tPolλ [81] have
indicated that dNTP binding induces only a repositioning of four active site side chains
and a minor shift in the position of two β-strands. Taken together, our MS-based
footprinting data indicate that the solution-phase and solid-phase structures of tPolλ were
similar but not identical, especially at the local structure surrounding R275.
Comparing the MS-based footprinting spectra of fPolλ, dPolλ and tPolλ (Table
3.2) in the same substrate binding states reveals remarkable similarity, with the exception
of R50, R55, and R57 which reside in the BRCT domain and were expected not to be
probed by HPG in dPolλ and tPolλ. Given that nearly all probed Arg residues were
modified to similar extents, and that fPolλ, dPolλ, and tPolλ were subjected to conditions
121
designed to mimic discrete states of the catalytic cycle, it is reasonable to assume that this
polymerase does not undergo a radical conformational change involving Arg residues as
it carries out gap-filling DNA synthesis. However, these findings do not exclude the
possibility of small, local structural rearrangements within the polymerase active site as
catalysis occurs.
Structural and Functional Roles of R386
Table 3.2 illustrates that nucleotide binding to the binary complexes of all three
Polλ constructs protected R386 from chemical modification. In the crystal structure of
tPolλ gapped DNA ddTTP (Figure 3.5), a strong salt bridge (2.90 and 3.04 Å) likely
forms with the –phosphate of the bound ddTTP due to charge-charge attraction. Thus, it
is most likely the salt bridge that prevents the guanidinium moiety of R386 to react with
HPG. Such a salt bridge perhaps strengthens the ground-state binding affinity of an
incoming nucleotide, positions it for catalysis, and stabilizes pyrophosphate, the leaving
group. This possibility was strongly supported by the kinetic data of R386A and R386E,
two point mutants of fPolλ (Table 3.3). Relative to wild-type fPolλ, R386A catalyzed
correct dTTP incorporation into single-nucleotide gapped DNA with a 3-fold lower kp, 6fold higher Kd, and 17-fold lower incorporation efficiency (kp/Kd). These kinetic effects
from the side chain of R386 could be contributed to the positive charge, size, or a
combination of both properties. When R386 was mutated to a glutamic acid residue
which is of similar size but opposite charge, the kp was reduced by 780-fold, the Kd was
increased by 385-fold, and the nucleotide incorporation efficiency was decreased by
122
300,000-fold. Thus, the positive charge of R386 has a more important role in nucleotide
binding and catalysis than the size of its side chain. The R386E mutation could repel the
negatively-charged dNTP and significantly weaken its binding. Since the rate-limiting
step of the kinetic mechanism for nucleotide incorporation catalyzed by Polλ has not
been established, the charge-charge repulsion could either destabilize the transition state
or alter the positioning of dNTP during catalysis. Interestingly, in the crystal structure of
the product ternary complex tPolλ•nicked DNA•pyrophosphate [81], R386, R420, and
Mg2+ stabilize the negative charges on the pyrophosphate leaving group and facilitate
catalysis (Figure 3.5 part (C)). The R386E substitution is likely to inhibit the formation of
the pyrophosphate product through charge-charge repulsion during catalysis. Thus, the
R386E mutation could compromise coordination of both dNTP and pyrophosphate in the
enzyme active site, leading to the greatly reduced kp.
Conservation of R386 and R420 in Other DNA Polymerases
The structural and functional importance of R386 and R420 suggested that other
DNA polymerases may use similar positively-charged residues, arginine or lysine, to
anchor both the dNTP and pyrophosphate. Thus, we analyzed the available X-ray crystal
structures of the other ternary complexes in order to determine whether this feature was
conserved among DNA polymerases (Table 3.4). The X-ray crystal structure of
Polβ gapped DNA ddCTP shows that R149 and R183 are oriented near the triphosphate
group of ddCTP in a manner similar to Polλ’s R386 and R420 [121]. For DNA
polymerase Mu and TdT, these two members of the X-family possess the structural
123
homolog of R420 which corresponds to R323 for DNA polymerase Mu and R336 for
TdT [329, 330]. However, in lieu of a positively-charged residue structurally homologous
to R386 and/or R420, the negative charge on the dNTP phosphate can be stabilized by
utilizing lysine, histidine, asparagine, or possibly the amide nitrogen of the peptide
backbone (Table 3.4) [329, 330]. As an example, DNA polymerase IV from Sulfolobus
solfataricus employs a network of residues, an arginine (R51), a lysine (K159), a
phenylalanine (F11), and two tyrosines (Y10 and Y48), to cooperatively stabilize the
negatively-charged phosphates through salt bridge formation and hydrogen-bonding
interactions [331, 332].
After examining the published ternary structures of several DNA polymerases
listed in Table 3.4, we concluded that the presence of positively-charged residues, either
arginine or lysine, at the dNTP and pyrophosphate binding sites of a DNA polymerase
active site is almost universal. The alignment of the amino acid sequences of several
DNA polymerases, including the A-, B-, X-, and Y-family members for which crystal
structures are not available, is given in Figure 3.9. Although DNA polymerases are
structurally and functionally quite diverse, all DNA polymerases analyzed in this work
showed that at least one-positively charged residue was conserved in each DNA
polymerase family (Figure 3.9). In addition to the two metal ion mechanism proposed by
Steitz [122], our finding reveals another conserved feature among the DNA polymerases
examined herein. However, based upon our mutagenesis results, these positively-charged
124
residues are not an absolute requisite for catalysis, since the catalytic activity of Polλ
R386A remained fairly robust (Table 3.3).
In summary, our spectrometry-based protein footprinting method confirmed that
tPolλ, dPolλ, and fPolλ do not undergo a dramatic conformational change during
catalysis in the solution phase. Moreover, our work identified the importance of
stabilizing the negative charges of an incoming nucleotide and the pyrophosphate
product, a feature shared by a myriad of DNA polymerases.
125
3.6. Figures
Figure 3.1 Domain structure of human fPolλ, dPolλ, tPolλ, and Polβ. Each domain, with
amino acid residue numbers indicated above, is shown as a rectangle. The N-terminal 35
residues of fPolλ contain a nuclear localization signal motif as represented by the line.
126
Figure 3.2 Gap-filling DNA polymerase activity of fPolλ following HPG modification.
Reactions of fPolλ (10 μM) and 22-18/41G-mer (60 μM) were initiated by the addition of
100 µM dCTP at 37 ˚C and terminated after 1 minute by the addition of 0.37 M EDTA.
The negative control reaction (- Control) did not have dCTP. “HPG-E” and “HPG-ED”
denote HPG-modified apo-fPolλ and the HPG-modified binary complex (fPolλ•2218/41G-mer), respectively. The positive control reaction (+ Control) contained
unmodified fPolλ.
127
Figure 3.3 Representative segments of the MALDI-ToF MS spectra. (A) Apo-tPolλ
treated with HPG. (B) Binary complex of tPolλ•22-18/41G-mer DNA pre-formed and
then subjected to HPG modification. (C) Ternary complex of tPolλ•22ddC-18/41G-mer
DNA•dCTP was pre-formed and then treated with HPG. (D) The spectrum of unmodified
apo-tPolλ.
128
Figure 3.3
129
Figure 3.4 Tryptic digestion map of human fPolλ. Residues 0-575 are encoded by the
fPolλ gene. The protein contained N-terminal and C-terminal hexahistidine tags. The
tryptic peptide peaks that were detected by our MALDI-ToF instrument are underlined.
Arrow heads indicate that the peptide sequence continues on the next lower line at the
arrow tail. Protein domains are colored as follows: Yellow, N-terminal nuclear
localization sequence and hexahistidine tag; Orange, BRCT domain; Grey, Proline-rich
domain; Purple, dRPase domain; Blue, Fingers subdomain; Red, Palm subdomain; Green,
Thumb subdomain and C-terminal hexahistidine tag.
130
A
B
(continued)
Figure 3.5 Crystal structure of tPolλ detailing the interactions of R386, R275, ddTTP,
and the DNA template. The crystal structure of the ternary complex of tPolλ (blue),
gapped DNA substrate (black), and ddTTP (multi-colored) [81]. (A) Overall structure of
tPolλ ternary complex. (B) Close-up view detailing the interaction between R386 (red)
and ddTTP. (C) Close-up view detailing the interaction between R275 (red) and the 5’phosphate of the downstream primer terminus.
131
Figure 3.5 (continued)
C
132
1.2
1
0.6
k
obs
-1
(s )
0.8
0.4
0.2
0
0
10
20
30
40
50
60
70
dTTP ( M)
A
(continued)
Figure 3.6 Concentration dependence on the rate of dTTP incorporation into 21-19/41Amer (Table 3.1). (A) Pre-steady state kinetic parameters for fPolλ R386A. A preincubated solution of enzyme (300 nM) and 5′-[32P]-labeled 21-19/41A-mer DNA (30
nM) was mixed with increasing concentrations of dTTP for various times prior to being
quenched by 0.37 M EDTA. The observed rate constants (kobs) were plotted against the
concentrations of dTTP and the data were fit to Equation 2 (Section 3.3). For fPolλ
R386A, a kp of 1.3 ± 0.1 s-1 and a Kd of 15 ± 5 µM were determined. (B) Pre-steady state
kinetic parameters for fPolλ R386E. A kp of 0.005 ± 0.001 s-1 and a Kd of 1000 ± 410 µM
were determined.
133
Figure 3.6 (continued)
0.003
0.0025
0.0015
k
obs
-1
(s )
0.002
0.001
0.0005
0
0
200 400 600 800 1000 1200 1400 1600
dTTP ( M)
B
134
Figure 3.7 Arginine residues modified by HPG in the crystal structure of the ternary
complex of tPolλ. The ternary complex of tPolλ is colored blue. The gapped DNA
substrate is colored black. The incoming ddTTP is shown in multiple colors [81], and the
locations of arginine residues modified by HPG in apo-tPolλ are shown as red space
filling models.
135
Figure 3.8 Active site of tPolλ. tPolλ (blue) in the ternary complex with a gapped DNA
(black), after the chemistry step but before product release [81]. The charge of the leaving
group, pyrophosphate (PPi, red-orange), is stabilized by two Arg residues (red) and a
divalent metal ion (red sphere).
136
A
B
(continued)
Figure 3.9 Y-family DNA polymerase sequence alignment. Conservation of positively
charged residues involved in stabilizing the triphosphate moiety and/or the pyrophosphate
product. Based upon X-ray crystal structural analysis, amino acid residues within 4.0 Å
of the triphosphate moiety (red) were identified. Amino acid sequences of selected (A) Afamily, (B) B-family, (C) X-family, and (D) Y-family DNA polymerases were aligned
using ClustalW2. Conserved residues are shaded in blue and the conserved catalytic
aspartic acid residues are shaded in green.
137
Figure 3.9 (continued)
C
D
138
3.7. Tables
21-19/41A-mer
5’-CGCAGCCGTCCAACCAACTCA CGTCGATCCAATGCCGTCC-3’
3’-GCGTCGGCAGGTTGGTTGAGTAGCAGCTAGGTTACGGCAGG-5’
22-18/41G-mer
5’-CGCAGCCGTCCAACCAACTCAC GTCGATCCAATGCCGTCC-3’
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
22ddC-18/41G-mer
5’-CGCAGCCGTCCAACCAACTCAC GTCGATCCAATGCCGTCC-3’
3’-GCGTCGGCAGGTTGGTTGAGTGGCAGCTAGGTTACGGCAGG-5’
Table 3.1 DNA substrates. The downstream 18-mer and 19-mer strands were 5′phosphorylated. “C” denotes ddCMP.
139
Residue
R50
R55
R57
R275
R323
R386
R441
R446
R484
R485
R549
R561
R568
Apo
NP
NP
NP
+
+
+
+
+
+
+
+
+
+
tPolλ
Binary Ternary
NP
NP
NP
NP
NP
NP
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Apo
NP
NP
NP
+
+
+
+
+
+
+
+
+
+
dPolλ
Binary Ternary
NP
NP
NP
NP
NP
NP
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Apo
+
+
+
+
+
+
+
+
+
+
+
+
+
fPolλ
Binary Ternary
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Table 3.2 Summary of modified arginine residues. “+” indicates Arg residues susceptible
to modification by HPG. “-” indicates Arg residues shielded from HPG in the
nucleoprotein complexes. “NP” indicates that the indicated Arg residue is not present in
the protein constructs.
140
kp
Kd
kp/Kd
(s-1)
(μM)
(μM-1s-1)
WTb
3.9 ± 0.2
2.6 ± 0.4
1.5
1
R386A
1.3 ± 0.1
15 ± 5
0.087
17
R386E
0.005 ± 0.001
1000 ± 410
0.000005
300,000
fPolλ Mutant
Efficiency Ratioa
Table 3.3 Kinetic parameters of dTTP incorporation into single-nucleotide gapped 2119/41A-mer catalyzed by fPolλ variants at 37 C. aCalculated as (kp/Kd)WT/(kp/Kd)Mutant.
b
Values for the WT enzyme are from Fiala et. al. [288].
141
β-phosphate
DNA
polymerase
(PDB)
Human Polλ
(1XSN)
Family
X
None
Human Polλ
(1XSP)
X
Linked to primer
Human Polβ
(1BPY)
X
None
R183 (2.84 and 2.85
Å)
Pol mu
(2IHM)
X
None
R323 (3.23 Å)
Mouse TdT
(1KEJ)
Dpo4
(1JX4)
X
R454 (2.57 Å)
R336 (3.57 Å)
Y
R51 (3.13 Å)
Y10 (3.40 Å)
Dpo4
(2ATL)
Y
K159 (3.64 Å)
R51 (2.82 Å)
K159 (2.93 Å)
Y48 (3.71 Å)
F11 (3.03 and 3.03
Å)
K159 (2.93 Å)
Dpo4
(2AGO)
Y
None when
linked to primer
K159 (3.58, 3.65 and
3.82 Å)
Dbh
(3BQ1)
Y
None
F11 (2.93 Å)
Y10 (3.34 Å)
-phosphate
R420 (3.00 and 3.10
Å)
S417 (3.06 Å)
R420 (2.82 and 3.74
Å)
S417 (2.65 Å)
-phosphate
Notes and Reference
R386 (2.90
and 3.04 Å)
Backbone of S417
[81]
R386 (2.77,
3.39, 3.73 and
3.77 Å)
G426 (3.08 Å)
None
PPi bound
Backbone for S417
and G426
[81]
Some R nearby but
>4.0 Å
[121]
Side chain and
backbone for H329
[330]
Backbone for H342
[88]
dADP,
backbone of Y10
[331]
dCTP,
backbone of Y10 and
F11
[333]
K325 (2.83 Å)
H329 (2.93
and 3.05 Å)
H342 (2.84 Å)
N/A
R51 (2.89 and
3.53 Å)
Y10 (3.09 Å)
K159 (3.12
and 3.27 Å)
Y48 (2.55 Å)
R51 (3.04 and
3.94 Å)
Y10 (3.13 and
3.28 Å)
K160 (3.08,
3.17, and 3.59
Å)
R50 (3.17 and
3.22 Å)
dGMP-PPi structure
backbone of Y10
[334]
Backbone of Y10 and
F11
[329]
Table 3.4 Positively-charged residues that potentially stabilize the triphosphate moiety of
an incoming nucleotide and/or pyrophosphate product. (continued)
142
Table 3.4 (continued)
Yeast Pol eta
(2R8J)
Y
R73 (3.24 and
3.35 Å)
R73 (3.02 and 3.15
Å)
R67 (3.64 and
4.01 Å)
K289 (2.89 Å)
Human Pol iota
(2FLL)
Y
None
C37 (3.63 Å)
Human Pol kappa
(2OH2)
Y
None
F111 (2.95 Å)
Rev1
(2AQ4)
Y
None
N414 (2.94 Å)
T7
(1SKR)
A
K522 (2.85 Å)
H506 (2.75 Å)
G478 (3.40 Å)
Bacillus
stearothermophilus
DNA polymerase I
Fragment
(1LV5)
Taq
(2KTQ)
Taq
(3KTQ)
A
K706 (3.10 Å)
H682 (3.59 Å)
I657 (3.79 and 3.90
Å)
R71 (2.92 and
3.02 Å)
K77 (3.60 Å)
K214 (2.60 Å)
F38 (3.34 Å)
R144 (2.74,
3.64 and 3.69
Å)
K328 (2.81 Å)
A110 (3.28 Å)
R408 (2.69
and 2.77 Å)
K525 (3.02 Å)
C365 (2.62 Å)
R518 (2.83
and 3.33 Å)
K522 (3.60 Å)
G478 (2.71 Å)
R702 (2.91
and 3.18 Å)
Q656 (3.21Å)
A
None within 4.0
Å
K663 (3.01 and
3.94 Å)
None within 4.0 Å
RB69
(2OZS)
B
K560 (3.19 Å)
N564 (3.72 Å)
Phi29
(2PYL)
B
None
HIV-1 RT
(2IAJ)
RT
None
N387 (3.50 Å)
K383 (3.60 and 3.89
Å)
None
A
H639 (3.10 Å)
143
dCTP, 3′-Cisplatin
structure, AA
numbering is
according to PDB file
[335]
Backbone for C37
and F38
[336]
Backbone for A110
and F111
[337]
Backbone for C365
[338]
Backbone for G478
[339]
Backbone for Q656
[108]
None within
4.0 Å
R659 (2.97
and 2.87 Å)
open ternary
[340]
active ternary
[340]
R482 (2.80
and 2.94 Å)
K560 (2.80 Å)
K371 (3.90 Å)
[341]
K219 (3.07 Å)
Mutant, no DNA
RNA/DNA substrate,
ATP is bound
[343]
[342]
Chapter 4 - Preliminary investigation of the mechanism of Y-family DNA
polymerases using mass spectrometry
4.1. Introduction
The widely variable biochemical and structural properties of non-canonical DNA
polymerases, and especially those of the Y-family, suggest their mechanism of DNA
polymerization may vary considerably from the more well established mechanisms used
by replicative DNA polymerases. Replicative polymerases frequently exploit (to varying
degrees) the energy of nucleotide binding to drive a rate-limiting protein conformational
change preceeding a fast chemistry step [315] as a mechanism to enhance polymerase
fidelity. Other DNA polymerases employ mechanisms to enhance fidelity that utilize
rate-limiting transition states to enhance polymerase fidelity. For example, in Polβ (rabbit
origin) the free energy difference in chemical transition states between correct and
incorrect nucleotide base pairing [344]. Furthermore Pol , like other replicative
polymerases, is shown to utilize the induced-fit conformational change mechanism to
select and incorporate correct nucleotides during polymerization [143]. Interestingly
however, Pol is suggested to experience protein conformational as the rate-limiting step
for the incorporation of both matched and mismatched nucleotides [143]. In addition,
144
The Y-family enzymes possess relatively flexible and solvent accessible active
sites in order to accommodate bulky DNA lesions [331, 345]. However, Y-Family DNA
polymerases catalyze DNA synthesis over undamaged DNA with low fidelity and poor
processivity [331, 346-348]. The Y-family DNA polymerases have been identified in all
three domains of life, e.g. four in humans (DNA polymerases , , , and Rev1), two in
Escherichia coli (DNA polymerases IV and V) and one in S. solfataricus (Dpo4) (Section
1.2). Because Dpo4 can be expressed in E. coli and purified with a high yield, and
because it is the only functional Y-family enzyme in S.solfataricus, it has been
extensively studied in vitro as a prototype Y-family enzyme. Dpo4 catalyzes DNA
synthesis on an undamaged DNA template with a fidelity of one error per 1,000 to 10,000
nucleotide incorporations based on pre-steady-state kinetic analysis from 37 to 56 °C [30,
349, 350].
pre-steady state kinetic investigation into the mechanism of DNA polymerization
utilized by Dpo4 [351] suggests that although Dpo4 follows the induced-fit mechanism to
select correct nucleobase pairings, the rate-limiting step for incorrect nucleotide
incorporation is limited by chemistry, not conformational change [351]. Therefore,
employing methods similar to those described in Section 3.3, our ongoing research seeks
to further elucidate the mechanism of DNA polymerization and nucleotide selection
utilized by Dpo4 and possibly the mechanisms used by Y-family DNA polymerases in
general.
145
4.2. Materials
Preparation of Dpo4
Full-length Dpo4 fused to a C-terminal His6 tag was overexpressed and purified in
E. coli as described previously [352].
Synthetic oligodeoxyribonucleotides
The oligodeoxyribonucleotides (See Table 2.1) were purchased from Integrated
DNA Technologies (Coralville, IA) and purified by denaturing polyacrylamide gel
electrophoresis (17% acrylamide, 8 M urea, Tris-borate-EDTA running buffer). Their
concentrations were determined by UV absorbance at 260 nm with calculated extinction
coefficients. Each single-nucleotide gapped DNA substrate was prepared by heating a
mixture of 22-mer, and 41-mer in a 1:1.15 molar ratio, respectively, for 8 min at 95 °C
and then cooling the mixture slowly to room temperature over 3 h as described previously
[319].
Reaction buffer D
Reaction buffer D contained 50 mM Tris-Cl (pH 8.0 at 37 °C), 5 mM MgCl2, 50
mM NaCl, and 0.1mM EDTA and 10% glycerol. This reaction buffer was optimized
previously for transient state kinetic analysis of Dpo4 [30]. All reactions reported herein
were carried out in reaction buffer D at 25 °C, and all concentrations refer to the final
concentration of the components after mixing.
146
4.3. Methods
Mass spectrometry-based protein footprinting assay
Dpo4 (10 µM), Dpo4 (10 μM)•22/41-mer (20 μM) binary complex, and Dpo4 (10
μM)•22ddC/41-mer (20 μM)•dCTP (2 mM) ternary complex were subjected to chemical
modification by Sulfo-N-Hydroxysuccinimido-Biotin (Sulfo-NHS-Biotin). Sulfo-NHSBiotin reacts efficiently with primary amines with the concomitant release of Sulfo-Nhydroxysuccinimide, to form stable amide bonds and adds (226 Da) to the weight of
reacted lysine residues [322]. Sulfo-NHS-Biotin was adopted for footprinting purified
free Dpo4, and its binary and ternary complexes following empirical observations
indicating that lysine residues are more easily modified than arginine residues in this
enzyme. The Sulfo-NHS-Biotin treatments were carried out at 25 ˚C for 30 minutes and
terminated by the addition of 160 mM (final concentration) lysine in its free form. Dpo4
was then separated from DNA and dNTP by SDS-PAGE. The protein bands were
excised, destained, dehydrated, and digested with 1 g of trypsin in 50 mM NH4HCO3 at
25 C overnight.
Small molecular weight peptides were analyzed by MALDI-ToF MS using
AXIMA-CFR instrument (Shimadzu Scientific Instruments). The samples were analyzed
with an α-cyano-4-hydroxycinnamic acid matrix as described previously [324]. Sequence
data and Protein Prospector v4.0.6 (http://prospector.ucsf.edu) were used to identify
Dpo4 peptide peaks. Modified lysine residues were assigned by identifying mass peaks
147
that appear only in the spectra of Biotin-modified Dpo4 and that have a molecular weight
corresponding to the sum of the predicted peptide fragment plus the 226 Da Biotin
adduct. For accurate quantitative analysis of the modified peptide peaks, at least two
unmodified proteolytic peptide peaks were used as internal controls. A protection was
considered to be significant when the intensity of the given modified peptide peak
derived from Sulfo-NHS-Biotin treated free Dpo4 was reduced at least 10-fold in the
context of the nucleoprotein complexes. A modified peptide peak was considered
unprotected when the intensities of the given peptide obtained from free protein and
nucleoprotein complexes were within ± 20% of each other.
148
4.4. Preliminary results
Comparing the MS spectra for the Apo, Binary and Ternary complexes of Dpo4
after treatment with Sulfo-NHS-Biotin reveals that 17 out of a possible 39 (Table 4.1).
The Apo and Binary complex spectra suggest that there may be little change in the
structure of Dpo4 as it binds to DNA and forms the Binary complex. However,
comparing the Ternary complex to the Apo and Binary complexes, K278 and K282 are
shown to be protected from chemical modification.
Interestingly, when the crystal structure of the ternary complex of Dpo4 [353] is
examined closely, it can be seen that the primary amine of K282 is moved to within 3.28
Å of the carbonyl backbone oxygen of K339, strongly suggesting the presence of a
hydrogen bond. However, in the crystal structures of the Apo [353] and Binary [354]
complexes, K282 does not appear to form such a bond. Therefore, the observed
protection may be the result of the formation of a stable hydrogen bond between K282
and the backbone carbonyl oxygen of K339. Formation of such a bond would likely
require a conformational change in Dpo4 and may be evidence of such a change
occurring upon formation of the Ternary structure, but interestingly not upon formation
of the binary structure.
Finally, similar analysis of the crystal structures of Dpo4 does not reveal an
obvious reason for the observed protection of K278 (Table 4.1). However, it is possible
that factors such as crystal packing may have distorted the native position of K278 in one
149
or all three of the crystal structures. Therefore, no conclusion about this residue can yet
be drawn.
150
4.5. Figures
Figure 4.1 Crystal structure of Dpo4 showing K282.
151
Figure 4.2 Crystal structure of Dpo4 in the binary complex.
152
Figure 4.3 Crystal structure of Dpo4 in the ternary complex.
153
4.6. Tables
Lys Residue
52
56
65
146
148
152
159
193
195
201
212
221
223
243
278
282
329
Apo
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Binary
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Ternary
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Table 4.1 Summary of modified lysine residues in Dpo4. “+” indicates that the Lys
residue was susceptible to modification by Sulfo-NHS-Biotin. “-” indicates that the Lys
residues was shielded from Sulfo-NHS-Biotin modification in the nucleoprotein
complexes.
154
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