GENOMIC CONSEQUENCES OF DNA OXIDATION BY PEROXYNITRITE
by
William Louis Neeley
A.B., Chemistry (2001)
Washington University in St. Louis
Submitted to the Department of Chemistry in Partial
Fulfillment of the Requirements for the Degree of
DOCTOR OF PHILOSOPHY IN BIOLOGICAL CHEMISTRY
at the
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
February 2006
 2006 Massachusetts Institute of Technology. All Rights Reserved.
The author hereby grants to MIT permission to reproduce
and to distribute publicly paper and electronic
copies of this thesis document in whole or in part.
Signature of Author:
Department of Chemistry
November 14, 2005
Certified by:
John M. Essigmann
William and Betsy Leitch Professor of Chemistry and Toxicology
Thesis Supervisor
Accepted by:
Robert W. Field
Haslam and Dewey Professor of Chemistry
Chairman, Department Committee on Graduate Students
This doctoral thesis has been examined by a committee
of the Department of Chemistry as follows:
Steven R. Tannenbaum
Underwood-Prescott Professor of Toxicology and Chemistry
Chairman
John M. Essigmann
William and Betsy Leitch Professor of Chemistry and Toxicology
Thesis Supervisor
Peter C. Dedon
Professor of Toxicology and Biological Engineering
Gerald N. Wogan
Professor of Chemistry and Biological Engineering
2
GENOMIC CONSEQUENCES OF DNA OXIDATION BY PEROXYNITRITE
by
William Louis Neeley
Submitted to the Department of Chemistry on November 14, 2005
in Partial Fulfillment of the Requirements for the Degree of
DOCTOR OF PHILOSOPHY IN BIOLOGICAL CHEMISTRY
ABSTRACT
The radicals nitric oxide and superoxide are produced endogenously by activated
macrophages and neutrophils and combine in a diffusion-limited reaction to form
peroxynitrite, a powerful oxidizing and nitrating agent capable of damaging a variety of
biomolecules, including DNA. Of the four nucleobases of DNA, guanine has the lowest
oxidation potential and thus considerable attention has been given to the study of the
oxidation of this base by peroxynitrite. A variety of DNA lesions are generated from
guanine including guanidinohydantoin, spiroiminodihydantoin, oxaluric acid, urea, 2aminoimidazolone, and 5-guanidino-4-nitroimidazole. In order to assess the biological
significance and consequences of peroxynitrite-damaged DNA, it is essential that these
lesions be characterized for their genotoxic and mutagenic potential. This work focuses
on the elucidation of those properties.
In the first study, the 2’-deoxynucleoside of 5-guanidino-4-nitroimidazole was
chemically synthesized and incorporated into an oligonucleotide by the phosphoramidite
method. In the second study, the genotoxic and mutational properties of 2aminoimidazolone and 5-guanidino-4-nitroimidazole were determined in wild-type
uninduced and SOS-induced E. coli. In the third study, oxaluric acid was found to
hydrolyze to urea in a reaction catalyzed by magnesium cations and bicarbonate. The
genotoxic and mutational properties of oxaluric acid and urea were determined in wildtype uninduced and SOS-induced E. coli. In the fourth study, the genotoxic and
mutational properties of guanidinohydantoin and spiroiminodihydantoin were determined
in wild-type uninduced E. coli. In the fifth study, the genotoxic and mutational properties
of guanidinohydantoin, spiroiminodihydantoin, oxaluric acid, urea, 2-aminoimidazolone,
and 5-guanidino-4-nitroimidazole were determined in wild-type, polymerase II deficient,
polymerase IV deficient, polymerase V deficient, and polymerase II / polymerase IV /
polymerase V deficient E. coli under both uninduced and SOS-induced conditions. All of
the lesions studied were potent sources of mutations in vivo. Guanidinohydantoin,
spiroiminodihydantoin, urea, and 5-guanidino-4-nitroimidazole were significant blocks to
replication and were strongly dependent upon induction of the SOS system. Polymerase
V was responsible for the majority of translesion synthesis.
Thesis Supervisor: John M. Essigmann
Title: William and Betsy Leitch Professor of Chemistry and Toxicology
3
ACKNOWLEDGMENTS
Thus far in life, I have experienced no greater professional honor than the
privilege of attending MIT and receiving permission to write this thesis. No two people
have affected my life more, or are more responsible for my ability to reach this point,
than Mom and Dad. They have provided wisdom and love to me throughout my life, and
without their guidance I would have few achievements of which to speak. My intense
desire to make them proud is the single largest source of my motivation each and every
day. My siblings Andrena, David, Tom, and Margaret are my life-long best friends, and I
am immensely grateful for the love and support they have always provided, especially
during my time in graduate school. My love for and devotion to all of you forever
remains unquestioned.
I befriended an amazing group of people in graduate school. All of you are
destined for greatness and it’s going to be an amazing experience watching your careers
develop. For all the good times at the Muddy and elsewhere, cheers to Yuri Alekseyev,
Jeannette Allen, Charlie Hamilton, Paul Henderson, Hector Hernandez, Shawn Hillier,
Dave Laitar, Kaushik Mitra, Kyle Proffitt, Ryan Rieth, Jen Robbins, Peter Rye, Mo
Seyedsayamdost, Uday Sharma, and Amrit Sinha.
The Essigmann Laboratory has been a fantastic place to work in large part
because of the people there I have been privileged to know. I thank Paul Henderson for
getting me started on the oxidative DNA damage project, for teaching me many of the
techniques needed to conduct the research, and for being a good mentor. I also
collaborated with Drs. Yuriy Alekseyev, James Delaney, and Sarah Delaney. Thank you
all for your diligence, insight, and great scientific conversations. I am very grateful to
Drs. Uday Sharma and Kaushik Mitra for providing invaluable advice and instruction to
me on the art of organic synthesis. If my experiments succeeded, often a good suggestion
from a group member was responsible for this outcome. Bob Croy, Nicole Dinaut, Aida
Herrera, Shawn Hillier, John Marquis, Charles Morton, Pei-Sze Ng, Kyle Proffitt,
Jennifer Robbins, Denise Rodriguez, and Peter Rye contributed in one way or another
and I am very thankful to you all. I also thank our administrative assistant Kim Bond
Schaefer.
I thank my collaborators Jacquin Niles and Feng Gu from the Tannenbaum
Laboratory, James Muller and Cindy Burrows from the University of Utah, Sang Soo
Hah from Lawrence Livermore National Laboratory, and Olga Rechkoblit and Dinshaw
Patel from Memorial Sloan-Kettering Cancer Center for their hard work.
Instrumentation facilities in the Department of Chemistry and Biological
Engineering Division were essential for the work described in this thesis. I thank Li Li,
Dave Bray, and Pete Wishnok for their invaluable help.
The Department of Chemistry has been a phenomenal place to study chemistry.
The classes I took during my first year were excellent because of the professors teaching
them. Many thanks to Professors Dave Bartel (Department of Biology), Steve Buchwald,
4
Rick Danheiser, Barbara Imperiali, Alex Klibanov, Tam Rajbhandary (Department of
Biology), Larry Stern, and Joanne Stubbe for their efforts. I also thank Susan Brighton
for her devotion to the Department and graduate education.
I thank my Thesis Committee, Professors Pete Dedon, Steve Tannenbaum, and
Gerry Wogan for their valuable advice and for sacrificing their time. A tremendous
thank you must go to my advisor, Professor John Essigmann. John is a brilliant scientist
and has the rare ability to empathize with the people around him. Few men I have met in
this life are as kind, generous, and selfless as John Essigmann. You have been an
outstanding advisor and I thank for you accepting me into your laboratory.
WILLIAM L. NEELEY
Massachusetts Institute of Technology
Cambridge, Massachusetts
October 2005
5
For Mom and Dad
6
CONTENTS
COMMITTEE PAGE
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ABSTRACT
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ACKNOWLEDGMENTS .
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TABLE OF CONTENTS .
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ABBREVIATIONS.
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LIST OF FIGURES
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LIST OF SCHEMES
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LIST OF TABLES
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CHAPTER 1. Mechanisms of Formation, Genotoxicity, and Mutation of Guanine
Oxidation Products: A Survey of the Literature
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1.1.
Introduction
1.2.
Chemistry of Nitric Oxide
1.3.
Mechanisms of Guanine Oxidation Product Formation in Vitro
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1.3.1. Products derived directly from guanine
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1.3.2. Products derived directly from 7,8-dihydro-8-oxoguanine .
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1.3.3. Type II photosensitized oxidation of guanine and 7,8-dihydro8-oxoguanine .
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1.4.
Structural Aspects of Guanine Oxidation Products .
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1.5.
Repair of Guanine Oxidation Products
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1.6.
Genotoxic and Mutational Properties .
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1.6.1. In vitro properties of guanine oxidation products
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1.6.2. In vivo properties of guanine oxidation products
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1.7.
Biological Relevance .
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1.8.
Concluding Remarks .
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1.9.
Acknowledgment
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1.10.
References
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CHAPTER 2. Efficient Synthesis of DNA Containing the Guanine Oxidation-Nitration
Product 5-Guanidino-4-nitroimidazole: Generation by a Postsynthetic Substitution
Reaction
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2.1.
Abstract
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2.2.
Introduction
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2.3.
Experimental Procedures
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2.4.
Results and Discussion
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2.5.
Acknowledgment
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2.6.
References
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122
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CHAPTER 3. In Vivo Bypass Efficiencies and Mutational Signatures of the Guanine
Oxidation Products 2-Aminoimidazolone and 5-Guanidino-4-nitroimidazole
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126
3.1.
Abstract
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3.2.
Introduction
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3.3.
Experimental Procedures
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3.4.
Results
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3.5.
Discussion
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3.6.
Acknowledgment
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140
3.7.
References
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147
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CHAPTER 4. Urea Lesion Formation in DNA as a Consequence of 7,8-Dihydro-8oxoguanine Oxidation and Hydrolysis Provides a Potent Source of Point Mutations
153
4.1.
Abstract
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154
4.2.
Introduction
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155
4.3.
Experimental Procedures
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157
4.4.
Results and Discussion
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162
4.4.1. Hydrolysis of Oa to form Ua .
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162
4.4.2. In vivo mutagenesis by Oa and Ua under normal or SOS-induced
conditions
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4.5.
Acknowledgment
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4.6.
References
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179
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8
CHAPTER 5. The Hydantoin Lesions Formed From Oxidation of 7,8-Dihydro-8oxoguanine are Potent Sources of Replication Errors in Vivo
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185
5.1.
Abstract
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5.2.
Introduction
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5.3.
Experimental Procedures
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5.4.
Results
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5.5.
Discussion
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5.6.
Acknowledgment
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5.7.
References
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216
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CHAPTER 6. Effect of Escherichia coli Polymerases II, IV, and V on the Bypass
Efficiencies and Mutational Signatures of Guanine Oxidation Products
6.1.
Abstract
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217
6.2.
Introduction
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6.3.
Experimental Procedures
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6.4.
Results
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228
6.4.1. Translesion bypass efficiency .
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228
6.4.2. Mutation type and frequency .
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230
6.5.
Discussion
6.6.
6.7.
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234
Acknowledgment
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238
References
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262
FUTURE DIRECTIONS .
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CURRICULUM VITAE .
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271
9
ABBREVIATIONS
AP, apurinic/apyrimidinic
8-oxoG, 7,8-dihydro-8-oxoguanine
Ca, cyanuric acid
PAGE, polyacrylamide gel
Gh, guanidinohydantoin
electrophoresis
HPLC, high-pressure liquid
PNK, polynucleotide kinase
chromatography
pol α, calf thymus polymerase α
IPTG, isopropyl-β-D-thiogalactoside
pol β, human polymerase β
Iz, 2-aminoimidazolone
pol I, DNA polymerase I
Kf, Klenow fragment of DNA
pol II, DNA polymerase II
polymerase I
pol IV, DNA polymerase IV
Kf (exo−), Klenow fragment of DNA
pol V, DNA polymerase V
polymerase I without exonuclease
REAP, restriction endonuclease and
activity
post-labeling analysis of mutation
MALDI-TOF, matrix assisted laser
frequency
desorption ionization-time of flight
ss, single-stranded
8-MeOdG, 8-methoxy-2’-
Sp, spiroiminodihydantoin
deoxyguanosine
THF, tetrahydrofuran
MW, molecular weight
TLC, thin-layer chromatography
8-NO2-G, 8-nitroguanine
Ua, Urea
NI, 5-guanidino-4-nitroimidazole
WT, wild-type
Oa, oxaluric acid
X-gal, 5-bromo-4-chloro-3-indolyl-β-D-
ODN, oligodeoxynucleotide
galactoside
ONOO−, peroxynitrite
Z, oxazolone
10
LIST OF FIGURES
Figure 1.1.
Guanine oxidation products discussed in Chapter 1 .
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45
Figure 1.2.
Structure of DGh and other proposed precursors of Oa
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46
Figure 2.1.
1
Figure 2.2.
13
Figure 2.3.
1
Figure 2.4.
13
Figure 2.5.
UV-vis spectrum of 3A in CD2Cl2
Figure 2.6.
1
Figure 2.7.
13
Figure 2.8.
H NMR spectrum of 2 in DMSO-d6 .
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91
C NMR spectrum of 2 in DMSO-d6 .
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92
H NMR spectrum of 3A in CD2Cl2 .
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93
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UV-vis spectrum of 3 in CD2Cl2
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98
Figure 2.9.
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99
Figure 2.10.
13
C NMR spectrum of 4 in CD3OD .
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100
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101
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102
C NMR spectrum of 5 in CD3OD .
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103
Figure 2.14. UV-vis spectrum of 5 in H2O, pH 7 .
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104
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105
C NMR spectrum of 6 in CD2Cl2
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106
H NMR spectrum of 7 in CD3CN
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107
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108
Figure 2.19. ESI mass spectrum for 8NI and 9NI .
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109
Figure 2.20. MALDI-TOF mass spectrum for 8NI and 9NI
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110
Figure 2.21. Digestion analyses of 8NI and 9NI using SVPD
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111
Figure 2.22. Digestion analysis of 9NI using nuclease P1 .
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112
Figure 2.23. Representative melting curves for 8NI
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113
Figure 2.24. Representative melting curves for 8G
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115
Figure 2.26. Agarose gel analysis of the M13 genome constructions
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Figure 2.27. HPLC traces and UV-vis spectrum of 8NI
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117
C NMR spectrum of 3A in CD2Cl2 .
H NMR spectrum of 3 in CD2Cl2
C NMR spectrum of 3 in CD2Cl2
H NMR spectrum of 4 in CD3OD
Figure 2.11. gCOSY spectrum of 4 in CD3OD
Figure 2.12.
1
Figure 2.13.
13
.
H NMR spectrum of 5 in D2O
Figure 2.15.
1
Figure 2.16.
13
Figure 2.17.
1
Figure 2.18.
31
H NMR spectrum of 6 in CD2Cl2
P NMR spectrum of 7 in CD3CN
Figure 2.25. Energy-minimized (AM1) structure of dNI (5)
11
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.
Figure 3.1.
Primary products of ONOO− oxidation and nitration of guanine
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141
Figure 3.2.
Construction of genomes containing a site specific modification
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142
Figure 3.3.
Assay for determining the efficiency of DNA polymerase bypass
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143
Figure 3.4.
Bypass efficiency of Iz, NI, and a THF abasic site relative to G
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144
Figure 3.5.
REAP assay for determining mutation type and frequency .
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145
Figure 3.6.
REAP assay results for G, Iz, and NI .
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146
Figure 4.1.
Translesion DNA polymerase bypass and REAP assays
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169
Figure 4.2.
ESI mass spectra and HPLC traces (insets) of 8-oxoG, Oa, and Ua .
170
Figure 4.3.
Hydrolysis of Oa to form Ua in an oligonucleotide .
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171
Figure 4.4.
Oxalate formation curves for the decomposition of Oa
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172
Figure 4.5.
Relative bypass efficiency for Oa and Ua
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173
Figure 4.6.
SOS induction allows preferential bypass of blocking lesions
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174
Figure 4.7.
TLC showing mutation frequency and type .
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175
Figure 5.1.
Structures of lesions and the oligonucleotide sequence used .
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201
Figure 5.2.
HPLC chromatogram for the oligonucleotide containing Gh .
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202
Figure 5.3.
HPLC chromatogram for the oligonucleotides containing Sp
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203
Figure 5.4.
MALDI-TOF mass spectra for Gh
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204
Figure 5.5.
MALDI-TOF mass spectra for Sp1 .
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205
Figure 5.6.
MALDI-TOF mass spectra for Sp2 .
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206
Figure 5.7.
Translesion DNA polymerase bypass data for Gh, Sp1, and Sp2
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207
Figure 5.8.
REAP assay and mutation frequency analysis for Gh, Sp1, and Sp2
Figure 6.1.
Primary products of ONOO− oxidation and nitration of G
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239
Figure 6.2.
HPLC trace and MALDI-TOF spectrum of 8-oxoG oligonucleotide .
240
Figure 6.3.
HPLC trace and MALDI-TOF spectrum of Gh oligonucleotide
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241
Figure 6.4.
HPLC trace and MALDI-TOF spectrum of Sp1 oligonucleotide
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242
Figure 6.5.
HPLC trace and MALDI-TOF spectrum of Sp2 oligonucleotide
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243
Figure 6.6.
HPLC trace and MALDI-TOF spectrum of Oa oligonucleotide
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244
Figure 6.7.
HPLC trace and MALDI-TOF spectrum of Ua oligonucleotide
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245
Figure 6.8.
HPLC trace and MALDI-TOF spectrum of NI oligonucleotide
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246
Figure 6.9.
Genome quantification gel and standard curve
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247
Figure 6.10. Bypass efficiency of DNA lesions relative to G in wild-type cells .
248
12
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208
Figure 6.11. Bypass efficiency of DNA lesions in pol II deficient cells
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249
Figure 6.12. Bypass efficiency of DNA lesions in pol IV deficient cells .
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250
Figure 6.13. Bypass efficiency of DNA lesions in pol V deficient cells .
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251
Figure 6.14. Bypass efficiency of DNA lesions in triple knockout cells .
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252
Figure 6.15. REAP assay results for wild-type cells
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253
Figure 6.16. REAP assay results for 8-oxoG
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254
Figure 6.17. REAP assay results for Gh
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255
Figure 6.18. REAP assay results for Sp1 .
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256
Figure 6.19. REAP assay results for Sp2 .
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257
Figure 6.20. REAP assay results for Oa
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Figure 6.21. REAP assay results for Ua
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Figure 6.22. REAP assay results for NI
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13
LIST OF SCHEMES
Scheme 1.1. Proposed mechanisms for G oxidation products
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47
Scheme 1.2. Proposed mechanisms for 8-oxoG oxidation products
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48
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49
Scheme 1.3. Proposed mechanisms for 8-oxoG products by peroxynitrite
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50
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118
Scheme 2.2. Synthesis of 5 and the convertible nucleoside phosphoramidite 7
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119
Scheme 4.1. Oa hydrolyzes to from Ua and oxalate
Scheme 1.4.
O2 oxidation of G and 8-oxoG
.
Scheme 2.1. Synthesis of the convertible nucleoside 4
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176
Scheme 4.2. Proposed mechanism for catalytic hydrolysis of Oa .
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177
14
.
LIST OF TABLES
Table 1.1.
Common names that appear in the literature for G oxidation products
51
Table 1.2.
Excision of Gh and Sp by Nei, Nth, and MutM
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52
Table 1.3.
Summary of in vitro standing start primer extension results .
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53
Table 1.4.
Bypass efficiencies and mutation frequencies for 8-oxoG products .
54
Table 2.1.
NOE difference for 4 in CD3OD (%) .
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120
Table 2.2.
Melting Temperatures and Free Energies of Duplexes
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121
Table 4.1.
Bypass efficiencies and mutation frequencies for 8-oxoG products .
178
Table 5.1.
Bypass data for G, Gh, Sp1, Sp2, and AP
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209
Table 6.1.
E. coli strains used in this work
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261
15
.
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CHAPTER 1
Mechanisms of Formation, Genotoxicity, and Mutation of
Guanine Oxidation Products: A Survey of the Literature
16
Chapter 1 – The Literature
1.1. Introduction
Endogenous and exogenous oxidants continually assault the genome. A variety of
deleterious processes and conditions (1) may result from DNA oxidation, including aging
(2-5), carcinogenesis (6-8), and neurological syndromes such as Alzheimer’s disease (9)
and amyotrophic lateral sclerosis (ALS) (10). DNA oxidation can result from its reaction
with compounds of environmental origin such as transition metals (11) or from reactive
oxygen species released during metabolism of exogenous organic compounds such as
benzo(a)pyrene (12). DNA also can be oxidized by reactive oxygen and nitrogen species
produced during cellular processes such as respiration and inflammation (13, 14).
Oxidation of DNA leads to the formation of an assortment of oxidation products
(15). Relative to the other DNA nucleobases, oxidation of guanine (G) occurs most
readily due to its low oxidation potential (16). Sequence context modulates the oxidation
potential of G in DNA. For example, ab initio calculations and experimental data show
that the 5’-G of stacked GG and GGG sequences oxidize more easily than a lone G in
DNA and that the 5’-G of GGG oxidizes more easily than the 5’-G of GG (17, 18). The
resultant radical cations (holes) generated in DNA from one electron abstraction can
migrate long distances before being trapped (19-21). Consequently, there is speculation
that GC rich domains outside the coding regions of genes may serve to protect the
genome from mutagenesis by oxidation products derived from trapped holes (22). 7,8Dihydro-8-oxoguanine (8-oxoG), an oxidation product of G and commonly used
biomarker of oxidative stress, is more easily oxidized than G and can therefore serve as a
better trap (23). Stacking with G improves the trapping ability (lowers the oxidation
potential) of 8-oxoG in the sequence context 5’-(8-oxoG)G-3’. Interestingly, in the
sequence context 5’-G(8-oxoG)-3’, G is preferentially oxidized with a ratio of about 1.5:1
demonstrating the strong influence of sequence context on oxidation potential (24).
Numerous studies show a predominance of GCÆAT, GCÆTA, and GCÆCG
mutations after replication of in vitro oxidized DNA in cells or upon exposure of cells to
oxidants (25-42). Importantly, 8-oxoG does not induce GCÆCG mutations, so these
results suggest additional G-derived lesions are responsible for the observed mutagenesis.
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Chapter 1 – The Literature
Indeed, a multitude of products resulting from G oxidation has been reported (Figure 1.1,
see Table 1.1 for the full names).
Many reports over the last several years have detailed the formation and
properties of G oxidation products. This review will focus specifically on the products
shown in Figure 1.1 and summarize the current understanding with regard to the
mechanisms of formation, effect on DNA structure and stability, repair potential, and
genotoxic and mutagenic prowess of these lesions. 8-OxoG was recently reviewed (15)
and will not be discussed except in the context of its oxidation products.
Formamidopyrimidine-G (FapyG) lesions have also been reviewed recently (43).
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Chapter 1 – The Literature
1.2. Chemistry of Nitric Oxide
A specific focus of the work in this thesis is the role of nitric oxide (•NO) and its
derivatives in carcinogenesis. Therefore, a brief discussion of the chemistry of •NO is
appropriate for placing the contents of this thesis in context.
•
NO is an important biological molecule that is involved in cellular signaling and
the immune response. An enzyme, known as nitric oxide synthase (NOS), uses Larginine, O2, and NADPH as substrates to generate •NO, citrulline, and presumably H2O
(44). Three general isoforms of this enzyme exist and they can be divided into two
groups.
The first group consists of Ca2+/calmodulin dependent isoforms, namely
neuronal NOS (nNOS) (45) and endothelial NOS (eNOS) (46), whereas the second group
is generally Ca2+/calmodulin independent and consists of inducible NOS (iNOS) (47).
With regards to inflammation, iNOS is the most important isoform since it is induced in a
variety of tissues including macrophages upon exposure of the tissues to signals such as
γ-interferon and lipopolysaccharide (LPS) (48). Whereas small amounts of •NO are
produced by nNOS and eNOS for use as a signaling agent (pM-nM), iNOS produces
much higher levels of •NO (µM) for cytotoxic purposes (49).
The chemistry of •NO has been reviewed (50, 51) and will be summarized here.
The relatively unreactive •NO reacts to form several reactive products including nitrous
anhydride (N2O3) and peroxynitrite (ONOO−).
N2O3 is the major nitrosating agent
formed from •NO, and the rate of formation is first order with respect to O2 and second
order with respect to •NO.
The reaction of N2O3 with DNA results in nitrosative
deamination.
Formation of ONOO− results from the combination of •NO with O2•−, which react
at a rate between 6.6 and 19 × 109 M-1 s-1. It is important to note that the consumption of
O2•− by superoxide dismutase occurs at a rate several fold slower than the rate of reaction
with •NO thereby allowing the formation of ONOO− to compete with O2•− dismutation.
ONOO− can protonate to form peroxynitrous acid (ONOOH), which has a pKa of 6.8. In
19
Chapter 1 – The Literature
the absence of CO2 and at physiological pH, ONOOH decomposes with a t1/2 of 1.9 s (52)
to form •NO2 and •OH in a solvent cage. Approximately 67% of the time, the two
radicals recombine to form NO3− and H+; however about 33% of the time, the radicals
escape the solvent cage and become free radicals that can participate in other chemistries.
DNA damage by these ONOO− decomposition products occurs primarily on the sugar
moiety, leading to strand breaks and oxidized abasic site formation.
The addition of CO2 to the reaction mixture profoundly changes the chemistry of
ONOO− and decreases the t1/2 to about 50 ms. CO2 reacts with ONOO− at a rate of 3-6 ×
104 M-1 s-1 to form nitrosoperoxycarbonate (ONOOCO2−). ONOOCO2− decomposes to
CO3•− and •NO2 that are initially in a solvent cage.
As in the case of ONOOH
decomposition, these radicals recombine about 67% of the time to form NO3− and CO2
and about 33% of the time diffuse out of the solvent cage becoming free radicals. The
major route of ONOO− decomposition should occur via CO2 since CO2 exists at
concentrations up to 1 mM physiologically. A difference of central importance caused by
the presence of CO2 is the shift in the predominant DNA damage from deoxyribose to
guanine bases. In the following sections, the current understanding of guanine base
damage is reviewed.
20
Chapter 1 – The Literature
1.3. Mechanisms of Guanine Oxidation Product Formation in Vitro
Oxidation products of G are believed to result from two main pathways, the
difference being the presence or absence of 8-oxoG as an intermediate. The first two
sections deal with products formed primarily as a result of one- and two-electron
oxidations of G and 8-oxoG. Each section is divided according to reactive species, and
the products formed as a result of reaction with these species are discussed. The third
section summarizes findings from 1O2 oxidation of G and 8-oxoG.
The major
mechanistic pathways are summarized in Schemes 1.1-1.4.
1.3.1. Products derived directly from guanine
A variety of oxidants abstract a single electron from G such as HO• (53, 54),
radicals derived from ONOO− (55-58), Br2•− and SO4•− (59), light (photoionization) (60,
61) and riboflavin (type I photosensitization) (53, 62, 63). This reaction generates a
guanine radical cation (G•+) that, with a pKa of 3.9 at the nucleoside level (59),
deprotonates at physiological pH to produce the neutral guanine radical (G(−H)•). The
deprotonation of G•+ at the nucleoside level and in DNA occurs with a rate constant on
the order of 107 s-1 at neutral pH (64). The lifetime of this radical depends on the
presence of reactive species, but in the absence of these species, the lifetime can reach ~5
seconds (65).
The reaction of G(−H)• with radicals such as O2•−, •NO2, and CO3•− occurs rapidly
and results in the formation of oxidative DNA lesions. A major product of this process is
Iz and its hydrolysis product Z (reviewed in (66)). In the literature, these products are
sometimes referred to together (Iz + Z) and at other times separately (Iz or Z).
Combination of O2•− with G(−H)• occurs with a rate constant of (4.7 ± 1.0) × 108 M-1 s-1
in both single- and double-stranded oligonucleotides.
This reaction results in the
formation of Iz as the major product (Scheme 1.1) and 8-oxoG, Sp, and Gh as minor
products (67), although it is likely Sp and Gh result from further oxidation of the 8-oxoG
produced during the reaction.
In contrast to single-stranded oligonucleotides,
dGMP(−H)• and G(−H)• nucleoside react faster with O2•− at rates of (1.3 ± 0.3) × 109 M-1
21
Chapter 1 – The Literature
s-1 (68) and about 3 × 109 M-1 s-1, respectively (69). Since these reactions occur in
aerated solution, Cadet originally proposed that under γ-radiolysis conditions dG(−H)•
(formed by reaction of dG with HO•) reacts with 3O2 (53).
However, γ-radiolysis
produces hydrated e− that are trapped by 3O2 to form O2•− at a rate of 1.9 × 1010 M-1 s-1
(70), and more recent experiments show that 3O2 reacts with G(−H)• at least three orders
of magnitude slower than does O2•−, making the reaction with O2•− kinetically more
favored (69). At the nucleoside level, Iz and its hydrolysis product Z are the major
products of HO• and riboflavin (type I photosensitization) oxidation (71) and represent
80% of the products (53).
The combination of G(−H)• with •NO2 (generated photochemically) has been
studied, and the reaction proceeds with a rate of ~4.3 × 108 M-1 s-1 in single- and doublestranded DNA, with the major products being NI and 8-NO2-G (Scheme 1.1) (72). The
formation of these products indicates reaction at the C5 (for NI) or C8 (for 8-NO2-G)
position of G(−H)•. NI is a chemically stable lesion (73, 74), but 8-NO2-G depurinates
with a half-life of about 1 h at 37 °C (pH 7.2) at the nucleoside level (41). Half-lives of
about 20 h at 23 °C (pH 7) (75) and 31 h at 25 °C (pH 8) (76) have been reported for 8NO2-G in single-stranded oligonucleotides, though a half-life as low as 4 h has also been
reported for 8-NO2-G in DNA at 37 °C (pH 7.4) (77). Joffe et al. demonstrated that the
environment of the parent G affects the ratio of 8-NO2-G to NI formation and found that
the ratio decreases from 3.4 in 2’,3’,5’-tri-O-acetylguanosine to 2.1-2.6 in single-stranded
oligonucleotides to 0.8-1.1 in double-stranded oligonucleotides. The maximum yield of
these lesions in a single-stranded oligonucleotide containing one G residue is 28% (74).
Campbell and coworkers recently proposed a mechanism for dissociation of
guanosine diphosphate (GDP) from Ras proteins (proteins involved in signal
transduction) through oxidation of GDP to NI. •NO activates Ras through the exchange
of bound GDP for bound GTP, and the study demonstrates that •NO2 produced from the
oxidation of •NO by O2 reacts with the protein to form a thiyl radical at a cysteine residue
that oxidizes the bound GDP. Key interactions between the nucleotide and the protein
22
Chapter 1 – The Literature
are disrupted during this process and cause the nucleotide to dissociate from the protein
and combine with a second equivalent of •NO2 to form 5-NO2-GDP, which collapses to
form the nucleotide diphosphate of NI (78, 79).
Treatment of G with ONOO− also generates 8-NO2-G and NI (55). ONOO−
breaks down to form •NO2 and either HO• or CO3•−, depending on the absence or
presence of CO2, and therefore can cause a net oxidation and nitration of G (51).
Reaction of CO3•− with dGMP occurs with a rate of (6.6 ± 0.7) × 107 M-1 s-1, which is
about 10 times faster than the reaction with dATP, dCTP, or dTTP. CO3•− reacts with a
G-containing oligonucleotide at a rate of (1.9 ± 0.2) × 107 M-1 s-1 (80). Treatment with a
50-fold excess of ONOO− produces NI with a yield of 2.3 ± 0.2% in a single-stranded
oligonucleotide and 0.3 ± 0.05% in a double-stranded oligonucleotide (73).
The
oxidation of 8-nitropurines by ONOO− has been investigated. Oxidation of 9-ethyl-8nitroxanthine (a model compound of 8-NO2-G) produces uric acid (a model compound of
8-oxoG), suggesting 8-oxoG may result from oxidation of 8-NO2-G (81). N2-NO2-dG
has been observed to form as a minor product when •NO and O2 are bubbled through a
solution of dG, and the yield of N2-NO2-dG increases with increasing levels of •NO or O2
and with increasing pH. 8-NO2-dG, 8-nitroxanthine, and xanthine also form during this
reaction. Incubation at pH 7.4 and 37 °C for one week does not cause decomposition of
N2-NO2-dG, indicating the lesion is a stable product (82). Another product from the
reaction of ONOO− with dG, nox-G, has been reported once (83).
Mn–TMPyP/KHSO5 (a two electron oxidant of G) oxidizes dG to generate dG+.
Presumed capture of the cation by HSO5− leads to the formation of Iz in 90% yield (84).
Oxidation of Iz by Mn–TMPyP/KHSO5 forms a putative N-oxide of Iz that has a half-life
of 2 h at room temperature and pH 7 (84). Oxidation of a GpT dinucleotide by Mn–
TMPyP/KHSO5 generates both Iz and DGh (85).
The half-life of DGh in an
oligonucleotide is about 5 h at 37 °C in H2O (86) and in a dinucleotide is 8 h at 0 °C in
pH 6.5 buffer (87). In one study, hydrolysis of DGh was proposed to form Iz (88), but
this transformation does not occur (87). Hydrolysis of DGh instead leads to Oa and, at
23
Chapter 1 – The Literature
the dinucleotide level, NMR data show that Oa adopts a linear structure with a bond
between N1 of Oa and C1’ of the sugar (85, 89) rather than a branched structure with a
bond between N3 of Oa and the sugar. Experiments with ESR spectroscopy at 77 K
reveal that further oxidation products of 8-oxoG that contain α-diketo groups, such as Oa,
can trap electrons to produce anion radicals (90). Also, Oa hydrolyzes to form the
tertiary lesion urea (Ua) which, historically, is more typically associated with pyrimidine
oxidation (91). Divalent metal cations and HCO3− catalyze the hydrolysis of Oa to Ua,
and the half-life of Oa nucleoside under these conditions is about 40 h (92).
The structure of DGh was determined by reducing the putative DGh with NaBH4
and then characterizing the resultant Gh product with NMR (87, 89). Other structures (3,
4, 5) (Figure 1.2) with molecular formulae identical to DGh and also leading to the
formation of Oa have been proposed to form under other oxidation conditions (93-95);
however, Meunier and coworkers have proposed that these other compounds are actually
DGh.
Reaction of 3’,5’-di-O-acetyl-dG with HOCl at pH 7.4 generates many products
including 8-Cl-dG, Sp, Gh, Iz, Z, and DIz (96). At neutral pH, Sp is the major product,
whereas at pH 5 Iz, DIz, and Gh predominate. In accordance with observations from
previous studies of pH dependence (97, 98), Gh is the principal product when the pH is
less than 5 (96). The Sp and Gh lesions formed during the oxidation by HOCl originate
from 8-oxoG and are discussed in more detail in the next section.
1.3.2. Products derived from 7,8-dihydro-8-oxoguanine
As with G, one electron oxidation of 8-oxoG forms a radical cation (8-oxoG•+).
Additionally, due to the low oxidation potential of 8-oxoG (E7 = 0.74 V/NHE) (23)
relative to G (E7 = 1.29 V/NHE) (16), 8-oxoG can serve as an effective hole trap in DNA.
The oxidation of 8-oxodG by G(−H)• nucleoside occurs with a rate of 4.6 × 108 M-1s-1
(23). Upon oxidation, the pKa of the 8-oxoG nucleoside decreases from 8.6 (99) to 6.6
for 8-oxoG•+; therefore, 8-oxoG•+ and 8-oxoG(−H)• are present in similar proportions at
neutral pH (23, 100).
24
Chapter 1 – The Literature
Shafirovich and coworkers investigated the oxidation of 8-oxoG-containing
oligonucleotides using kinetic laser spectroscopy and mass spectrometry (101). In this
system, irradiation of a 2-aminopurine and 8-oxoG-containing oligonucleotide induces
the one-electron oxidation of 8-oxoG. The irradiation also produces hydrated e− that are
captured by O2 to form O2•−. Reaction of 8-oxoG(−H)• with O2•− occurs with a rate
constant of ~1 × 108 M-1s-1 in single- and double-stranded DNA and leads primarily to
the formation of Oa through the intermediate DGh (Scheme 1.2).
When Cu,Zn-
superoxide dismutase (Cu,Zn-SOD) is added to the reaction, the lifetime of 8-oxoG(−H)•
is 3-5 seconds in single-stranded DNA and 15-20 seconds in duplex DNA. 8-OxoG(−H)•
exhibits low reactivity with O2 (<102 M-1 s-1) and, in the presence of Cu,Zn-SOD, instead
combines with H2O to form Sp as the major product (101). If 8-oxoG(−H)• is not
oxidized prior to addition of H2O, then formation of Sp likely involves addition of H2O to
8-oxoG•+. Provided the pKa of 8-oxoG•+ does not differ greatly in an oligonucleotide
versus at the nucleoside level, a substantial amount of 8-oxoG•+ should be present in the
pH range of 7-7.5 that is generally used experimentally in G and 8-oxoG oxidation
mechanistic studies. For close to 20 years, the structure of Sp was misassigned as 4,8dihydro-4-hydroxy-8-oxo-2’-deoxyguanosine (102-106). Evidence from HPLC, UV-vis
spectroscopy, and ESI-MS/MS lead to the reassignment of the structure as Sp (107), and
further proof of the structure was provided using SELINQUATE NMR (108).
The role of pH in the photooxidation of 2’,3’,5’-tri-O-acetyl-8-oxoG by riboflavin
was investigated by Burrows and coworkers, and the amounts of each product formed
were recorded (97). Riboflavin reacts primarily by a type I mechanism (103, 109), which
leads to a one-electron oxidation of 8-oxoG. At pH 7, the predominant product is Sp,
whereas Gh and Iz + Z are trace products. The second major product that makes up the
remaining mass balance was assigned as Iaox (4), but another possible structure for this
compound is DGh (Figure 1.2) (87). At pH 6, 4 makes up more than 50% of the product
mixture and Sp and Gh balance the remaining mass in approximately equal proportions.
Addition of superoxide dismutase (SOD) to the reaction decreases the amount of 4
formed to about 25% of the product mixture.
25
Sp and Gh again form in similar
Chapter 1 – The Literature
proportions and make up the remaining mass. At pH 8.6, Iz + Z and Sp are the major
products and 4 is the minor product, but in the presence of SOD, Sp is the major product
and Iz + Z are the minor products. Riboflavin can reduce O2 to O2•− (110), so the
differences in product distribution at the various pH values are possibly due to a
difference in reactivity of O2•− with the 8-oxoG radical when it is protonated versus
neutral and a competing reaction of O2•− with protonated riboflavin at pH 7.
An
interesting difference in the chemistry of G versus 8-oxoG is that Iz + Z are only formed
from 8-oxoG in substantial amounts under alkaline conditions (97).
There was controversy about whether Gh exists as a mixture of four isomers
(diastereomers of two structural isomers, Gh and Ia) or two diastereomers (87, 95). The
current consensus is that Gh is an equilibrating mixture of two diastereomers, although Ia
may exist in very low quantities (below the detection limit of NMR spectroscopy) (111).
8-OxoG in a single-stranded oligonucleotide reacts with CO3•− with a rate
constant of (3.2 ± 0.4) × 108 M-1 s-1, which is about 10 times faster than reaction with the
same oligonucleotide containing a single G instead of 8-oxoG [(2.4 ± 0.3) × 107 M-1 s-1].
Reaction of a self-complementary oligonucleotide containing a single G residue reacts
about half as fast as a single-stranded oligonucleotide containing a single G [(1.4 ± 0.2) ×
107 M-1 s-1], and an oligonucleotide containing no G residues reacts even slower [(5.7 ±
0.6) × 106 M-1 s-1]. The major end product of CO3•− oxidation of G and 8-oxoG is Sp in
both single and double-stranded DNA, and the formation of Sp from G appears to go
through 8-oxoG as an intermediate (Schemes 1.1 and 1.2) (112). Whereas reaction of
CO3•− with a G-containing single-stranded oligonucleotide generates Sp with a maximum
yield of 60%, continued production of CO3•− causes a slow degradation of Sp (112),
presumably due to oxidation of the lesion. Interestingly, CO3•− is the source of the
oxygen atom incorporated into Sp when this lesion forms from 8-oxoG through this
scheme. Oxidation of G to Sp proceeds through 8-oxoG as an intermediate and results in
the incorporation of two oxygen atoms from two equivalents of CO3•− (113).
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Chapter 1 – The Literature
Another mechanism of Sp formation (Scheme 1.2) involves •NO2 addition to 8oxoG(−H)• and occurs with a rate constant of ~4.3 × 108 M-1s-1 in single- and doublestranded DNA, yielding Sp as the major product at pH 7.5. The oxygen atom in Sp
originates from water, not •NO2 as determined isotopically (72).
This result has
significant mechanistic implications for ONOO−-related chemistry since •NO2 is a
product of ONOO− breakdown.
Tannenbaum and coworkers have investigated the role of ONOO− flux in the
identity and distribution of 8-oxoG oxidation products (98). Two groups of products
were identified with the major mechanistic difference being the identity of the attacking
nucleophile after the initial oxidation of 8-oxoG (Scheme 1.3). When a bolus of ONOO−
is used (high initial concentration of ONOO−), ONOO− can add nucleophilically to the
oxidized 8-oxoG to form DGh as the major product along with NO2-DGh and CAC as
minor products. When ONOO− is infused (low flux), water is the source of exogenous
oxygen atoms and the resultant products are Sp, Gh, and HICA (114). The ratio of Sp to
Gh varies dramatically depending on the pH of the reaction solution. Below pH 5.8, Gh
forms preferentially, whereas above pH 5.8 Sp predominates. It was therefore proposed
that the pKa of the Sp and Gh precursor, 5-hydroxy-8-oxo-7,8-dihydroguanine (5-HO-8oxoG), is 5.8. Initial studies of peroxynitrite oxidation of 8-oxoG revealed that at low
[ONOO−]/[DNA]
ratios
(<5),
3a-hydroxy-5-imino-3,3a,4,5-tetrahydro-1H-
imidazo[4,5d]imidazol-2-one (5) is the major product and hydrolysis of this compound
leads to Oa.
A more recent study showed that 5 is likely DGh (87).
At high
[ONOO−]/[DNA] ratios (>10), Ca and its precursor CAC are the major products (94).
Whereas the discussion above shows that the ratio of Sp to Gh formed from
oxidation of 8-oxoG is highly dependent on pH, other factors are also important.
Burrows and coworkers investigated the oxidation of 2’,3’,5’-tri-O-acetyl-8-oxoG by the
one-electron oxidants Na2IrCl6, K3Fe(CN)6, and CoCl2/KHSO5 and found that at pH 7,
the major product of the reaction is Sp regardless of the oxidant used (115). However,
starting with unprotected 8-oxodG, Henderson and coworkers found that Gh is the major
27
Chapter 1 – The Literature
oxidation product over the pH range 4.6 to 8.4 (116). In duplex DNA, oxidation of 8oxoG by Na2IrCl6 also produces Gh as the major product (117, 118). These results reveal
an underlying complexity to the mechanism of Sp and Gh formation. In the presence of
peptides, crosslinks between Sp or Gh and a peptide form upon oxidation of 8-oxoG in
DNA (119).
Oxidation of duplex DNA with Cr(VI)/ascorbate (ascorbate reduces Cr(VI) to
Cr(IV)) at pH 7.0 converts 1.44 ± 0.06% of all guanines to Sp when 50 uM Cr(VI) is
used and 0.315 ± 0.05% of guanines to Sp when 3.1 uM Cr(VI) is used, and the amount
of Sp formed is about 20-fold higher than 8-oxoG (120). Reaction of peroxo-Cr(V) with
calf thymus DNA at pH 7.2 yields Gh as a major product and 8-oxoG as a minor product,
and reaction of 8-oxodG under the same conditions also yields Gh as the major product
(121). These contrasting results with respect to which product forms predominantly
suggest that at least in some cases the reaction pathways to Sp and Gh in duplex DNA are
influenced by the nature of the oxidant.
Treatment of an oligonucleotide containing A, T, C, and G with 100 uM Cr(V)Salen shows piperidine cleavable G specific oxidation, and the amount of piperidine
cleavage does not increase upon posttreatment with Na2IrCl6 suggesting that any 8-oxoG
formed is further oxidized. Using an oligonucleotide containing a site-specific 8-oxoG
lesion, oxidation with Cr(V)-Salen occurs at the site of the 8-oxoG lesion with high
selectivity and generates both Sp and Gh (122, 123).
Reaction of 8-oxodG with HOCl produces Sp as the major product, and the
mechanism is believed to involve addition of the highly electrophilic Cl+ to the C4-C5
double bond. A solution of 1 mM 8-oxodG with 0.5 mM HOCl forms Sp in 76% yield
with respect to the amount of HOCl present. For comparison, oxidation of 8-oxoG with
CoCl2/KHSO5 or ONOO−/N-AcCys produces Sp with yields of 32% and 4%, respectively
(124, 125). Certain biological compounds hinder the oxidation of dG and 8-oxodG. For
example, the tea polyphenol epigallocatechin gallate (EGCG) inhibits the oxidation of dG
and 8-oxodG by HOCl; however, the inhibition effect was greater for 8-oxodG (126).
28
Chapter 1 – The Literature
Other biological compounds increase the efficiency of oxidation product formation. At
low concentrations of HOCl (100 uM), Sp forms at levels 6.6 fold higher than 8-Cl-dG,
and micromolar amounts of nicotine increase the yield. Nicotine apparently improves the
reaction yield through the formation of a nicotine-Cl+ adduct that reacts with dG and 8oxodG more efficiently than HOCl. Higher concentrations of nicotine (above 50 uM)
cause an increase in the yield of 8-oxoG at the expense of Sp (125). Burrows and
coworkers recently reported that oxidation of 8-oxoG in the presence of spermine
produces Ua (127).
1.3.3. Type II photosensitized oxidation of guanine and 7,8-dihydro-8-oxoguanine
DNA is susceptible to oxidation by 1O2 (128-136). Photoexcitation of dGMP
with UVB light generates 1O2 by type II photosensitization (energy transfer from a
photoexcited sensitizer to 3O2), and 1O2 then reacts with the nucleobase (137) to form the
5’-monophosphates of Sp and Iz + Z (138, 139).
An earlier study noted that UV
irradiation of the A, T, C, and U nucleobases produces 1O2; however, irradiation of G
(free base), dGMP, and TpG do not cause 1O2 to accumulate (140), suggesting the
reaction of 1O2 with the guanine nucleobase occurs quickly relative to the production of
1
O2. The reaction of 1O2 with G is a [4+2] cycloaddition with the imidazole ring to form
an endoperoxide (106, 141-145), which leads to the formation of 5-HO-8-oxoG (Scheme
1.4) (146). 5-HO-8-OxoG subsequently degrades to Sp or Gh, depending on the reaction
conditions. As an aside, a previous study by Loeb and coworkers showed that methylene
blue and white light treatment (a source of 1O2) of an M13mp2 single-stranded genome
causes the formation of an unidentified DNA lesion that is both a block to replication and
induces 66% GÆC and 27% GÆT mutations (31). Given what is now known about the
mechanisms of G oxidation product formation and the mutational properties of these
DNA lesions (see section 1.6 and Table 1.4), it is likely the DNA damage observed by
Loeb and coworkers is Sp.
Two pathways to 5-HO-8-oxoG from the cycloaddition product have been
proposed. One pathway proposes that two equivalents of 1O2 and two reduction steps
result in the formation of 5-HO-8-oxoG, with no oxygen atom incorporation from H2O
29
Chapter 1 – The Literature
(Scheme 1.4, path A) (146). This mechanism is inconsistent with a prior study where 18O
from H218O was found in the final Sp product. The proposed mechanism in the latter
case involves only 1 equivalent of 1O2 (Scheme 1.4, path B) (111). However, Foote and
coworkers note that in their mechanism (path A) exchange of H2O with the HOO- or HOgroups at C5 would account for the incorporation of oxygen from H2O (146). Oxidation
of dG by 1O2 generates a minor product, DIz, in addition to 8-oxodG, Iz + Z, and Sp. DIz
forms directly from dG, not from 8-oxodG, and hydrolyzes to form Iz + Z (147).
As shown in Scheme 1.4, path B, 1O2 adds to 8-oxoG across the C4 and C5 bond
by [2+2] cycloaddition to form a dioxetane intermediate (148), which opens to form 5hydroperoxy-8-oxo-4,8-dihydroguanine (5-HOO-8-oxoG) and eventually 5-HO-8-oxoG
(146). The structures of 5-HOO-8-oxoG and 5-HO-8-oxoG were initially misassigned as
the respective 4-isomers (148). The distribution of end products from 1O2 oxidation of 8oxoG varies depending on the environment of the 8-oxoG base. At the nucleoside level,
oxidation of the 8-oxoG base by methylene blue photosensitization generates Ca, Iz +Z,
and Sp with yields of 50%, 35%, and <10%, respectively (149, 150), whereas Oa is by far
the major product in single-stranded oligonucleotides (86).
Methylene blue can
participate in type I photosensitization to a small extent (151), so Cadet and coworkers
investigated the oxidation of 8-oxoG by a “clean” source of 1O2 (produced thermally,
rather than photochemically) and found that essentially no Ca forms, suggesting other
methylene blue-related reactions may be involved in the production of Ca (152). Iz and
DGh appear to form from the intermediate 5-HOO-8-oxoG (Scheme 1.4).
The
photosensitized oxidation of 8-oxoG nucleoside by methylene blue in the presence of a
thiol generates Sp as the major product. Presumably, the thiol reduces the 5-HOO-8oxoG intermediate to 5-HO-8-oxoG, thereby allowing Sp to form (107). This result also
suggests that Iz and DGh result from the 5-HOO-8-oxoG intermediate. Two additional
compounds,
6-amino-2,5-diimino-2,5-dihydro-3H-pyrimidin-4-one
(1)
and
1,3,5-
triazepane-2,4,6,7-tetrone (2), result from 1O2 oxidation of 8-oxoG, but to date have only
been observed in organic solvent (Figure 1.1) (144, 148).
30
Chapter 1 – The Literature
1.4. Structural Aspects of Guanine Oxidation Products
Relatively few structural studies have been reported for G oxidation products.
Helix melting temperature (Tm) experiments represent the majority of the investigations.
DNA melting studies indicate duplexes containing Sp or Gh paired opposite A, T, C, or G
are destabilized relative to duplexes containing a G:C pair at the same position.
Additionally, duplexes consisting of one strand that contains an internally placed Sp or
Gh and a complementary strand that occupies the region 3’ of the lesion with no base
opposite the lesion experience a slight thermodynamic destabilization compared to when
a G occupies the site of the lesion.
The degree of thermodynamic destabilization
generally increases as the length of the complementary strand increases (153). In a
14mer, the presence of a Ca:C base pair reduces the melting temperature by 15 °C to 40 ±
1 °C in comparison to an unmodified oligonucleotide containing a G:C pair at the same
location (154). NI paired with A, T, C, or G decreases the melting temperature of a
19mer duplex by 10-12 °C (155).
A computational study of the Sp nucleoside reveals the rings of the diastereomeric
lesions are essentially planar and perpendicular to each other (156). In comparison to G,
the conformational freedom is restricted. The authors also compare the tautomeric forms
that the lesions may adopt. Of the two possible amino forms, one was chosen to compare
with the imino form and is 1 kcal/mol lower in energy (156). Uncertainty remains as to
which of the two amino forms is preferred. Molecular dynamics simulations show that
the Sp diastereomers generally are situated in the major groove of B-DNA and that
intercalation structures severely disrupt duplex structure (157). Differences in duplex
stability were noted for one diastereomer versus the other. For example, the lesions adopt
both syn and anti conformations, but the opposing base and the stereochemistry of the
lesion determine the preferred conformation. The lesions also form hydrogen bonds with
the opposing base, and the R isomer forms hydrogen bonds more favorably than the S
isomer.
31
Chapter 1 – The Literature
Ua exists as a mixture of α- and β-anomers, and an NMR study showed that both
anomers are present in a duplex 12mer. The β-anomer aligns interhelically with the base
stack and shares at least one hydrogen bond with the complementary base T (158).
32
Chapter 1 – The Literature
1.5. Repair of Guanine Oxidation Products
E. coli has three enzymes, MutM, MutY, and MutT, that belong to the GO system
and protect the cell from mutagenesis by 8-oxoG (159, 160). Polymerases incorporate A
opposite 8-oxoG, which leads to GÆT mutations, but also incorporate C opposite the
lesion, which produces no mutation. MutM excises 8-oxoG lesions when they are paired
opposite C,T, and G, but shows weak activity towards 8-oxoG:A pairs (161). In contrast,
MutY removes A when paired opposite 8-oxoG (162). Thus, the excision of A when
paired with 8-oxoG by MutY gives the cellular replication machinery a second
opportunity to insert a C opposite the lesion, which in turn allows the lesion to be excised
by MutM.
Endonuclease III (Endo III or Nth) (163) recognizes many oxidized
pyrimidines (164, 165) and is of interest since most of the G oxidation products lack the
two ring purine structural feature. Recently, endonuclease VIII (Nei) was found to
possess activity for 8-oxoG paired with G, A, or C. Nei is hypothesized to prevent
mutations resulting from 8-oxoG incorporation in the nascent strand opposite A in the
template strand (166).
Several groups have tested 8-oxoG oxidation products as substrates for repair
enzymes. Table 1.2 lists the kinetic properties of E. coli Nei, Nth, and MutM for the
excision of Sp and Gh when paired opposite C, G, and A. Base pairs containing G are
the preferred substrates for Nei, whereas base pairs containing C are preferred by MutM.
Nth shows similar specificity for base pairs containing C or G. Significantly, only Nei
can efficiently excise Sp and Gh when paired opposite A (167). Another study showed
qualitatively that MutM excises Sp and Gh when paired with A, T, C, or G, although the
rate of removal of the lesion opposite A is markedly lower (118). The ability of MutY to
excise A, C, or G when paired with Sp or Gh has been investigated. Although MutY
shows negligible glycosylase activity towards these base pairs, the enzyme does form
stable complexes with the lesion-containing duplex oligonucleotides.
Additionally,
removal of Sp or Gh by MutM is inhibited by 10 nM MutY when the lesions are paired
with G but not when they are paired with C, suggesting the two enzymes interact to
prevent potentially mutagenic repair processes. Nth was inhibited in all cases by MutY,
33
Chapter 1 – The Literature
whereas Nei was inhibited only when a higher concentration of MutY (100 nM) was used
(118, 167).
Oa and Z paired opposite the four natural nucleobases are efficiently removed by
MutM and Nth. Comparison of the catalytic efficiencies for the repair of Oa:C or Z:C
versus 8-oxoG:C reveals that 8-oxoG is a better substrate by 2- and 4-fold, respectively.
In contrast, Oa:C and Z:C are much better substrates for Nth than 5-hydroxycytosine:G
by about 100-fold (168, 169). A separate study showed MutM excises Oa quantitatively,
whereas 70% of Z and less than 10% of Ca are excised when these three lesions are
paired opposite C (170). Gasparutto et al. found that neither MutM nor Nth could excise
Ca from a duplex containing a Ca:C base pair (154). NI is a weak substrate for MutM
and is not a substrate for Nth under the conditions tested (73).
Eukaryotic repair enzymes also show activity towards some of the lesions
discussed in this review. Murine NEIL1 and NEIL2 efficiently excise Sp and Gh from
single-stranded oligonucleotides. Both enzymes also excise Gh from double-stranded
DNA opposite all four native bases with similar efficiency. NEIL1 shows activity for Sp
in duplex DNA; however, NEIL2 shows little activity, but does bind to the Sp-containing
oligonucleotide. The two enzymes show little to no activity for single- or doublestranded substrates containing 8-oxoG (171).
The eukaryotic homologues of MutM, known as OGG, have been screened for
activity towards Sp- and Gh-containing oligonucleotides.
The human homologue,
hOGG1, shows activity only for 8-oxoG, with a 1000-fold preference for 8-oxoG paired
with C over 8-oxoG paired with A. The yeast homologues, yOGG1 and yOGG2, remove
Sp and Gh from duplexes, and the base paired with Sp or Gh does not significantly affect
the rate. Whereas yOGG1 removes Sp and Gh at similar rates, yOGG2 removes Sp more
efficiently than Gh. For comparison, the rate of removal of Sp and Gh by the yOGG
enzymes is about 20-30 fold lower than the removal of 8-oxoG by hOGG1 or MutM.
The ability of the yOGG proteins to excise Sp and Gh regardless of the identity of the
34
Chapter 1 – The Literature
complementary base indicates the action of the yOGG proteins on these lesions is a
mutagenic process except when the complementary base is C (172).
Human methylpurine DNA N-glycosylase (MPG) removes Ca when paired with
any of the natural nucleobases, although Ca:C and Ca:T pairs are removed more
efficiently than Ca:G and Ca:A pairs. Other enzymes tested in this study include MutM,
Nth, AlkA, Ogg1, Ntg1, Ntg2, hOgg1, and hNth1, but these enzymes do not show
activity for Ca-containing duplex substrates. Comparison of the rate of excision of 7methylguanine:C versus Ca:C shows that the removal of 7-methylguanine is only 1.6-fold
higher than that for the removal Ca indicating that Ca:C is a relatively good substrate for
MPG (173).
Incorporation of 8-oxodGMP by DNA polymerases during replication induces
ATÆCG mutations.
The E. coli enzyme MutT hydrolyzes the triphosphate of 8-
oxodGTP with a specificity 1000-fold higher than dGTP in order to prevent the
incorporation of this lesion into the cellular genome (174, 175). A human homolog,
denoted as hMTH1 (human MutT Homolog-1), exists and also possesses 8-oxodGTPase
activity (176, 177). The ability of MutT or its homologs to hydrolyze triphosphates of
the oxidation products discussed in this review has not been reported; however, given that
the DNA repair enzymes discussed above often act on G oxidation products besides 8oxoG, it would not be surprising if the triphosphates of these lesions are also substrates
for MutT, its eukaryotic homologs, or another class of enzymes operating in a similar
fashion to maintain the informational integrity of the nucleotide pool.
35
Chapter 1 – The Literature
1.6. Genotoxic and Mutational Properties
The genotoxic and mutational properties of DNA lesions illustrate potentially
deleterious aspects of lesion formation and therefore are critical metrics. The following
sections review in vitro and in vivo mutagenesis studies involving G oxidation products.
In the in vitro studies, the ability of a polymerase to extend a primer annealed to a
template containing a single lesion at a defined position (known as a site-specifically
modified template) is determined along with the identity of the base inserted opposite the
lesion. These experiments are advantageous because the conditions can be precisely
defined, the kinetic parameters for replication of a lesion by a specific polymerase can be
determined, and the experiments can be completed relatively quickly. While critical for
defining enzymatic properties, in vitro studies do not present a complete analysis of DNA
lesion properties since they neglect the complexity of the cellular environment. In vivo
experiments assess the genotoxic and mutagenic properties of a DNA lesion in the
presence of all replication and repair components, or in the presence of these components
minus specifically deleted or dysfunctional gene products, and thus provide a highly
relevant appraisal of DNA lesion properties. The in vivo experiments discussed below
use a site-specifically modified single-stranded M13 vector that can replicate in E. coli as
a model for in vivo leading strand DNA replication (178). From these studies, the effect
of the E. coli replication and repair machinery on a DNA lesion can be determined.
1.6.1. In vitro properties of guanine oxidation products
Sp and Gh are strong blocks to replication by Klenow fragment (Kf) and Klenow
fragment exo− [Kf (exo−)] (Table 1.3). In the presence of all four deoxynucleotides, the
major product of standing start primer extension in the case of 8-oxoG-containing
template is the full length product, but in the case of Sp- or Gh-containing template, only
trace amounts of full length product are detected. For both Sp- and Gh-containing
templates, the polymerase inserts predominantly A and G opposite the lesions, which
would result in GÆT and GÆC mutations, respectively (117, 153). In a different
sequence context, Sp and Gh are complete blocks to full length primer extension by Kf
(exo−), and the polymerase inserts almost entirely A opposite the lesions (123). Under
running start conditions where the primer must be extended by two bases prior to
36
Chapter 1 – The Literature
insertion opposite the lesion, similar results are obtained as under standing start
conditions with the exception of Gh, which is bypassed with much higher efficiency
when the proceeding two bases in the template are G rather than A. This result suggests
that the ability of Kf to bypass Gh is dependent on the stability of the duplex region
upstream of the lesion (153).
Replication of Iz using DNA polymerase I (pol I) or the Kf shows that the lesion
codes exclusively as a C, but it is inefficiently bypassed (62). Ab initio studies of Iz in
vacuo and using a self-consistent reaction field method of solvation show that the lesion
mimics the hydrogen bonding face of C, thereby providing a rationale for the GÆC
mutations induced by this lesion (62, 63). Ca does not block replication when Kf (exo−)
is used. The polymerase inserts A most readily opposite Ca, but also G to a much smaller
extent (154). Primer extension using Kf (exo−) reveals that A is incorporated opposite Oa
and that full length extension results.
When Taq DNA polymerase is used as the
replicating polymerase, both A and G are inserted opposite the lesion, although Oa blocks
synthesis to some degree. Oa inhibits replication by DNA polymerase β (pol β) in the
presence of individual nucleotide triphosphates and all four triphosphates (168).
In a related study, Kf (exo−) inserts A opposite Z, and in the presence of all four
nucleotide triphosphates, a small amount of full length product is observed. Z blocks
replication by pol β, whereas Taq incorporates A and, to a smaller extent, C and G
opposite the lesion; however, Taq does not fully extend the primer (169). Ua is a block
to replication (179, 180).
Kf (exo−), pol β, and T7 DNA polymerase can insert a
nucleotide before and opposite Ua, but Ua inhibits full length extension of the primer.
Kinetic analysis of insertion by Kf (exo−) reveals A and G are incorporated opposite the
lesion (179).
The bypass and coding properties of DNA containing NI were previously studied
using the purified DNA polymerases Kf (exo−), calf thymus polymerase α (pol α), and
human pol β (73). All three polymerases could insert a nucleotide opposite the lesion
and, although a kinetic analysis was not performed, the data suggest that extension of the
37
Chapter 1 – The Literature
NI-containing base pair by Kf (exo−) and pol α is rate limiting and results in poor
translesion synthesis. The nucleotides A and G are incorporated opposite NI by Kf (exo−)
and pol α, with Kf (exo−) favoring insertion of C opposite the lesion. Interestingly, pol β
efficiently bypasses the lesion and incorporates mostly C, rendering the lesion non-toxic
and non-mutagenic.
The mutational properties of 8-NO2-G were investigated using mammalian DNA
polymerases (76). Running start primer extension reactions in the presence of all four
dNTPs show that 8-NO2-G inhibits DNA synthesis by pol α and pol β such that
nucleotides were incorporated one base prior to the lesion and opposite the lesion, but
negligible amounts of full length product result. Primarily, C is incorporated opposite the
lesion resulting in no mutation, but to a lesser extent A and T are also inserted. Under
similar conditions, pol η and pol κ∆C fully extend the primer and experience only minor
blocking by the lesion. Pol η inserts similar amounts of C and A preferentially and also
inserts G and T. Pol κ∆C mainly inserts A, but also inserts C, G, and T opposite the
lesion.
1.6.2. In vivo properties of guanine oxidation products
The in vivo DNA polymerase bypass efficiencies of the oxidation products
relative to G are given in Table 1.4. In general, the lesions can be separated into two
groups based on the bypass efficiency. The first group consists of Oa, Ca, Iz, Z, and Gh,
all of which are either well-bypassed or moderately well-bypassed. The second group
includes Ua, Sp, and NI, which are poorly bypassed (92, 181-184). For comparison, the
bypass efficiency of 8-oxoG in the same system is 85-90% (92, 181, 182). The bypass
efficiencies of Ca, Oa, Ua, Iz, Z, and NI were also determined in SOS-induced E. coli.
SOS induction upregulates the expression of more than 40 proteins in E. coli (185)
including three DNA polymerases, pol II (186), pol IV (187), and pol V (188), which are
known as translesion synthesis polymerases because of their ability to bypass DNA
lesions (189). The bypass efficiency of NI increases 8-fold under these conditions,
whereas no significant increase in the bypass efficiency occurs for the other 5 lesions (92,
181, 183, 184).
38
Chapter 1 – The Literature
The mutational properties for these lesions in single-stranded DNA as reported by
Essigmann and coworkers are presented in Table 1.4 (92, 181-183). In comparison to 8oxoG, which has a mutation frequency of about 3-7% in the same system, all of the
lesions studied are highly mutagenic, suggesting their formation in vivo could be
deleterious in the absence of effective repair mechanisms. SOS induction does not
significantly affect the mutational signature of Ca, Oa, or Z (92, 181) but does affect the
properties of Ua, Iz, and NI (92, 183). It is noted that repair systems that operate only on
lesions in double-stranded DNA could represent defenses against these lesions. As
indicated above, the replication properties of the lesions were evaluated in singlestranded genomes.
Interestingly, similar lesion structures do not infer similar in vivo bypass
efficiencies or mutational signatures. For example, Gh and NI share much structural
similarity but Gh is well-bypassed and codes almost exclusively as a C, whereas NI is
poorly bypassed and codes mostly as G but also as A, T, and C. Iz and Z also are
structurally similar, but Iz codes mostly as a C, whereas Z codes as an A. The mutational
signatures of the Sp diastereomers remain qualitatively similar but differ significantly.
These results clearly demonstrate that the identity of the lesion influences the rate of
polymerase bypass and the choice of dNTP inserted opposite the lesion and suggest that
in spite of the structural similarity the interactions between the lesion and the polymerase
active site differ enough to cause significant shifts in mutational signature. Structural
studies on polymerases in primer-template complexes with DNA lesions are needed to
help develop an understanding of the aforementioned important issues.
Comparison of the bacterial in vivo mutagenesis data to the in vitro mutagenesis
data with Kf indicates the two do not always correlate. This observation is not surprising
given that in vitro conditions vary dramatically from in vivo conditions and that E. coli
possesses four other known DNA polymerases besides pol I (Klenow fragment). The
results for Ua, Iz, and NI from SOS-induced E. coli strongly suggest involvement of at
least one of the SOS-inducible DNA polymerases, namely pol II, pol IV, and pol V. The
39
Chapter 1 – The Literature
differences in mutagenesis data from normal cells and SOS-induced cells show that
polymerases in SOS-induced cells do not necessarily insert dNTPs in the same ratio as do
the polymerases in normal cells.
Many eukaryotic DNA polymerases have been identified (190) and five (pol η,
pol ι, pol κ, pol ζ, and Rev1) are known to be involved in translesion DNA synthesis
(TLS) (191). Although some of the in vitro studies discussed earlier report findings for
eukaryotic polymerases, only one of these studies involves eukaryotic TLS polymerases
and reports quantitative enzymatic data (76).
Thus, numerous questions remain
concerning the eukaryotic mechanisms for bypassing guanine oxidation products.
40
Chapter 1 – The Literature
1.7. Biological Relevance
Of the lesions reviewed here, only 8-oxoG, 8-NO2-G and Sp have been detected
in vivo. Numerous studies report the detection of 8-oxoG in human cells at levels of 1100 lesions per 106 bases (192). However, given the abundance of 8-oxoG in cellular
DNA and its ease of oxidation relative to G, the in vivo existence of many of the lesions
discussed in this review is plausible. An important recent discovery is the in vivo
detection of Sp lesions by Sugden and coworkers (193). Previous studies showed that Sp
forms in DNA treated with Cr(VI) (potassium dichromate) (120) and that the BER
enzyme Nei excises Sp lesions (167). The authors treated Nei deficient E. coli with
Cr(VI), analyzed for Sp in the genomic DNA using mass spectrometry, and found the
amount of Sp is ~20-fold greater than in wild-type cells when treated with 500 µM
Cr(VI). The Nei deficient strain does not accumulate 8-oxoG lesions suggesting that
Cr(VI) oxidation competes with the MutM/MutY repair system. Essentially no increase
in the amount of Sp is observed for cells that are Nei proficient, including cells that are
also MutM/MutY deficient (193).
The detection of 8-NO2-G is useful as a biomarker of inflammation (194) and
forms as a result of a variety of conditions such as chronic hepatitis C infection (195),
intra-hepatic cholangiocarcinoma (196), Helicobacter pylori infection (197), and
inflammatory bowel disease (198). In vitro, 8-NO2-G and NI form in double-stranded
DNA in approximately equal proportions (74), so it seems reasonable that this lesion
should form under the same conditions that generate 8-NO2-G in vivo. Since NI is a
chemically stable lesion, unlike 8-NO2-G, this toxic and mutagenic lesion may
accumulate in cells in the absence of an efficient repair system.
The potential discovery of very low amounts of each lesion in the cell relative to
8-oxoG could argue against their biological relevance. However, since the mutation
frequency of many of these lesions is at least an order of magnitude higher than that for
8-oxoG, the amount of a well-bypassed lesion in the cell could be an order of magnitude
lower than the amount of 8-oxoG and still induce a similar number of mutations. As
suggested by the discovery of Sp in repair deficient bacterial cells, an efficient repair
41
Chapter 1 – The Literature
system may have evolved to combat the potent mutagenicity of these DNA lesions.
Consequently, discovery of more G oxidation products in cells may be contingent on
elimination of the specific DNA repair activities for the lesion in question. As yet
undiscovered repair activities may also preclude detection of other G oxidation products.
42
Chapter 1 – The Literature
1.8. Concluding Remarks
Oxidation of G yields a diverse array of products, and many questions remain
about the properties of these lesions.
Recent studies have provided a much better
understanding of the mechanisms of formation for these compounds, but additional
studies are needed to explain numerous observations, such as the dependence of the ratio
of Sp to Gh formation on the presence or absence of sugar protecting groups. Detailed
experimental structural studies of the lesions in DNA and their interactions with repair
proteins and DNA polymerases are also needed. Of note, NI, Ca, Sp, and Gh possess the
stability necessary for NMR and X-ray structure studies, and phosphoramidite building
blocks for automated DNA synthesis have been reported for NI (155) and Ca (154).
Many in vitro studies demonstrate the ability of various repair enzymes and polymerases
to act on G oxidation products. Further in vitro studies with TLS polymerases are
essential since these polymerases likely have a major and critical role in the processing of
the lesions. Studies in both prokaryotic and eukaryotic cells are needed to ascertain the
true level of involvement of specific repair proteins and DNA polymerases on the
oxidation products.
Detection of the G oxidation products in vivo must occur to
demonstrate the biological relevance of this chemistry. The recent discovery of Sp
lesions in vivo should encourage further investigations in this area.
43
Chapter 1 – The Literature
1.9. Acknowledgment
We thank the National Institutes of Health (CA26731 and CA080024) for
financial support.
44
Chapter 1 – The Literature
Figure 1.1. Guanine oxidation products discussed in Chapter 1.a
O
O
N
HN
N
H2N
G
N
N
R
H2N
O
O
NH2
N
HN
NH
N
R O
N
N
R
N
HN
R
N
Dz
H
N
O
N
R
O
O
N
R O
NH
O
N
HN
R
N
NH2
Iz
H
N
O
NH
NH2
N
HN
NO2
O2N
O
NH2
HN
R
N
H
O
O
N
O H2N
NH
R
N
N
H H
R
H2N
N
O
N
R
HN
N
R
O
HN
O
N
R
Ca
O
NH
HN
NH2
O
N
R
2
O
NH
O
O
1
NH
The names of these compounds are listed in Table 1.1.
45
HICA
NH
HN
NH2
O
N
R
O
O
CAC
Oz
NH
R
O
O
OHH
N
HO
Ua
Ia
N NH2
N
R
O
H2N
NH O
O NH2
N
H
NI
O
Oa
Gh
O
N
H2N
N
R
N
O
HO
O
N
N
H H R
H2N
N
H
O2N
NH
N2-NO2-G
Pa
Sp (S)
HN
O
DGh
O
N
R
N
N
HN
8-NO2-G
H
N
H2N
Sp (R)
a
H2N
O
NH
NO2-DGh
O
O
8-oxoG
N
R
N
N
HN
N
R
N
H
N
O
NH2
O2N
HN
O
O
H
N
HN
H2N
OH
N
N
N
ONO R
nox-G
O
Chapter 1 – The Literature
Figure 1.2. Structure of DGh and other proposed precursors of Oa.
O
NH
H2N
N
DGh
H
N
O
N
R
N
H2N
N
H
N
O
O
NH
R
H2N
3
N
O
N
N
R
4
46
NH2
O
H
N
N
O
HN
N
N
H OH
R
5
Chapter 1 – The Literature
Scheme 1.1. Proposed mechanisms for formation of major oxidation products directly from G by one electron processes [adapted
from Refs (55, 67, 113)].
O2N
NH
N
N
H
N
R
H2N
O NO
2
N
N
H2N
N
HN
H2N
O
N
G
N
R
[O]
H2N
N
G
N
R
+
H
N
N
G( H)
H2N
N
H2N
N
H2N
+ O2
N
R
+H HN
2
N
N
N
R
H2N
+ H2O
H2N
HCONH2
N
R
N
N
+
N
+ CO3
NO2
H2N
H
H2N
47
N
NH
R
+ H2O
N
N
R
NO2
8-NO2-G
N
R
OH
O
O
[H]
H
H
N
R
HN
H2N
O
H
O
O
H
N
N
N
R
8-oxoG
H2N
O
O
H2N
N
NH
R
Z
O
HCO3
O
N
O
Iz
N
HN
N
HN
N
O
O
N
R
O
NH
[O] (?)
N
N
N
N
R
O
O
HN
CO2
N
+ NO2
O
N
OH
N
N
N
R
O
O
N
N
H2N
H2N
+ O2
+
+H
O
N
HN
N
N
R
+ NO2
NI
O
N
O
Chapter 1 – The Literature
Scheme 1.2. Proposed mechanisms for formation of major oxidation products directly from 8-oxoG by one electron processes
[adapted from Refs (72, 92, 101, 113)].
O
HN
H2N
N
O OH
O H
H
H
+
N H
N
HN
H
O
N
N
H2N
N
R
R
O
O H
H
N H
O
N
R
O
HN
H2N
O
H
N
O
N
N
R
[O]
HN
H2N
8-oxoG
N
R
Z
Iz
pH > 7
H2N
N
N
OH
O H
N
N
R
N
H2N
H
N
O
H
N
R
CO2
O
_7
pH <
5-HOO-8-oxoG
H2N
N
H2N
Ua
O
N H
R O
48
NH
R
2+
H
N
DGh
O
O
Mg + HCO3
(catalysts)
oxalate
+ O2
+ H+
O
NH
H
N
Gh
H
O
N
O
N
N
H H R
8-oxoG( H)
8-oxoG
O
O
N
O R
O
NH
O
O
NH
Sp
H2N
+ CO3
N
R
N
N
HN
HCO3
O O
H
N
N
H2N
+ O2
+ H2O
H+
NO2
H+
+ NO2
O
H
N
O
N
N
O
N
R
N
O
H2N
5-HO-8-oxoG
NO2
N
H2N
e
H+ H N
2
O
O OH
H
N
N
H
O
H
N
O
N
R
guanidine
Pa
O + H2O
O
HN
R
O
N
H
Oa
OH
O
Chapter 1 – The Literature
Scheme 1.3. Proposed mechanisms for formation of major products of 8-oxoG oxidation
by ONOO− [adapted from Refs (98, 114)].
O
[O]
8-oxoG
8-oxoG
[O]
O
N
N
H2N
O
N
R
N
+ ONOO N
+
+H HN N
2
Gh
H2N
N
O
H2N
O
N
H2N
N
5-HO-8-oxoG
N
Sp
O
+ H2O
guanidine
O
NH
O
N
R
N
R
Ca
NH
OH
N
H2N
N
R
H2N
O
H
N
N
N
R
O
N
O
O
N
R
H2N
HICA
N
O
O
H
N
N
N
R
N
low [ONOO ] products
[H]
O
NH
H
N
N
N
R
O
DGh
Pa
Oa
high [ONOO ] products
49
O
ONOO
H2N
O
NO2-DGh
NH
CAC R
+ NO2
O2N
O
+ H2O
urea
ONOO
CO/CO2
O
O OH
H
N
HO
HN
O
O
+ H2O
NH
O OH
H
N
N
H
N
O
R
O
N
R
N
N
N
H OHR
H2N
H2N
+ H2O
N
R
O OH
H
N
N
O OH
H
N
N
O
NO2
H+
H2O
HO OH OH
H
N
N
ONO
OH
N
Ua
Chapter 1 – The Literature
Scheme 1.4. 1O2 oxidation of G and 8-oxoG [adapted from Refs (111, 146)].
O
HO
O
N
N
R
NH
N
[H]
H
N
N
R
NH2
O2
G
[4+2]
H
O N
N
R O
O2
NH2 [2+2]
N
O
HO
N
O
N
R
O
N
N
H
O
NH
NH2
B
HO
O
N
N
R
N
N
H
N
N
O
N
R
NH2
N
_7
pH <
NH2
N
O
NH2
HO O
H
N
N
N
R
N
NH2
5-HO-8-oxoG
50
pH >
7
DGh
Iz
5-HOO-8-oxoG
[H] or H2O
H2O O
O
HOO O
H
N
N
N
R
NH2
8-oxoG
O
N
1
NH
O
A
1
O
5.8
pH >
Sp
pH <
5.8
Gh
Chapter 1 – The Literature
Table 1.1. Common names that appear in the literature for G oxidation products.
abbr.
8-oxoG
8-NO2-G
N2-NO2-G
NI
NO2-DGh
DGh
Pa
Oa
Ua
HICA
Sp1 + Sp2
Gh
Ia
CAC
Ca
DIz
Iz
Z
nox-G
1
2
common names
7,8-dihydro-8-oxoguanine
8-nitroguanine
N2-nitroguanine
5-guanidino-4-nitroimidazole; nitroimidazole
N-NO2-N'-(2,4-dioxo-imidazolidin-5-ylidene)-guanidine
dehydroguanidinohydantoin
parabanic acid
oxaluric acid
urea
4-hydroxy-2,5-dioxo-imidazolidine-4-carboxylic acid
spiroiminodihydantoin; 2-imino-5,5’-spirodihydantoin
guanidinohydantoin; 5-guanidinohydantoin
iminoallantoin
2,4,6-trioxo[1,3,5]triazinane-1-carboxamidine
cyanuric acid
diiminoimidazolone
imidazolone; 2-aminoimidazolone; 2,5-diaminoimidazolone
oxazolone; 2,2,4-triamino-2H-oxazol-5-one
4,5-dihydro-5-hydroxy-4-(nitrosooxy)-guanine
6-amino-2,5-diimino-2,5-dihydro-3H-pyrimidin-4-one
[1,3,5]triazepane-2,4,6,7-tetraone
51
Chapter 1 – The Literature
Table 1.2. Kinetic parameters for the excision of Gh and Sp in duplex oligonucleotides by Nei, Nth, and MutM (adapted from Ref
(167)).
Nei
a
(nM).
b
(min-1).
substrate
Kma
kcatb
kcat/Kmc
Sp⋅C
Sp⋅G
Sp⋅A
Gh⋅C
Gh⋅G
Gh⋅A
30 ± 4.0
38 ± 5.2
40 ± 6.0
40 ± 5.8
35 ± 3.8
45 ± 5.0
1.6
5.8
2.5
1.0
6.1
3.0
0.05
0.15
0.06
0.03
0.17
0.07
c
kinetic parameters
Nth
a
Km
kcatb
kcat/Kmc
32 ± 4.5 0.50
0.015
34 ± 5.0 1.14
0.030
N.D.
39 ± 3.8 0.39
0.010
42 ± 4.8 0.50
0.012
N.D.
(min-1 nM-1). N.D., not determined.
52
MutM
Kma
25 ± 3.0
75 ± 8.0
22 ± 2.5
65 ± 7.0
kcatb
6.0
1.5
N.D.
5.9
1.7
N.D.
kcat/Kmc
0.24
0.02
0.26
0.03
Chapter 1 – The Literature
Table 1.3. Summary of in vitro standing start primer extension results.
guanine oxidation
d
Sp
Gh
Iz
Ca
Oa
Z
pola
Kf
Kf
Kf
Kf
Kf
Taq
pol β
Kf
Taq
Ua
pol β
Kf
NI
pol β
T7 pol
Kf
8-NO2-G
pol α
pol β
pol α
pol β
pol κ∆C
pol η
opposing
b G
A,
A, G
C
A>G
A
A, G
A
A>C,G
none detected
A, G
C>A, G
A, G
C
C>A, T
C>A, T
A>C,G, T
C, A>G, T
53
insertion
ffi +i
+
+
+
+
extension
ffi −i
−
−
+
+
−
−
−
−
−
−
−
−
−
−
+
−
+
+
+
+
+
+
+
+
+
+
−
−
+
+
−
−
+
+
references
(117, 123, 153)
(117, 123, 153)
(62)
(154)
(168)
(168)
(168)
(169)
(169)
(169)
(179)
(179)
(179)
(73)
(73)
(73)
(76)
(76)
(76)
(76)
Chapter 1 – The Literature
Table 1.4. Summary of bypass efficiencies and mutation frequencies for 8-oxoG oxidation products in wild-type E. coli (%).
SOS
−
−
−
−
+
+
+
+
a
Ref (181).
b
property
bypass
GÆTf
GÆC
GÆA
bypass
GÆT
GÆC
GÆA
Ref (183).
c
8-oxoGd
~88
7
<1
<1
N.S.D.
6.8
1.3
N.S.D.
Ref (182).
eluting isomer, see Ref (182).
f
Ca a
65 ± 8
97
<1
<1
N.S.D.
N.S.D.
N.S.D.
N.S.D.
d
NIb
7.0 ± 1.6
22
8.9
19
57 ± 1
N.S.D.
2.5
13
Ref (92).
e
guanine oxidation product
Izb
Za
Gh c
Sp1 c,e
60 ± 5 57 ± 13 75 ± 5
9±3
1
86
1.4
27
88
<1
98
72
2
<1
<1
<1
N.S.D.
N.S.D.
N.D.
N.D.
5.5
N.S.D.
N.D.
N.D.
75
N.S.D.
N.D.
N.D.
3.4
N.S.D.
N.D.
N.D.
Sp2 c,e
9±4
41
57
<1
N.D.
N.D.
N.D.
N.D.
Oa d
51 ± 3
99
<1
<1
N.S.D.
95
3.8
<1
Ua d
11 ± 5
99
<1
<1
N.S.D.
43
46
10
Sp1 refers to the isomer eluting faster by HPLC, and Sp2 refers to the slower
GÆT implies a G∗:A initial pairing, which would resolve to a G:CÆT:A mutation upon further
replication (G∗ = guanine oxidation product under study). N.S.D., no statistical difference. N.D., not determined.
54
Chapter 1 – The Literature
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Chapter 1 – The Literature
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Mitra, S. (2001) Repair of hydantoins, one electron oxidation product of
8-oxoguanine, by DNA glycosylases of Escherichia coli. Nucleic Acids
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(168) Duarte, V., Gasparutto, D., Jaquinod, M., Ravanat, J., and Cadet, J. (2001)
Repair and mutagenic potential of oxaluric acid, a major product of
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synthesis opposite oxazolone and repair of this DNA damage using
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Peroxynitrite-induced secondary oxidative lesions at guanine
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(171) Hailer, M. K., Slade, P. G., Martin, B. D., Rosenquist, T. A., and Sugden, K. D.
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acid, a stable and mutagenic oxidation product of 8-oxo-7,8dihydroguanine. Int. J. Radiat. Biol. 80, 21-27.
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73
CHAPTER 2
Efficient Synthesis of DNA Containing the Guanine
Oxidation-Nitration Product 5-Guanidino-4-nitroimidazole:
Generation by a Postsynthetic Substitution Reaction
74
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
2.1. Abstract
A convertible nucleoside was synthesized and used to prepare the 2’deoxynucleoside of 5-guanidino-4-nitroimidazole (NI), a putative in vivo product of the
reaction of peroxynitrite with guanine (G). The convertible nucleoside was incorporated
into an oligodeoxynucleotide (ODN) by the phosphoramidite method and converted postsynthetically to yield an ODN containing the NI lesion site-specifically. A melting
temperature study revealed that duplexes containing NI are greatly destabilized relative to
duplexes containing G at the same site.
75
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
2.2. Introduction
Of the four nucleobases of DNA, guanine has the lowest oxidation potential, and
thus considerable attention has been given to the study of the oxidation of this base (1).
The DNA lesions 7,8-dihydro-8-oxoguanine (8-oxoG) and 8-nitroguanine (8-nitroG) are
two major guanine (G) oxidation products that are formed by reaction with the
endogenous oxidant peroxynitrite (ONOO−) (2).
These lesions are also labile to
oxidation and form a variety of secondary oxidation products (3-5). These products may
result from in vivo oxidation by oxidants other than ONOO− (1, 6), and many are
susceptible to hydrolysis or further oxidation. In contrast, 5-guanidino-4-nitroimidazole
(NI) forms directly from the oxidation of G by ONOO− and is both chemically stable and
resistant to tested forms of enzymatic repair in vitro (7-9). Additionally, this lesion may
be well-bypassed and mutagenic, as suggested by in vitro primer extension assays (7). NI
therefore may contribute to the mutagenic spectrum of DNA damage induced by ONOO−,
and its measurement may be useful as a specific biomarker of ONOO−-mediated
oxidation.
Evaluation of the chemical and biological properties of oxidative DNA lesions is
facilitated by the availability of oligodeoxynucleotides (ODN) and, eventually, genomes
containing these lesions site-specifically (1). Most commonly, oxidative lesions are
prepared site-specifically by damaging a specific base within an ODN (10-12). This
approach facilitates and accelerates the initial characterization of these lesions while
circumventing the common problem of lesion instability, a major obstacle to
incorporation of these damaged bases by automated DNA synthesis. However, low
yields and the formation of multiple products that are often difficult to separate limit the
use of ODN sequences to those containing only one G residue. The present methods for
preparing NI-containing ODNs are hindered by both of these characteristics (7, 8). In
advance of biological evaluation of NI mutagenicity and genotoxicity, we sought to
develop a synthetic method for the preparation of NI lesions site-specifically within
ODNs that is high-yielding and independent of base sequence.
76
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
A convertible nucleoside phosphoramidite offers a convenient route to
incorporating an unnatural nucleoside at a defined site within an ODN (13).
This
approach has been used successfully to prepare ODNs containing nucleosides that are
unstable to DNA synthesis conditions (14, 15), containing structural analogs of a given
base, and with additional functionality (16-22). In the present case, the use of the
convertible nucleoside approach would avoid potential protection-deprotection schemes
with the guanidino functionality of NI, enabling an efficient synthesis of a
phosphoramidite building block. Herein, we describe the synthesis of a convertible
nucleoside phosphoramidite for the site-specific incorporation of an NI lesion into an
ODN by postsynthetic substitution and the effect of this lesion on duplex stability.
Although we only present the synthesis of the biologically relevant NI lesion, the method
allows access to other potentially useful alternative structures.
77
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
2.3. Experimental Procedures
Materials. Solvents were purchased anhydrous from Sigma-Aldrich, except for THF,
which was distilled from CaH2 prior to use. Reagents were purchased and used without
further purification from Sigma-Aldrich, except for 3,5-di-O-toluoyl-α-1-chloro-2deoxy-D-ribofuranose, which was purchased from Chemgenes Corporation. 5(4)-Bromo4(5)-nitro-1H-imidazole (2) (NMR spectra are shown in Figures 2.1 and 2.2) was
prepared as previously described from 5(4)-bromo-1H-imidazole (1) (Sigma-Aldrich)
(23). All reactions were conducted under argon atmosphere. DNA synthesis reagents
and standard phosphoramidites were purchased from Glen Research. Unmodified ODNs
were purchased from IDT, Inc. and HPLC purified prior to use. SVPD was purchased
from ICN Biomedical. Nuclease P1 and alkaline phosphatase were purchased from
Roche Applied Science.
1
H NMR (500 MHz) and
Varian Unity Inova spectrometer.
13
C NMR (125.8 MHz) spectra were obtained on a
31
P NMR (121.5 MHz) spectra were obtained on a
Varian Mercury spectrometer and referenced to 85% H3PO4. HRMS was performed by
the MIT Department of Chemistry Instrumentation Facility. ESI-MS was performed by
the MIT Center for Cancer Research and Howard Hughes Medical Institute Biopolymers
Laboratory. MALDI-TOF MS were acquired on a PerSeptive Biosystems Voyager-DE
STR spectrometer (337 nm laser). Absorption spectra and Tm curves were obtained on a
Varian Cary 100 or 300 Bio UV-vis spectrophotometer equipped with a temperature
controller unit.
5-Bromo-4-nitroimidazole-3’,5’-di-O-toluoyl-β-2’-deoxy-D-ribofuranose (3) and 4Bromo-5-nitroimidazole-3’,5’-di-O-toluoyl-β-2’-deoxy-D-ribofuranose (3A)
Compound 2 (0.500 g, 2.60 mmol) was treated with NaH (60% dispersion in mineral oil,
0.115 g, 2.87 mmol) in 15 mL CH3CN for 25 min with stirring. 3,5-Di-O-toluoyl-α-1chloro-2-deoxy-D-ribofuranose (1.11 g, 2.87 mmol) was added in portions and the
reaction stirred for 3.5 h. The mixture was then diluted with 100 mL of CH2Cl2 and
washed with 100 mL of H2O. The aqueous layer was back extracted twice with 50 mL of
78
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
CH2Cl2 and the combined organic layers were dried over Na2SO4, evaporated at room
temperature on a rotary evaporator, suspended in a minimal amount of 30% hexanes in
CH2Cl2, and loaded onto a silica column prepared in 30% hexanes in CH2Cl2. The first
isomer was eluted with a step gradient of 17.5-0% hexanes:CH2Cl2 and the second isomer
was eluted with a step gradient of 0-4% EtOAc:CH2Cl2. Fractions were pooled and
concentrated to yield 3A (0.499 g, 35%) as a white solid and 3 (0.716 g, 50%) as a white
foam. Isomer 3A: Rf = 0.48 (1:2 EtOAc:hexanes). 1H NMR (CD2Cl2, 500 MHz) (Figure
2.3): δ = 8.05 (1H, d, J = 0.8 Hz), 7.96 (2H, m), 7.81 (2H, m), 7.28 (4H, m), 6.68 (1H, t,
J = 6.2 Hz), 5.60 (1H, dt, J = 3.1, 6.2 Hz), 4.76-4.66 (3H, m), 3.10 (1H, ddd, J = 3.1, 6.0,
14.6 Hz), 2.56 (1H, dt, J = 6.6, 14.6 Hz), 2.43 (3H , s), 2.41 (3H, s); 13C NMR (CD2Cl2,
125.8 MHz) (Figure 2.4): δ = 166.5, 166.3, 145.3, 145.2, 137.2, 130.25, 129.98, 129.91,
129.82, 126.93, 126.86, 122.3, 90.9, 84.7, 74.5, 64.2, 41.3, 22.02, 21.98; UV-vis (EtOH)
λmax (Figure 2.5):
240, 315 nm; ESI-HRMS calc’d for C24H22N3O7Br [(M+Na+)]
566.0533, found 566.0520.
Isomer 3:
Rf = 0.30 (1:2 EtOAc:hexanes).
1
H NMR
(CD2Cl2, 500 MHz) (Figure 2.6): δ = 7.96 (2H, d, J = 8.2 Hz), 7.88 (1H, s), 7.85 (2H, d,
J = 7.9 Hz), 7.28 (4H, dd, J = 7.9, 21.1 Hz), 6.25 (1H, dd, J = 6.1, 7.6 Hz), 5.68 (1H, m),
4.74-4.63 (3H, m), 2.96 (1H, ddd, J = 2.2, 5.9, 14.3 Hz), 2.61 (1H, m), 2.43 (3H, s), 2.41
(3H, s);
13
C NMR (CD2Cl2, 125.8 MHz) (Figure 2.7): δ = 166.5, 166.3, 145.4, 145.2,
134.1, 130.2, 129.95, 129.92, 129.83, 127.0, 126.8, 103.6, 88.4, 84.3, 75.0, 64.2, 40.1,
22.02, 21.96; UV-vis (EtOH) λmax (Figure 2.8): 241, 296 nm; ESI-HRMS calc’d for
C24H22N3O7Br [(M+Na+)] 566.0533, found 566.0552.
5-Bromo-4-nitro-β-2’-deoxy-D-ribofuranose (4)
Guanidine hydrochloride (0.175 g, 1.83 mmol) was treated with 0.5 M NaOMe in MeOH
(3.53 mL, 1.76 mmol) in 36 mL MeOH and the solution stirred for 15 min in an ice bath.
3 (0.400 g, 0.735 mmol) was added and the suspension stirred in an ice bath for 3.75 h.
The reaction was diluted with CH2Cl2 to form a 20% solution of MeOH in CH2Cl2 and
then filtered through a bed of silica. The silica was washed with 100 mL of 20% MeOH
in CH2Cl2, and the combined solutions evaporated. The mixture was adsorbed on to
silica (1.5 g) by suspending in MeOH (5 mL) and then evaporating to dryness. The dry
silica was suspended in CH2Cl2 and applied to a silica column prepared in CH2Cl2.
79
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Elution with a step gradient of 3-5% MeOH in CH2Cl2 afforded 4 as a white foam (0.204
g, 90%). Rf = 0.28 (10% MeOH:CH2Cl2). 1H NMR (CD3OD, 500 MHz) (Figure 2.9): δ
= 8.34 (1H, s, imidazolic H), 6.22 (1H, t, J = 6.1 Hz, 1’H), 4.50 (1H, m, 3’H), 4.03 (1H,
ddd, J = 3.6, 3.6, 7.2 Hz, 4’H), 3.80 (1H, dd, J = 3.2, 12.1 Hz, 5’H) , 3.73 (1H, dd, J =
3.7, 12.2 Hz, 5’H), 2.54 (2H, m, 2’H); 13C NMR (CD3OD, 125.8 MHz) (Figure 2.10): δ
= 136.6, 105.4, 90.0, 89.2, 71.7, 62.4, 42.5; gCOSY NMR (CD3OD, 500 MHz) (Figure
2.11); ESI-HRMS calc’d for C8H10N3O5Br [(M+Na+)] 329.9696, found 329.9698.
5-Guanidino-4-nitroimidazole-β-2’-deoxy-D-ribofuranose (5)
Guanidine hydrochloride (77.4 mg, 0.810 mmol) was dissolved in 1.62 mL of 0.5 M
NaOMe in MeOH. The solution was stirred, evaporated to dryness, and redissolved in 5
mL of absolute EtOH. To this solution was added 50.0 mg (0.162 mmol) of 4, and the
reaction refluxed for 2 h. The solvent was subsequently evaporated and the residue
adsorbed onto silica (0.5 g) by suspending in MeOH and then evaporating to dryness.
The dried silica was suspended in CHCl3 and charged to a silica column prepared in
CHCl3.
Elution with 15-30% MeOH in CHCl3 gave 5 as a bright yellow, very
hygroscopic solid (45 mg, 98%, by UV-vis spectrophotometry).
Rf = 0.24 (30%
MeOH:CH2Cl2). 1H NMR (D2O, 500 MHz) (Figure 2.12): δ = 7.73 (1H, s), 6.08 (1H, t,
J = 6.7 Hz), 4.52 (1H, dt, J = 4.0, 6.3 Hz), 4.03 (1H, m), 3.77 (1H, dd, J = 3.7, 12.5 Hz),
3.69 (1H, dd, J = 5.2, 12.5 Hz), 2.60 (1H, m), 2.46 (1H, ddd, J = 4.3, 6.6, 14.1 Hz); 13C
NMR (CD3OD, 125.8 MHz) (Figure 2.13): δ = 160.0, 143.4, 135.0, 132.2, 89.1, 85.5,
72.4, 63.1, 41.8; UV-vis (H2O, pH 7) λmax (log ε) (Figure 2.14): 230 nm (4.13), 380 nm
(3.82); ESI-HRMS calc’d for C9H14N6O5 [(M+Na+)] 309.0918, found 309.0917.
5’-O-(4,4’-Dimethoxytrityl)-5-bromo-4-nitroimidazole-β-2’-deoxy-D-ribofuranose
(6)
Compound 4 (105 mg, 0.341 mmol) was dried by coevaporation twice with 2 mL of
pyridine, once with 2 mL of toluene, and then by high-vacuum overnight.
4,4’-
Dimethoxytritylchloride (DMT-Cl) (0.260 g, 0.767 mmol), 4-dimethylaminopyridine
(DMAP) (4.2 mg, 10 mol %), and triethylamine (0.107 mL, 0.767 mmol) were combined
80
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
and dissolved in pyridine (3.4 mL). The solution was stirred for 5 min and then added to
a flask containing 4. The reaction was stirred for 3.5 h and then evaporated to dryness.
The mixture was dissolved in CH2Cl2 and applied to a column of neutral alumina,
Brockmann II, prepared in CH2Cl2 with several drops of pyridine. Elution with a step
gradient of 0-0.5% MeOH in CH2Cl2 yielded 0.179 g of a yellow mixture. This mixture
is unsuitable for the preparation of 7; thus, 0.208 g of the mixture were flashed through a
column of neutral alumina, Brockmann II, following the same procedure noted above.
Compound 6 was isolated as an off-white foam (0.181 g, 76%).
Rf = 0.46 (1:2
EtOAc:CH2Cl2). 1H NMR (CD2Cl2, 500 MHz) (Figure 2.15): δ = 7.79 (1H, s), 7.427.37 (2H, m), 7.33-7.21 (7H, m), 6.84 (4H, m), 6.13 (1H, t, J = 6.1 Hz), 4.56 (1H, br m),
4.15 (1H, ddd, J = 4.0, 4.0, 7.9 Hz), 3.78 (6H, s), 3.37 (2H, ddd, J = 4.5, 10.9, 15.4 Hz),
2.61 (1H, ddd, J = 4.4, 6.1, 13.6 Hz), 2.43 (2H, m);
13
C NMR (CD2Cl2, 125.8 MHz)
(Figure 2.16): δ = 159.2, 145.1, 136.0, 135.8, 134.6, 130.52, 130.50, 128.46, 128.45,
127.5, 113.69, 113.68, 103.7, 88.2, 87.6, 87.2, 71.6, 63.8, 55.7, 42.2; ESI-HRMS calc’d
for C29H28N3O7Br [(M+Na+)] 632.1003, found 632.0995.
5’-O-(4,4’-Dimethoxytrityl)-5-bromo-4-nitroimidazole-β-2’-deoxy-D-ribofuranose3’-O-2-cyanoethyl-N,N-diisopropylphosphoramidite (7)
Compound 6 (50 mg, 0.082 mmol) was dried by coevaporation twice with 2 mL of
pyridine, once with 2 mL of toluene, and then by high-vacuum overnight. To the flask
containing 6 was added 0.8 mL of CH2Cl2 and diisopropylethylamine (0.043 mL, 0.25
mmol). 2-Cyanoethyl N,N-diisopropylchlorophosphoramidite (0.027 mL, 0.12 mmol)
was added dropwise by syringe and the reaction was stirred for 45 min at room
temperature. The reaction was poured into 20 mL of saturated aqueous NaHCO3 and
extracted 3 times with 20 mL of CH2Cl2. The organic layers were combined, dried over
Na2SO4, and evaporated to an orange residue. The residue was applied to a column of
basic alumina, Brockman II-III, prepared in CH2Cl2. Elution with CH2Cl2 gave 7 as a
white solid (29 mg, 44%).
Rf = 0.49 and 0.58 (two diastereomers) (10% EtOAc:
CH2Cl2). 1H NMR (CD3CN, 500 MHz) (Figure 2.17): δ = 7.88 (1H, s), 7.42-7.34 (2H,
m), 7.32-7.19 (7H, m), 6.84 (4H, m), 6.12 (1H, dddd, J = 5.7, 5.7, 5.7, 11.2 Hz), 4.71
81
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
(1H, m), 4.18 (1H, d of br m, J = 21.2 Hz), 3.86-3.51 (10H, m including two s from
−OCH3 at 3.76 and 3.75 ppm), 3.37-3.20 (2H, m), 2.72-2.62 (3H, m), 2.53 (1H, t, J = 6.0
Hz), 1.19-1.04 (12H, m); 31P NMR (CD3CN, 121.5 MHz) (Figure 2.18): δ = 150.2, 150.0
(two diastereomers); ESI-HRMS calc’d for C38H45BrN5O8P [(M+Na+)] 832.2081, found
832.2057.
NOE-Difference NMR Spectroscopy. The protons of 4 were assigned using a 2D gCOSY
spectrum (Figure 2.11).
Approximately 5 mg of 4 were dissolved in CD3OD and
transferred to an NMR tube fitted with a J. Young valve. The solution was degassed with
four freeze-pump-thaw cycles, and then sealed. An NOE-difference experiment yielded
the results in Table 2.1.
Oligodeoxynucleotide Synthesis. ODNs were synthesized on an Applied Biosystems 391
DNA synthesizer using PAC-protected phosphoramidites. The activator reagent was 0.25
M 4,5-dicyanoimidazole in CH3CN, Cap A was 5% phenoxyacetic anhydride in
THF/pyridine, Cap B was 10% 1-methylimidazole in THF, the deblocking reagent was
3% dichloroacetic acid in CH2Cl2, and the oxidizer was 0.02 M I2 in pyridine/THF/H2O.
Syntheses were performed on a 1 µmol scale using 500-Å CPG supports and all coupling
efficiencies, including that of the non-natural nucleoside, were greater than 95% as
determined by trityl monitoring.
The phosphoramidite 7 (16 mg) was dried by
coevaporation twice with 2 mL of pyridine, once with 2 mL of toluene, and then by highvacuum overnight. 7 was manually coupled by dissolving in 150 µL activator (0.25 M
DCI in CH3CN) and then pipetting directly on the CPG support. Coupling proceeded for
20 minutes.
Following DNA synthesis, ODNs containing 5-bromo-4-nitroimidazole were
converted to ODNs containing NI by treating with 1 mL of a 0.5 M solution of guanidine
in THF for 30 min at room temperature (8NI) or with 1 mL of a 0.5 M solution of
guanidine in tert-butanol for 15 h at 55 °C (9NI). The guanidine solutions were prepared
by treating 48 mg (0.5 mmol) of guanidine hydrochloride with 1 mL (0.5 mmol) of 0.5 M
NaOMe in MeOH for 5 min, evaporating the MeOH, and redissolving in 1 mL of solvent.
8NI and 9NI were deprotected and cleaved from the solid support by treating with
82
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
concentrated NH4OH at room temperature for 4 hours and 2 hours, respectively. In the
case of 8NI, the guanidine was first washed from the solid support with 10 mL of THF
prior to treating with NH4OH.
After treating 9NI with NH4OH, the solution was
neutralized with triethylammonium acetate and acetic acid and then desalted using a
NAP-10 column (Sephadex G-25, Amersham Biosciences).
ODNs containing NI were purified by reversed-phase HPLC on a Varian
Microsorb-MV analytical C-18 column (4.6 mm × 25 cm, 5 µm), using 0.1 M
triethylammonium acetate and CH3CN as solvents. The concentration of CH3CN was
increased from 0-7% in 0.7 min, then 7-15% in 30 min with a 1.0 mL/min flow rate.
ESI Mass Spectrometry. Mass spectra (Figure 2.19) were acquired by direct infusion on a
Sciex Model API 365 triple stage mass spectrometer.
MALDI-TOF Mass Spectrometry. Mass spectra (Figure 2.20) were acquired using a
matrix consisting of anthranilic acid, nicotinic acid, and diammonium citrate (2:1:0.003)
in H2O and CH3CN as described by Zhang and Gross.(24) The instrument was operated
in the linear mode with negative polarity. The accelerating voltage was 25,000 V and the
grid voltage was 95% of the accelerating voltage. An extraction delay time of 150 ns was
used.
Enzymatic Digestion Analyses. The purity and content of the ODNs 8NI and 9NI were
established by HPLC analysis of the digested ODNs (Figure 2.21).
The following
conditions were used: 100 mM Tris-Cl, pH 8.8, 15 mM MgCl2, 500 mU SVPD, and 2
nmol ODN in 40 µL were incubated at 37 °C for 4.5 h, then 20 U of alkaline phosphatase
were added and the mixture incubated for an additional 2 h.
ODN 9NI was digested with nuclease P1 using these conditions: 30 mM NaOAc,
pH 5.3, 0.1 mM ZnSO4, 17 U nuclease P1, and 2 nmol 9NI in 50 µL were incubated at 37
°C for 2 h. Alkaline phosphatase (10 U, 10 µL) and 6.6 µL 10X buffer (500 mM Tris-Cl,
83
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
pH 8.5, 1 mM EDTA) were added and the solution incubated for an additional 2 h
(Figure 2.22).
Digestion mixtures were analyzed at 254 nm by reversed-phase HPLC with a
Supelco Supercosil C-18 analytical column (2.1 mm × 25 cm, 5 µm), using 150 mM
NH4OAc and CH3CN as eluting solvents. The concentration of CH3CN was increased
from 0-15% over 40 min at a flow rate of 0.25 mL/min.
Thermal Denaturation Studies.
Melting temperatures were determined in teflon-
stoppered 1 cm pathlength quartz cells. Absorbances were monitored at 260 nm and the
temperature was cycled between 20 and 85 °C at a rate of 0.5 °C per min with a 5 min
delay at each temperature extreme. Tm values and thermodynamic parameters were
extracted from the melting curves (Figures 2.23 and 2.24) using the program MeltWin
3.5. Solutions consisted of 0.75 µM ODN in phosphate buffer (5 mM sodium phosphate,
50 mM NaCl, pH 7.0) in a total volume of 1.32 mL. All duplexes obeyed an apparent
two state transition, and annealing curves matched melting curves.
Semi-Empirical MO Calculations. Calculations were performed using Hyperchem 6.03
on a Pentium IV 1.3 GHz personal computer. The geometry of the molecule was first
optimized using the MM+ molecular mechanics method (Hyperchem’s version of MM2),
and then optimized using the AM1 semi-empirical method without constraints. Several
minima of similar energy in addition to that shown in Figure 2.25 were found (including
syn conformations), and all depicted the guanidino group out of plane with respect to the
imidazole ring. Geometries were optimized until a root-mean-square gradient of less
than or equal to 0.01 kcal/(Å mol) was achieved.
M13 Genome Constructions. ODNs 8NI and 8G were inserted into an M13mp7(L2)
bacteriophage genome using a previously reported procedure (Figure 2.26) (25). The
desalted genomes (0.3 µg) were run on a 0.9% agarose gel in 0.5X TBE buffer. The gel
was run for 5 h at 100 V, and then stained in a 1X solution of SYBR Gold (Molecular
Probes, Inc.) in 1X TBE buffer (200 mL). The ligation efficiency of ODN insertion into
84
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
the genome was approximately 38% as determined by quantification of the stained DNA.
Circular wild-type M13 DNA and linear M13 DNA (linearized by digestion with EcoRI
restriction endonuclease) were used as markers.
85
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
2.4. Results and Discussion
The convertible nucleoside 4 was synthesized in three steps beginning with 5(4)bromo-1H-imidazole (1) (Scheme 1). 5(4)-Bromo-4(5)-nitroimidazole (2) was prepared
in 91% yield by nitrating 1 (23). Treatment of 2 with NaH in CH3CN, followed by
condensation with 3,5-di-O-toluoyl-α-1-chloro-2-deoxy-D-ribofuranose gave exclusively
two β-nucleosides, the 5-bromo-4-nitroimidazole isomer 3 in 50% yield and the 4-bromo5-nitroimidazole isomer in 35% yield. The structures of these isomers were assigned by
comparison of the UV spectra with those of literature analogs (26, 27). Interestingly, 4halo-5-nitroimidazoles have been reported to be resistant to nucleophilic displacement of
the halo substituent (28, 29). In the present study, the 5-bromo-4-nitroimidazole isomer
readily undergoes reaction at the site of the bromo substituent, while the 4-bromo-5nitroimidazole isomer was resistant to reaction at the bromo position under the same
conditions.
Saponification of the toluoyl esters of 3 was achieved in 90% yield by treatment
with guanidine in MeOH for 3.75 hours at 0 °C. Initial attempts to deprotect 3 using
reagents such as NaOMe in MeOH, NH4OH, and K2CO3 in MeOH were unsuccessful
because of decomposition of 3, presumably as a result of displacement of the bromo
substituent.
The β conformation of 4 was confirmed by NOE NMR spectroscopy.
Irradiating at the 1’ proton, we observed an NOE enhancement at the 4’ proton, and by
irradiating the imidazolic proton, NOE enhancements were observed at the 3’ proton and
5’ protons, thus confirming the structure. It is anticipated that 4 will serve as a branching
point for generating libraries of 5-substituted-4-nitroimidazole nucleosides and, when
incorporated into DNA, 5-substituted-4-nitroimidazole-containing ODNs, since previous
studies have shown the bromo substituent of 5-bromo-4-nitroimidazoles to be displaced
by carbon (27), sulfur and oxygen (29), and nitrogen (9) nucleophiles.
For later use as an analytical standard, the 2’-deoxynucleoside of NI (dNI, 5) was
synthesized from 4 in nearly quantitative yield by treatment with guanidine in refluxing
EtOH (Scheme 2). Having prepared 4 and 5, we then determined the stability of these
compounds to DNA synthesis conditions. The half-life of 4 was found to be less than 15
86
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
min in 3% trichloroacetic acid in CH2Cl2, but greater than 120 minutes in 2%
dichloroacetic acid in CH2Cl2. Compound 4 was also stable for at least 24 h under
standard oxidizing conditions used for DNA synthesis (0.02 M I2 in pyridine/H2O) and
therefore was determined to have the stability necessary for automated DNA synthesis.
Compound 5 was stable to treatment with concentrated NH4OH for at least 4 h at room
temperature. Under these conditions, phenoxyacetyl-protected ODNs can be cleaved
from the solid support and deprotected.
Compound 4 was protected as the 5’-O-dimethoxytrityl ether using DMT-Cl,
triethylamine, and a catalytic amount of DMAP in pyridine (Scheme 2). Two rounds of
purification of the tritylated product by flash chromatography were necessary and gave
the desired compound 6 in 76% yield. Compound 6 was subsequently treated with 2cyanoethyl N,N-diisopropylchlorophosphoramidite and diisopropylethylamine to produce
the phosphoramidite 7 in 44% yield. Alumina chromatography was necessary for the
purification of 6 and 7, as both were unstable to purification on silica, even in the
presence of triethylamine to reduce the acidity of the silica.
Compound 7 was incorporated into a 19mer ODN of mixed base sequence (8X,
see Table 2.2 for structure and nomenclature) using standard automated procedures, with
the exception that 3% dichloroacetic acid in CH2Cl2 was substituted for the deblocking
reagent to minimize abasic site formation. The coupling efficiency after 20 min was
greater than 95% as judged by trityl monitoring. Postsynthetic treatment of the DNA
with 0.5 M guanidine in THF at room temperature caused a rapid color change of the
solid support from white to bright yellow, indicating the production of NI. Deprotection
of the DNA and cleavage from the support was achieved with concentrated NH4OH for 4
h at room temperature. HPLC analysis of the crude 8NI (Figure 2.27) showed a single
major peak with absorbance maxima at 260 and 380 nm.
The purified 8NI was
characterized by nuclease and phosphatase digestion followed by HPLC analysis (Figure
2.21) and by MALDI-TOF (Figure 2.20) and ESI mass spectrometry (Figure 2.19).
87
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Digestion of 8NI with snake venom phosphodiesterase (SVPD) and alkaline
phosphatase yielded the five expected nucleosides, with the dNI peak eluting with the
same retention time and having the same UV-vis spectrum as the authentic standard 5. It
has been reported that SVPD is unable to cleave the phosphodiester bond in the sequence
context 5’-C[NI]-3’ (8).
We did not observe this effect in our sequence context;
however, in the sequence 5’-TT[NI]TTTTTTT-3’ (9NI), while SVPD could completely
digest the ODN, nuclease P1 could not and left an NI-containing dinucleotide (Figure
2.22). As previously reported, MALDI mass spectrometry allowed detection of two
degradation products, presumably of the nitro substituent induced by the MALDI laser
(8). MALDI-induced degradation has been previously observed during MALDI analysis
of aromatic nitro compounds (30, 31). Analysis by ESI mass spectrometry did not show
these degradation products.
Thermal denaturation studies were performed to determine the stability of
duplexes containing NI paired with each of the four natural bases (8NI⋅10Y) relative to
the stability of unmodified duplexes (8G⋅10Y) (Table 2.2, which also defines
nomenclature for 8 and 10). We found the NI lesion greatly destabilizes the duplex. The
presence of the NI lesion results in a 10−12 °C decrease in Tm relative to an unmodified
duplex containing a G⋅C base pair at the same position and a 4−6 °C decrease in Tm
relative to the least thermally stable mismatch of the unmodified duplex (G⋅T). A
comparison of all Tm values and free energies shows a greater difference in magnitude
among the unmodified duplexes than among the NI-containing duplexes, demonstrating
that the presence of the lesion generally reduces the effect of the opposing base on duplex
stability.
Given that a large reduction in thermal stability does not imply a large change in
duplex structure and similarity among thermal stabilities does not necessitate structural
similarities (32), it is not possible to draw conclusions from the thermal stabilities alone
about the effect of the NI lesion on duplex DNA structure. In an effort to probe the
origins of the observed duplex destabilization, a simple AM1 semiempirical method (33)
was used to optimize the geometry of 5 (Figure 2.25). The calculation predicts that the
88
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
guanidino group will be rotated out of plane in relation to the imidazole ring and extend
roughly 3 Å perpendicularly in either direction from the plane of the ring.
Accommodation of such a structure into the helix would require at minimum a local
distortion of the duplex and a disruption of the base π-stacking interactions.
We have site-specifically inserted 8NI into an M13 bacteriophage genome for
mutagenesis and genotoxicity studies (Figure 2.26) (25). The ligation efficiency of the
genome construction was similar to that observed for 8G. Mutagenesis and repair studies
using this construct will be reported elsewhere.
In summary, we have developed a synthetic route for the preparation of 5 and
provided a method for the incorporation of NI lesions into DNA by the phosphoramidite
method utilizing the convertible nucleoside approach.
Ready access to 5 will be
beneficial for use as an analytical standard in future studies aimed at detecting the NI
lesion in vivo, whereas the convertible nucleoside phosphoramidite will allow for the
rapid preparation of NI-containing ODNs for mutagenesis and DNA repair studies,
creation of libraries of 5-substituted-4-nitro analogs of NI (which may be useful as
structural probes), and preparation of NI-containing ODNs in sufficient quantities for
NMR and X-ray structure studies.
89
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
2.5. Acknowledgment
Paul T. Henderson (Lawrence Livermore National Laboratory) and John M.
Essigmann (MIT) contributed to this work. We thank Dr. Uday Sharma for helpful
discussions, the Sadighi laboratory at MIT for assistance with NOE NMR experiments,
Prof. Catherine L. Drennan for the use of her UV-vis spectrophotometer, and Hector H.
Hernandez for assistance with Tm experiments. We also acknowledge the MIT Biological
Engineering Division Mass Spectrometry Laboratory and the Department of Chemistry
Instrumentation Facility. Financial support was provided by NCI award P01-CA26731
and by NSF awards DBI-9729592 and CHE-9808061.
90
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.1. 1H NMR spectrum of 2 in DMSO-d6.
91
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.2.
13
C NMR spectrum of 2 in DMSO-d6.
92
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.3. 1H NMR spectrum of 3A in CD2Cl2.
93
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.4.
13
C NMR spectrum of 3A in CD2Cl2.
94
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.5. UV-vis spectrum of 3A in CD2Cl2.
Absorbance
240 nm
315 nm
250
300
350
Wavelength / nm
95
400
450
500
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.6. 1H NMR spectrum of 3 in CD2Cl2.
96
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.7.
13
C NMR spectrum of 3 in CD2Cl2.
97
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.8. UV-vis spectrum of 3 in CD2Cl2.
Absorbance
241 nm
296 nm
250
300
350
400
Wavelength / nm
98
450
500
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.9. 1H NMR spectrum of 4 in CD3OD.
99
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.10.
13
C NMR spectrum of 4 in CD3OD.
100
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.11. gCOSY spectrum of 4 in CD3OD.
101
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.12. 1H NMR spectrum of 5 in D2O.
102
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.13.
13
C NMR spectrum of 5 in CD3OD.
103
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.14 UV-vis spectrum of 5 in H2O, pH 7.
AU
230 nm
0.5
0.4
0.3
380 nm
0.2
0.1
0
220
270
320
370
Wavelength / nm
104
420
470
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.15. 1H NMR spectrum of 6 in CD2Cl2.
105
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.16.
13
C NMR spectrum of 6 in CD2Cl2.
106
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.17. 1H NMR spectrum of 7 in CD3CN.
107
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.18.
31
P NMR spectrum of 7 in CD3CN.
108
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.19. (Top) ESI mass spectrum for 8NI: calc’d 5882.9, found 5884.0. (Bottom)
ESI mass spectrum for 9NI: calc’d 3024.0, found 3024.6.
109
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.20. (Top) MALDI-TOF mass spectrum for 8NI: calc’d 5882.9, found 5882.1.
(Bottom) MALDI-TOF mass spectrum for 9NI: calc’d 3024.0, found 3025.0.
5866.34
100
90
5851.46
% Intensity
80
70
60
50
5882.07
40
30
20
10
0 5400
5600
5800
6000
6200
6400
3300
3500
Mass (m/z)
2993.33
100
90
% Intensity
80
3009.11
70
60
50
40
30
3024.98
20
10
0 2500
2700
2900
3100
Mass (m/z)
110
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.21. Digestion analyses of 8NI (Top) and 9NI (Bottom) using SVPD and
alkaline phosphatase. The insets show the UV-vis spectra of the dNI peaks.
mAU
125
AU
dG
dA
100
2.0
75
50
25
1.5
250
300
350
400
450 nm
1.0
dC
0.5
T
dNI
0.0
0
AU
10
20
Retention Time / min
30
T
mAU
100
1.5
75
50
25
1.0
250
300
350
400
450 nm
0.5
dNI
0.0
0
10
20
Retention Time / min
111
30
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.22. Digestion analysis of 9NI using nuclease P1 and alkaline phosphatase. The
inset shows the UV-vis spectrum of the dinucleotide peak.
AU
mAU
1.5
500
T
400
300
200
100
1.0
250
300
350
400
450 nm
dNI-T
or
T-dNI
0.5
0.0
0
20
10
Retention Time / min
112
30
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.23. Representative melting curves for 8NI annealed with 10G (top left), 10A (top right), 10C (bottom left), and 10T (bottom
right).
0.33
0.32
Absorbance
Absorbance
0.32
0.31
0.30
0.29
0.28
0.29
0.28
0.26
20
30
40
50
60
Temperature / °C
70
80
20
0.31
0.31
0.30
0.30
0.29
0.28
0.27
40
50
60
70
80
70
80
0.29
0.28
0.27
0.26
0.26
0.25
20
30
Temperature / °C
Absorbance
Absorbance
0.30
0.27
0.27
0.26
0.31
30
40
50
60
70
0.25
20
80
Temperature / °C
30
40
50
60
Temperature / °C
113
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.24. Representative melting curves for 8G annealed with 10G (top left), 10A (top right), 10C (bottom left), and 10T (bottom
0.30
0.31
0.29
0.30
Absorbance
Absorbance
right).
0.28
0.27
0.26
0.25
0.24
20
0.29
0.28
0.27
0.26
0.25
30
40
50
60
70
0.24
20
80
30
0.30
0.30
0.29
0.29
0.28
0.27
0.26
0.25
60
70
80
70
80
0.28
0.27
0.26
0.25
0.24
0.24
0.23
20
50
Temperature / °C
Absorbance
Absorbance
Temperature / °C
40
30
40
50
60
Temperature / °C
70
0.23
20
80
114
30
40
50
60
Temperature / °C
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.25. Energy-minimized (AM1) structure of dNI (5). The imidazole ring and
nitro group are in the plane of the page, while the carbon and two of the nitrogen atoms
of the guanidino group are projecting through the back of the page.
115
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.26. Agarose gel analysis of the M13 genome constructions. (1) Circular M13
DNA. (2) Linear M13 DNA. (3) Insertion of 8G into M13 DNA. (4) Insertion of 8NI
into M13 DNA.
Single-Stranded
Circular Genome
Single-Stranded
Linear Genome
1
2
3
4
116
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Figure 2.27. (Top) HPLC trace at 254 nm of the converted, deprotected 8NI and purified
Absorbance
8NI (inset). (Bottom) UV-vis spectrum of purified 8NI.
15.0 17.5 20.0 22.5 25.0 27.5
Retention Time / min
5
10
15
20
25
30
Absorbance
Retention Time / min
DNA
NI
250
300
350
400
Wavelength / nm
117
450
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Scheme 2.1. Synthesis of the convertible nucleoside 4.
N
N
H
Br
b) H2SO4
N
H
1
p-TolO
O
p-TolO
NO2 a) NaH, CH CN
3
b) p-TolO
Br
O
N
a) HNO3, EtOH
N
NO2
N
Br
2
Guanidine
HO
o
O
MeOH, 0 C
3
HO
118
Cl
p-TolO
N
NO2
N
Br
4
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Scheme 2.2. Synthesis of 5 and the convertible nucleoside phosphoramidite 7.
HO
O
N
NO2
N
Br
DMTO
DMT-Cl, DMAP
TEA, Pyridine
4
HO
HO
NO2
NH
O
HO
N
N
H
NO2
N
Br
6
HO
DIPEA, CH2Cl2
Cl
P
(i-Pr)2N
O(CH2)2CN
Guanidine
EtOH, reflux
N
O
N
DMTO
O
NH2
O
5
(i-Pr)2N
119
N
NO2
N
Br
7
O(CH2)2CN
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Table 2.1. NOE difference for 4 in CD3OD (%).
observed proton
imidazolic H
1’H
2’H
3’H
4’H
5’H
irradiated proton
1’H
imidazolic H
1.41

1.73

6.43
3.27
0.28
2.65
2.20
0.49
0.10
1.47
120
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
Table 2.2. Melting Temperatures and Free Energies of Duplexesa.
5’-GCG AAG ACC GXA GCG TCC G-3’
3’-CGC TTC TGG CYT CGC AGG C-5’
8X
10Y
X
Y
NI
NI
NI
NI
G
G
G
G
G
A
C
T
G
A
C
T
Tm (± 0.5 °C)
57.0
56.4
55.8
54.8
61.0
62.4
66.9
60.6
a
−∆G37ºC (kcal/mol)
16.8 ± 0.1
16.7 ± 0.0
16.7 ± 0.1
16.0 ± 0.0
19.3 ± 0.0
20.4 ± 0.0
23.0 ± 0.1
19.4 ± 0.0
Conditions: 50 mM sodium chloride, 5 mM sodium phosphate, pH 7.0. Tm and −∆G37ºC
values are the average of three melts.
121
Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
2.6. References
(1) Cadet, J., Bellon, S., Berger, M., Bourdat, A. G., Douki, T., Duarte, V., Frelon, S.,
Gasparutto, D., Muller, E., Ravanat, J. L., and Sauvaigo, S. (2002) Recent
aspects of oxidative DNA damage: guanine lesions, measurement and
substrate specificity of DNA repair glycosylases. Biol. Chem. 383, 933943.
(2) Burney, S., Niles, J. C., Dedon, P. C., and Tannenbaum, S. R. (1999) DNA
damage in deoxynucleosides and oligonucleotides treated with
peroxynitrite. Chem. Res. Toxicol. 12, 513-520.
(3) Kornyushyna, O., Berges, A. M., Muller, J. G., and Burrows, C. J. (2002) In vitro
nucleotide misinsertion opposite the oxidized guanosine lesions
spiroiminodihydantoin and guanidinohydantoin and DNA synthesis past
the lesions using Escherichia coli DNA polymerase I (Klenow fragment).
Biochemistry 41, 15304-15314.
(4) Lee, J. M., Niles, J. C., Wishnok, J. S., and Tannenbaum, S. R. (2002)
Peroxynitrite reacts with 8-nitropurines to yield 8-oxopurines. Chem. Res.
Toxicol. 15, 7-14.
(5) Tretyakova, N. Y., Niles, J. C., Burney, S., Wishnok, J. S., and Tannenbaum, S.
R. (1999) Peroxynitrite-induced reactions of synthetic oligonucleotides
containing 8-oxoguanine. Chem. Res. Toxicol. 12, 459-466.
(6) Burrows, C. J., Muller, J. G., Kornyushyna, O., Luo, W., Duarte, V., Leipold, M.
D., and David, S. S. (2002) Structure and potential mutagenicity of new
hydantoin products from guanosine and 8-oxo-7,8-dihydroguanine
oxidation by transition metals. Environ. Health Perspect. 110, 713-717.
(7) Gu, F., Stillwell, W. G., Wishnok, J. S., Shallop, A. J., Jones, R. A., and
Tannenbaum, S. R. (2002) Peroxynitrite-induced reactions of synthetic
oligo 2'-deoxynucleotides and DNA containing guanine: formation and
stability of a 5-guanidino-4-nitroimidazole lesion. Biochemistry 41, 75087518.
(8) Joffe, A., Mock, S., Yun, B. H., Kolbanovskiy, A., Geacintov, N. E., and
Shafirovich, V. (2003) Oxidative generation of guanine radicals by
carbonate radicals and their reactions with nitrogen dioxide to form site
specific 5-guanidino-4-nitroimidazole lesions in oligodeoxynucleotides.
Chem. Res. Toxicol. 16, 966-973.
(9) Niles, J. C., Wishnok, J. S., and Tannenbaum, S. R. (2001) A novel
nitroimidazole compound formed during the reaction of peroxynitrite with
2',3',5'-tri-O-acetyl-guanosine. J. Am. Chem. Soc. 123, 12147-12151.
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Chapter 2 – Synthesis of 5-Guanidino-4-nitroimidazole
(10) Henderson, P. T., Delaney, J. C., Gu, F., Tannenbaum, S. R., and Essigmann, J.
M. (2002) Oxidation of 7,8-dihydro-8-oxoguanine affords lesions that are
potent sources of replication errors in vivo. Biochemistry 41, 914-921.
(11) Kino, K., and Sugiyama, H. (2001) Possible cause of G-C-->C-G transversion
mutation by guanine oxidation product, imidazolone. Chem. Biol. 8, 369378.
(12) Shafirovich, V., Mock, S., Kolbanovskiy, A., and Geacintov, N. E. (2002)
Photochemically catalyzed generation of site-specific 8-nitroguanine
adducts in DNA by the reaction of long-lived neutral guanine radicals with
nitrogen dioxide. Chem. Res. Toxicol. 15, 591-597.
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strategy for users. Annu. Rev. Biochem. 67, 99-134.
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oligonucleotide suicide substrate for DNA methyltransferases. J. Org.
Chem. 57, 2989-2991.
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precursor of 2-aminoimidazolone, a major guanine oxidation product in
one-electron oxidation of DNA. J. Am. Chem. Soc. 121, 10836-10837.
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site-specific modification of RNA: The convertible nucleoside approach.
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containing 5-(N-aminoalkyl)carbamoyl-2'-deoxyuridines by a new
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nuclease-resistance properties. Bioconjugate Chem. 8, 271-280.
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124
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(32) Plum, G. E., Grollman, A. P., Johnson, F., and Breslauer, K. J. (1995) Influence
of the oxidatively damaged adduct 8-oxodeoxyguanosine on the
conformation, energetics, and thermodynamic stability of a DNA duplex.
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125
CHAPTER 3
In Vivo Bypass Efficiencies and Mutational Signatures of
the Guanine Oxidation Products 2-Aminoimidazolone and
5-Guanidino-4-nitroimidazole
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Chapter 3 – Iz and NI are Mutagenic in Vivo
3.1. Abstract
The in vivo mutagenic properties of 2-aminoimidazolone and 5-guanidino-4nitroimidazole, two products of peroxynitrite oxidation of guanine, are reported. Two
oligodeoxynucleotides of identical sequence, but containing either 2-aminoimidazolone
or 5-guanidino-4-nitroimidazole at a specific site, were ligated into single-stranded
M13mp7L2 bacteriophage genomes. Wild-type AB1157 Escherichia coli cells were
transformed
with
the
site-specific
2-aminoimidazolone-
and
5-guanidino-4-
nitroimidazole-containing genomes, and analysis of the resulting progeny phage allowed
determination of the in vivo bypass efficiencies and mutational signatures of the DNA
lesions. 2-Aminoimidazolone was efficiently bypassed and 91% mutagenic, producing
almost exclusively G to C transversion mutations.
In contrast, 5-guanidino-4-
nitroimidazole was a strong block to replication and 50% mutagenic, generating G to A,
G to T and, to a lesser extent, G to C mutations. The G to A mutation elicited by 5guanidino-4-nitroimidazole implicates this lesion as a novel source of peroxynitriteinduced transition mutations in vivo. For comparison, the error-prone bypass DNA
polymerases were over-expressed in the cells by irradiation with UV light (SOS
induction) prior to transformation. SOS induction caused little change in the efficiency
of DNA polymerase bypass of 2-aminoimidazolone; however, bypass of 5-guanidino-4nitroimidazole increased nearly 10-fold. Importantly, the mutation frequencies of both
lesions decreased during replication in SOS-induced cells. These data suggest that 2aminoimidazolone and 5-guanidino-4-nitroimidazole in DNA are substrates for one or
more of the SOS-induced Y-family DNA polymerases and demonstrate that 2aminoimidazolone and 5-guanidino-4-nitroimidazole are potent sources of mutations in
vivo.
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Chapter 3 – Iz and NI are Mutagenic in Vivo
3.2. Introduction
Oxidative damage of DNA is implicated as a cause of aging (1-3), carcinogenesis
(4-6), a variety of noncancerous diseases such as Alzheimer’s disease and cardiovascular
disease (7), and the progression to acquired immunodeficiency syndrome in HIV-infected
patients (8). The reactive species responsible for DNA damage are generated by common
endogenous processes (9) such as respiration and inflammation (10, 11).
During
inflammation, an assortment of reactive oxygen and nitrogen intermediates are generated
by activated immune system cells (11), and reaction of these molecules with DNA
produces dozens of oxidized nucleobase derivatives (12).
The radicals nitric oxide (•NO) and superoxide (O2•−) are produced by macrophages
and neutrophils (two types of inflammatory cells) upon immune response activation (13,
14).
These radicals combine in a diffusion-limited reaction to form peroxynitrite
(ONOO−) (15), a powerful oxidizing and nitrating agent capable of damaging a variety of
biomolecules (16-18), including DNA (19). Under physiological conditions, ONOO−
rapidly combines with CO2 to form nitrosoperoxycarbonate (ONOOCO2−), which
subsequently undergoes homolysis to produce carbonate radical (CO3•−) and nitrogen
dioxide (•NO2) (20-22). These radicals are believed to be responsible for the oxidation
and nitration of DNA caused by exposure to ONOO− (23-25).
Since guanine possesses the lowest redox potential of the four DNA nucleobases (E7
= 1.27 V vs. NHE) (26), it is preferentially oxidized by ONOO− compared to the other
natural nucleobases (27). Several products are formed directly from guanine residues in
DNA including 7,8-dihydro-8-oxoguanine (8-oxoG), 8-nitroguanine (8-NO2-G), 2aminoimidazolone (Iz), and 5-guanidino-4-nitroimidazole (NI) (Figure 3.1) (28). The
lesions 8-oxoG and 8-NO2-G are highly susceptible to further oxidation (27) and yield a
variety of additional products (29, 30).
Although many of these guanine-derived
oxidation products have been characterized for their in vivo mutagenic potential (31, 32),
Iz and NI have received little attention. In order to assess the biological significance and
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Chapter 3 – Iz and NI are Mutagenic in Vivo
consequences of oxidatively damaged DNA, it is essential that these lesions be
characterized for their genotoxic and mutagenic potential.
In the work presented here, we report the in vivo genotoxic and mutagenic properties
of Iz and NI. Oligodeoxynucleotides (ODNs), site-specifically modified with Iz or NI,
were synthesized using a previously described method (33) and a procedure developed in
our laboratory (34). The biological impact of unique Iz and NI lesions was addressed
under normal and SOS-induced conditions in wild-type AB1157 E. coli cells using viral
vectors containing the modified ODNs. Both lesions were bypassed by the E. coli
replication machinery and were substrates for SOS-induced error-prone DNA polymerase
bypass. Furthermore, each lesion was potently mutagenic during DNA replication.
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Chapter 3 – Iz and NI are Mutagenic in Vivo
3.3. Experimental Procedures
Oligodeoxynucleotides. DNA synthesis reagents were purchased from Glen Research.
Unmodified ODNs were purchased from IDT, Inc. and were purified by polyacrylamide
gel electrophoresis (PAGE). The 19mer ODN sequence used was 5’-GCG AAG ACC
GXA GCG TCC G-3’, where X is G, Iz, NI, or a tetrahydrofuran (THF) abasic site. The
19mer containing the THF abasic site analog was prepared as described (31).
The 19mer containing Iz was prepared as previously described (33). Briefly, 8methoxy-2’-deoxyguanosine (8-MeOdG) was incorporated into the 19mer ODN by the
phosphoramidite method and the ODN was deprotected and cleaved from the solid
support with concentrated NH4OH at 55 °C for 15 h. The 8-MeOdG-containing ODN
was purified by PAGE, and the 8-MeOdG subsequently converted to Iz by
photoirradiation with 365 nm light in the presence of riboflavin. The Iz-containing
19mer was purified by anion exchange HPLC on a Dionex NucleoPac PA-100 (4 × 250
mm) analytical column using 10% CH3CN in water (solvent A) and aqueous 1.5 M
NH4OAc (solvent B) as solvents. A flow rate of 1.0 mL/min was used and solvent B was
increased from 10% to 25% over 2.5 min, then 25% to 100% over 30 min. The purified
19mer was characterized using MALDI-TOF mass spectrometry (calc’d MW: 5824.8;
found: 5825.8) and by enzymatic digestion to nucleosides followed by HPLC analysis.
The following conditions were used for the enzymatic digestion: 50 mM Tris-Cl, pH 7.0,
0.1 mM ZnSO4, 18 units of nuclease P1, 12 units of alkaline phosphatase (both enzymes
from Roche Applied Science), and 2 nmol of Iz-containing 19mer in 50 µL were
incubated at room temperature for 30 min and then immediately analyzed by HPLC. For
the HPLC analysis, a Supelco Supercosil LC-18-DB (250 × 2.1 mm, 5 µm) column was
used with aqueous 150 mM NH4OAc as solvent A and CH3CN as solvent B. A flow rate
of 0.25 mL/min was used and solvent B was increased from 0% to 15% over 40 min.
Five peaks were observed with UV-vis spectra consistent with the nucleosides Iz, C, G,
T, and A.
The ODN containing NI was prepared by incorporating 5-bromo-4-nitroimidazole
into the 19mer sequence by the phosphoramidite method and subsequently treating the
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Chapter 3 – Iz and NI are Mutagenic in Vivo
19mer with 0.5 M guanidine in THF and then concentrated NH4OH to produce the 19mer
containing NI (34). The NI-containing 19mer was purified by C18 reversed phase HPLC
(34) and by anion exchange HPLC as described above, and characterized using MALDITOF mass spectrometry (calc’d MW: 5882.9; found: 5882.7) and by enzymatic digestion
with snake venom phosphodiesterase (ICN Biomedical) and alkaline phosphatase (34)
followed by HPLC analysis (same method as described for the 19mer containing Iz). For
the HPLC analysis, five peaks were observed with UV-vis spectra consistent with the
nucleosides NI, C, G, T, and A.
Genome Construction. Genomes were constructed in triplicate (Figure 3.2) on a 10 pmol
scale, as previously described (31, 32, 35). Briefly, the ss M13 DNA was linearized by
cleavage with EcoRI (30 U/pmol of DNA, 23 °C for 8 h) at a hairpin containing a single
EcoRI site (35). The genome was recircularized by annealing in the presence of the 5’phosphorylated 19mer insert and two “scaffold” ODNs (sequences that are partially
complementary to the 5’ and 3’ sides of the insert and the genomic DNA termini) and
incubating with 22.5 U/µL of T4 DNA ligase (NEB) for 2 h at 16 °C in a volume of 55
µL (31). Two short scaffolds are used in order to leave a ss gap at the site of the lesion,
thereby facilitating efficient ligation regardless of the lesion structure. The scaffold DNA
was removed using the exonuclease activity of T4 DNA polymerase (Amersham) by
treating with 0.25 U/µL for 1 h at 16 °C in a volume of 65 µL (36, 37). Under these
conditions, scaffold digestion was complete as determined by using a radiolabeled
scaffold and analyzing the reaction with PAGE and phosphorimagery. The genome
constructs were diluted to 115 µL with H2O, extracted with 100 µL of
phenol:chloroform:isoamyl alcohol (25:24:1) (Invitrogen), and desalted with Sephadex
G50 fine resin (Amersham Biosciences).
The stability of the lesion-containing 19mers was assessed using MALDI-TOF mass
spectrometry. The 19mers were exposed to the conditions used for genome construction,
with the exception that M13 DNA, other ODNs, and enzymes were excluded. The mass
spectra of the 19mers were essentially unchanged after exposure to the mock genome
construction conditions. The amount of circular, 19mer insert-containing genome in each
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Chapter 3 – Iz and NI are Mutagenic in Vivo
sample was quantified by agarose gel electrophoresis and phosphorimagery. This was
accomplished by annealing 1 pmol of a 5’-[32P]-labeled 30mer probe ODN (5’-TCC
CAG TCA CGA CGT TGT AAA ACG ACG GCC-3’) to 0.1 pmol of each genome
construct in a region of the genome that did not include the lesion-containing 19mer
insert.
The annealing solution consisted of 100 mM NaCl, 4.2% Ficoll, 0.042%
bromophenol blue, and 0.042% xylene cyanol FF in a volume of 15.5 µL. The genomeprobe mixtures (15.5 µL) were run on a 0.9% agarose gel in 1X TBE buffer for 4 h at 100
V, after which the free probe front was excised. The gel was run for an additional 3 h
and then transferred onto a glass plate and dried under a box fan for 36 h. The amount of
circular genome was quantified by phosphorimagery and the genome construct solution
volumes adjusted such that each solution used for cell transformation contained an equal
concentration of circular DNA.
Preparation of Electrocompetent Cells. 2 × 150 mL of LB media were each inoculated
with 1.5 mL from separate overnight cultures of wild-type AB1157 E. coli and grown on
a shaker at 37 °C to an A600 of approximately 0.4. Each culture was centrifuged,
resuspended in 25 mL of 10 mM MgSO4, and transferred to a large (150 × 15 mm) Petri
dish. The SOS system was induced in the cells by irradiating with 254 nm light (45 J/m2
of energy), immediately transferring the cells to 2 × 125 mL of 2×YT media, and
growing for 40 min at 37 °C with shaking. Uninduced cells were treated identically,
except without exposure to UV light. The 2×YT cultures were centrifuged, combined,
and washed with 2 × 175 mL of deionized water. The electrocompetent cells were
resuspended in 4 mL of a 10% solution of glycerol in water, stored at 4 °C, and used the
following day.
Translesion Bypass Efficiency. An equal amount of each circular genome construct was
mixed with 0.025 pmol of internal standard (wild-type circular single-stranded
M13mp7L2 DNA) and 100 µL of electrocompetent cells. The cell/genome mixtures
were electroporated and transferred to 10 mL of LB, generating at least 105 independent
transformed cells as determined by plating of an aliquot onto agar plates. The cultures
were incubated at room temperature for at least 30 min, and then incubated on a roller
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Chapter 3 – Iz and NI are Mutagenic in Vivo
drum for 4 h at 37 °C to amplify the progeny phage. The cells were spun down and the
progeny phage-containing supernatant retained. In this system, successful replication of
the genomes by the E. coli host leads to the production of progeny phage. When plated
on a lawn of NR9050 indicator E. coli in the presence of IPTG and X-gal (Gold
Biotechnology), progeny phage derived from genomes containing the 19mer insert
produce blue plaques if no mutation, a point mutation, or an in frame insertion or deletion
mutation occurs at the lesion site, whereas progeny phage from WT M13mp7L2
genomes, genomes lacking an insert (genetic engineering mutants), and lesion induced
out-of-frame insertions and deletions produce clear plaques (31). Thus, the amplified
progeny phage were diluted and plated such that approximately 1000-2000 total plaques
were produced per Petri dish, and the number of blue and clear plaques per plate were
counted. The number of normalized blue plaques resulting from each lesion-containing
genome relative to that of the normalized guanine control indicated the bypass efficiency
of a lesion (see Results for details). The normalization allowed direct comparison of
lesion bypass by DNA polymerase(s) between genomes containing different inserts,
regardless of unavoidable variation in factors such as electroporation efficiency and the
total number of plaques on each plate.
Mutation Type and Frequency.
Electrocompetent cells were transformed with each
genome construct, as described in the previous section, except that the internal standard
was excluded.
As before, the electroporations produced at least 105 independent
transformed cells. Amplified progeny phage (100 µL) and 10 µL of an overnight growth
of SCS110 E. coli were added to 10 mL of LB media and grown for 6 h at 37 °C on a
roller drum.
The cultures were centrifuged and the phage-containing supernatant
retained. Single-stranded phage DNA was isolated from 700 µL of each sample using a
QIAPrep Spin M13 kit (Qiagen). The region containing the lesion site was amplified by
PCR as previously described (31), except 0.7 µM of each primer was used, and the
resultant 101mer PCR product was purified using a QIAPrep PCR purification kit
(Qiagen). The REAP assay was used to determine the identity of the base at the site
formerly occupied by the lesion (31, 32, 35, 38).
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Chapter 3 – Iz and NI are Mutagenic in Vivo
3.4. Results
Genome construction. Convertible nucleoside phosphoramidites were used to introduce
each oxidized DNA lesion into a 19mer ODN at a defined site. Following incorporation
into the 19mer by automated DNA synthesis, 8-methoxyguanine was converted to Iz by
photoirradiation in the presence of riboflavin (33). Using a procedure recently developed
in our laboratory, NI was formed within an ODN from the convertible nucleoside 5bromo-4-nitroimidazole by treatment of the ODN with guanidine (34). Each ODN was
characterized by MALDI-TOF mass spectrometry and by nuclease and phosphatase
digestion followed by HPLC analysis.
Single-stranded M13mp7L2 viral genomes
containing Iz or NI at a specific site in the genome were constructed as shown in Figure
3.2. An aliquot of each genome construct, annealed to a radiolabeled ODN probe, was
analyzed by agarose gel electrophoresis and phosphorimagery to assess the yield of
circular, insert-containing genome.
Following quantification, the genome construct
solutions were normalized such that each contained an equivalent amount of circular,
insert-containing genome.
The stability of each lesion to the genome construction
conditions was confirmed by MALDI-TOF mass spectrometry after subjecting the lesioncontaining ODNs to the same conditions.
Translesion Bypass Efficiency. A viral plaque assay based on lacZ α-complementation
was employed for the determination of the lesion bypass efficiency (Figure 3.3). In this
system, phage produced from genomes containing the 19mer insert cause the formation
of blue plaques when plated on IPTG/X-gal indicator plates if either no mutation or a
point mutation occurs at the lesion site, and phage produced from WT M13mp7L2
genomes produce clear plaques. The number of phage that form blue plaques varies with
the efficiency of lesion bypass, and the rate of replication of the WT M13mp7L2
genomes is assumed to be independent of the identity of the insert-containing genomes.
Thus, a genome containing a lesion that partially blocks replication yields a lower
proportion of progeny phage relative to a genome containing a freely bypassed lesion.
Since the same amount of internal standard (WT M13mp7L2 genome) was used in each
mixture, the blue-to-clear ratio of each mixture can be directly compared after being
normalized to reflect the identical amount of internal standard in each mixture. The
134
Chapter 3 – Iz and NI are Mutagenic in Vivo
bypass efficiency of a lesion is scaled relative to guanine, which is defined as having a
bypass efficiency of 100%. From this logic the equation given in Figure 3.3 results,
where the term B(L)/B(G) is the definition of relative bypass efficiency and the term
C(G)/C(L) is the normalization factor. A limitation of this assay is that lesion induced
out-of-frame insertion and deletion mutations would also produce clear plaques and
therefore not be counted as bypass events. However, as is discussed in the next section
the lesions studied here induced negligible amounts of observable insertion and deletion
mutations and so a correction for these events was not performed.
In normal WT AB1157 E. coli, the bypass efficiency of Iz was 60 ± 5% (Figure
3.4). By contrast, NI was a much stronger replication block and was bypassed with an
efficiency of only 7.0 ± 1.6%. As a control, we also determined the bypass efficiency of
a synthetic THF abasic site. The THF lesion was bypassed with an efficiency of 5.8 ±
0.7%, in agreement with previous work showing that this lesion is a block to DNA
replication (31, 32, 39). The bypass efficiencies of the lesions were also determined in
cells with the SOS system induced. Under these conditions, the bypass efficiency of Iz
did not change significantly (71 ± 7%); however, the bypass efficiency of NI increased a
remarkable 8-fold to 57 ± 1%. The bypass efficiency of the THF lesion increased to 30 ±
2%, demonstrating that the SOS system was indeed induced.
Mutation type and frequency. The mutational signature of each lesion in WT AB1157 E.
coli was determined using the restriction endonuclease and post-labeling analysis of
mutation frequency (REAP) assay (Figure 3.5). Insertion and deletion mutations which
do not compromise the BbsI recognition site or the PCR primer sites are detectable by the
REAP assay during the PAGE purification step and were negligible in this work (for Iz,
no detectable frameshifts were observed; for NI, 0.5-1% −1 frameshifts were observed,
data not shown).
In normal cells, the mutation frequency of Iz was 91% versus 50% for NI (Figure
3.6). The mutations induced by Iz were essentially all GÆC mutations, as predicted by
molecular orbital calculations and in vitro primer extension experiments (40, 41). NI
135
Chapter 3 – Iz and NI are Mutagenic in Vivo
caused 8.9% ± 0.5% GÆC mutations, and a roughly equivalent amount of GÆA and
GÆT mutations (19% ± 2% and 22 ± 3%, respectively).
The mutation type and
frequency for each lesion was also determined in SOS-induced cells to investigate the
effect of the SOS response system on the coding properties of these lesions.
Interestingly, Iz and NI had somewhat lower overall mutation frequencies of 84% and
33%, respectively. The coding properties of Iz were more degenerate under SOS-induced
conditions. Whereas Iz caused 2.0 ± 0.1% GÆA and 1.1 ± 0.2% GÆT mutations
without SOS induced, the lesion now induced 3.4 ± 0.5% GÆA and 5.5 ± 0.8% GÆT
mutations. The mutational signature of NI also differed in SOS-induced cells, as the
lesion induced many fewer GÆA and GÆC mutations (13 ± 2% and 2.5 ± 0.6%,
respectively).
136
Chapter 3 – Iz and NI are Mutagenic in Vivo
3.5. Discussion
In order to assess the biological fate of Iz and NI in DNA, we have determined the
relative bypass efficiency and the mutation type and frequency of each lesion in vivo.
The Iz derivative is a major and ubiquitous in vitro product of guanine oxidation. In
addition to forming as a result of ONOO− oxidation (28), this lesion also forms when
guanine is oxidized by other reactants including hydroxyl radical (42), MnTMPyp/KHSO5 (43-45), and riboflavin (photooxidation) (40-42, 46). Direct formation
of Iz from 8-oxoG has also been observed (40, 47). On a biological timescale, Iz is stable
(t1/2 in single-stranded DNA ≈ 16 h at 37 °C) (33), but can hydrolyze to form an
oxazolone derivative.
Oxazolone is bypassed efficiently and exhibits a mutation
frequency similar to that of Iz in vivo (86%), but preferentially generates GÆT
transversion mutations (31). In distinct contrast, NI has been observed only as a result of
ONOO−-related chemistry (28, 48, 49). NI is a chemically stable lesion (when heated to
90 °C for 6 h at pH 7, ~10-15% decomposition of an ODN containing NI occurs) (49)
and is refractory to repair by Fpg glycosylase (MutM) and endonuclease III in vitro (48).
The chemical and biological properties of NI make this lesion a potential biomarker of
ONOO−-mediated oxidation of DNA (28, 48).
In vitro studies have been conducted with Iz and NI, and the results suggest they
would be both replication blocks and mutagenic in vivo. Ab initio studies of Iz in vacuo
and using a self-consistent reaction field method of solvation indicate this lesion should
give GÆC mutations because of its apparent ability to mimic the hydrogen bonding face
of C and to form a stable base pair with G (40, 41). Replication of the lesion using DNA
polymerase I (pol I), and the Klenow fragment of pol I (Kf), showed that the lesion
indeed codes exclusively as a cytosine, but is inefficiently bypassed (40). In contrast, the
site-specific bacteriophage replication experiments presented here showed that the lesion
was bypassed relatively efficiently and had a mutation frequency of 91%, suggesting one
or more polymerases in addition to pol I may be responsible for the bypass of Iz under
normal conditions.
137
Chapter 3 – Iz and NI are Mutagenic in Vivo
The bypass and coding properties of DNA containing NI were previously studied
using the purified DNA polymerases Kf (exo−), calf thymus polymerase α (pol α), and
human polymerase β (pol β) (48). All three polymerases could insert a nucleotide
opposite the lesion, and although a kinetic analysis was not performed, the data suggested
that extension of the NI-containing base pair by Kf (exo−) and pol α is rate limiting and
results in poor translesion synthesis.
The nucleotides A and G were incorporated
opposite NI by Kf (exo−) and pol α, with Kf (exo−) favoring insertion of C opposite the
lesion. Interestingly, pol β efficiently bypassed the lesion and incorporated mostly C,
rendering the lesion non-toxic and non-mutagenic. Our results differ from the in vitro
data since substantial amounts of T were incorporated opposite NI, thereby producing
GÆA mutations in vivo.
This observation indicates pol I is probably not solely
responsible for the bypass of NI since this polymerase did not incorporate T opposite the
lesion in vitro. The bypass efficiency of NI in WT cells shows that NI is a significant
block to replication and has a bypass efficiency similar to that of the THF synthetic
abasic site.
This observation correlates with our recent thermal melting study that
showed NI greatly destabilizes duplex DNA regardless of the identity of the base paired
opposite the lesion. To explain this observation, we proposed that NI adopts a nonplanar
and potentially DNA distorting conformation (34). Thus, the low bypass efficiency of NI
may result from the inability of NI to fit within the tight active site of a replicative DNA
polymerase (50).
Alternatively, the large increase in NI bypass efficiency when
replicated in SOS-induced cells may indicate the involvement of Y-family DNA lesion
bypass polymerases, which have less sterically restrictive active sites and can bypass
numerous types of lesions (51-58).
The differences in bypass efficiency and coding specificity of the lesions in
normal versus SOS-induced cells indicate that SOS-induced proteins contribute to the
processing of these lesions.
In normal cells, the number of SOS-inducible DNA
polymerase molecules per cell is approximately 50 for pol II (59), 250 for pol IV (60),
and fewer than 15 for pol V (61). Upon SOS induction, the number of DNA polymerase
molecules per cell increases to about 350 for pol II (59), 2500 for pol IV (60), and 200 for
pol V (61). It has been demonstrated that one or more of these polymerases are required
138
Chapter 3 – Iz and NI are Mutagenic in Vivo
to bypass certain DNA lesions, and the specific polymerases involved vary depending on
the sequence context in which the lesion resides (62-64). Experiments using polymerase
deficient cell lines would address which of the SOS-induced polymerases are involved in
the bypass of Iz and NI.
The mutational signatures of Iz and NI determined here using site-specifically
modified genomes correlate well with previous cell-based mutagenesis studies (19, 24).
In one report, a pSP189 shuttle vector containing the supF gene was treated with ONOO−
in the presence of NaHCO3 (a physiological buffer component that facilitates the reaction
of ONOO− with DNA by supplying CO2), followed by replication in MBL50 E. coli (24).
Essentially all mutations occurred at G-C base pairs and consisted of 55% GÆC, 31%
GÆT, and 11% GÆA mutations.
We previously showed in vivo that the 8-oxoG
secondary oxidation products cyanuric acid, oxaluric acid, and oxazolone give
exclusively GÆT mutations (31), whereas guanidinohydantoin generates exclusively
GÆC mutations and the spiroiminodihydantoin diastereomers yield a mixture of GÆC
and GÆT mutations (32). In both studies, 8-oxoG was slightly mutagenic and induced
only
GÆT
mutations,
so
the
discovery
that
guanidinohydantoin
and
the
spiroiminodihydantoin diastereomers generate GÆC mutations implicated these lesions
as the possible sources of the in vivo GÆC mutations found in the supF/pSP189 system.
The results in the present work implicate Iz as an additional in vivo source of GÆC
mutations. Importantly, prior to this work no ONOO−-derived lesions had been found to
give significant amounts of GÆA mutations. Our results implicate NI as a possible
source of the GÆA mutations observed during the supF/pSP189 study.
Iz and NI are formed directly from guanine upon oxidation by ONOO−. When
replicated in vivo, these lesions are bypassed and potently mutagenic, collectively
inducing GÆC, GÆT, and GÆA mutations. Our results support a role for Iz and NI as
contributors to the mutational spectrum observed when ONOO−-treated DNA is
replicated in vivo. If these lesions persist in cells, they may be powerful sources of
mutations.
139
Chapter 3 – Iz and NI are Mutagenic in Vivo
3.6. Acknowledgment
James C. Delaney (MIT), Paul T. Henderson (Lawrence Livermore National
Laboratory), and John M. Essigmann contributed to this work. We thank Dr. Yuriy O.
Alekseyev for helpful discussions.
We also acknowledge the MIT Biological
Engineering Division Mass Spectrometry Laboratory and the Department of Chemistry
Instrumentation Facility. Financial support was provided by NIH awards CA26731 and
CA080024 and by NSF awards DBI-9729592 and CHE-9808061.
140
Chapter 3 – Iz and NI are Mutagenic in Vivo
Figure 3.1. Primary products of ONOO− oxidation and nitration of guanine.
O
H
N
HN
H2N
O
O
N
dR
N
H2N
8-oxoG
H2N
H2N
O
N
NH
dR
N
NO2
N
dR
8-NO2-G
O
N
HN
N
N
HN
N
G
Iz
N
dR
N
NO2
NH
N
dR
N
H
NI
141
NH2
Chapter 3 – Iz and NI are Mutagenic in Vivo
Figure 3.2. Construction of genomes containing a guanine residue or DNA lesion at a
specific site. A 5’-phosphorylated 19mer oligodeoxynucleotide insert containing a
guanine residue or DNA lesion at a specific site (denoted by a solid circle) is positioned
within a ss WT M13mp7L2 bacteriophage genome using two DNA scaffolds. The 19mer
insert is ligated into the genome using T4 DNA ligase, and the scaffolds are digested
using the exonuclease activity of T4 DNA polymerase to produce the genome construct.
EcoRI Site
EcoRI
Guanine
or Lesion
Anneal Scaffolds
and Modified 19mer
WT M13mp7L2
Phage DNA
142
1. Ligate
2. Digest
Scaffolds
Genome
Construct
Chapter 3 – Iz and NI are Mutagenic in Vivo
Figure 3.3. Assay for determining the efficiency of DNA polymerase bypass of a DNA
lesion. Genome constructs containing a site-specific guanine residue (G) or DNA lesion
(X) are separately mixed with a known amount of internal standard (ss WT M13mp7L2
genome), and the equally proportioned mixtures are used to transform E. coli cells.
Progeny phage (denoted by a circle within a hexagon) derived from a genome construct
have a functional lacZ gene, whereas phage containing the WT M13mp7L2 genome do
not. Thus, the composition of the progeny phage mixtures can be assayed by plating on a
lawn of NR9050 indicator E. coli in the presence of IPTG and X-gal and counting the
number of blue and clear plaques produced by the phage infections.
G
X
+
+
G-Control Internal Standard
(WT M13mp7L2)
Lesion
Internal Standard
(WT M13mp7L2)
Transform E. coli
G
N
+
G-Control Internal Standard
Yields Blue
Yields Clear
Plaques
Plaques
+
Internal Standard
Lesion
Yields Clear
Yields Blue
Plaques
Plaques
Plate on lawn of
indicator E. coli
# of Blue Plaques = B(G)
# of Clear Plaques = C(G)
# of Blue Plaques = B(L)
# of Clear Plaques = C(L)
Relative Bypass Efficiency =
143
B(L) x C(G) x 100%
B(G) x C(L)
Chapter 3 – Iz and NI are Mutagenic in Vivo
Figure 3.4. Bypass efficiency of Iz, NI, and a THF abasic site relative to G. The results
shown are the average of three experiments, with the experimental error given as a 95%
confidence interval of the mean.
Bypass Efficiency / %
Normal Conditions
SOS-Induced Conditions
100
100
80
80
60
60
40
40
20
20
0
G
Iz
NI
THF
144
0
G
Iz
NI
THF
Chapter 3 – Iz and NI are Mutagenic in Vivo
Figure 3.5. REAP assay for determining the mutation type and frequency of a DNA
lesion. E. coli cells are transformed with genome constructs site-specifically modified
with a guanine residue or DNA lesion. Position N in the resultant progeny phage
genomes represents the site formerly occupied by the site-specific modification. The
region containing N is PCR amplified to yield a 101mer containing BbsI and HaeIII
recognition sites. The BbsI digestion product is radiolabeled and digested with HaeIII to
produce a 19mer with N radiolabeled. PAGE purification of the 19mer also allows
insertion and deletion mutations (appearing as ODNs of longer or shorter length than the
19mer) to be quantified, if present. Digestion of the 19mer with nuclease P1 and
subsequent partitioning of the nucleotide mixture by TLC allows the point mutation type
and frequency to be quantified using phosphorimagery.
BbsI Site HaeIII Site
N
PCR
5’
3’
GAAGACCG NAGC
CTTCTGGC YTCG
101mer
Progeny Phage
1. BbsI, Phosphatase
2. [γ- 32P]-ATP, Kinase 32P-NAGC
19mer
3. HaeIII
4. PAGE
3’
5’
3’ Nuclease 32
P-dNMP
TLC
Elution
C
G
A
Origin
Mutation Frequency =
C + T + A x 100%
C+T+A+G
145
Chapter 3 – Iz and NI are Mutagenic in Vivo
Figure 3.6. REAP assay results for G, Iz, and NI. A. Representative polyethyleneimine TLC analysis of point mutation type and
frequency. B. Quantification of the TLC data for G (top row), Iz (center row), and NI (bottom row). The graphical data shown are
the average of three experiments, with the experimental error given as a 95% confidence interval of the mean.
A
Normal Conditions
SOS-Induced Conditions
B
Normal Conditions
100
C
T
G
40
40
20
20
100
80
A
G
T
C
Iz
0
100
80
60
60
40
40
20
20
0
80
A
G
T
C
NI
0
100
80
60
60
40
40
20
20
0
G
Iz NI Marker
G Iz NI Marker
Base in M13 Genome Before Replication
80
60
100
A
G
60
0
Coding Specificity / %
Base Found at Lesion Site After Replication
80
SOS-Induced Conditions
100
A
G
T
C
0
G
A
G
T
C
G
T
C
G
T
C
Iz
A
NI
A
Base Found at Lesion Site After Replication
146
Chapter 3 – Iz and NI are Mutagenic in Vivo
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mutagenesis in E. coli: various combinations of translesion polymerases
(Pol II, IV and V) deal with lesion/sequence context diversity. DNA
Repair (Amst) 1, 159-167.
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CHAPTER 4
Urea Lesion Formation in DNA as a Consequence of 7,8Dihydro-8-oxoguanine Oxidation and Hydrolysis Provides
a Potent Source of Point Mutations
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
4.1. Abstract
The DNA oxidation product 7,8-dihydro-8-oxoguanine (8-oxoG) forms several
mutagenic oxidation products, including a metastable oxaluric acid (Oa) derivative. We
report here that a synthetic oligonucleotide containing Oa hydrolyzes under simulated “in
vivo” conditions to form a mutagenic urea (Ua) lesion.
Using the Oa 2'-
deoxyribonucleoside as a model, the hydrolysis rate depended strongly upon the
concentrations of bicarbonate and divalent magnesium. In buffered solutions containing
physiologically relevant levels of these species, the half-life of Oa nucleoside was
approximately 40 h at 37 °C. The mutagenic properties of Ua in DNA were investigated
using an M13mp7L2 bacteriophage genome containing Ua at a specific site.
Transfection of the lesion-containing genome into wild-type AB1157 Escherichia coli
allowed determination of the mutation frequency and DNA polymerase bypass efficiency
from the resulting progeny phage. Ua was bypassed with an efficiency of 11% compared
to a guanine control and caused a 99% GÆT mutation frequency, assuming the lesion
originated from G, which is at least an order of magnitude higher than the mutation
frequency of 8-oxoG under the same conditions.
SOS induction of bypass DNA
polymerase(s) in the bacteria prior to transfection caused the mutation frequency and type
to shift to 43% GÆT, 46% GÆC and 10% GÆA mutations. We suggest that Ua is
instructional, meaning that the shape of the lesion and its interactions with DNA
polymerases influence which nucleotide is inserted opposite the lesion during replication,
and that the instructional nature of the lesion is modulated by the size of the binding
pocket of the DNA polymerase. Replication past Ua, when formed by hydrolysis of the
8-oxoG oxidation product Oa, denotes a pathway that nearly quantitatively generates
point mutations in vivo.
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
4.2. Introduction
DNA damage mediated by reactive oxygen and nitrogen species is believed to
contribute to the etiology of human cancer, aging, and neurological disorders (1-3).
Examining the mutagenicity and repair of DNA oxidation products and providing a
structural basis for these effects are of fundamental importance for understanding their
genotoxic properties. The lesion 7,8-dihydro-8-oxoguanine (8-oxoG) is regarded as
perhaps the most critical product resulting from oxidative DNA damage due to its high
steady-state concentration in cells and a capability to mispair with A, which leads to
G→T transversion mutations (4-9). Recent studies in this laboratory and others have
shown that 8-oxoG is an unstable species under a variety of oxidative conditions, owing
to its low oxidation potential compared to the parent 2'-deoxyguanosine (10). A variety
of DNA lesions is formed when 8-oxoG is exposed to oxidative conditions (11-21). The
product distribution depends on the mechanism of action of the oxidant and the reaction
conditions. For example, peroxynitrite (ONOO−), a powerful oxidant that is formed in
vivo by the combination of nitric oxide with superoxide (22-24), reacts preferentially with
8-oxoG compared to the four canonical nucleobases to form several lesions including
cyanuric acid (Ca), imidazolone (Iz) and its hydrolysis product oxazalone (Oz),
guanidinohydantoin (Gh), spiroiminodihydantoin diastereomers (Sp1 and Sp2), and
oxaluric acid (Oa) (2). The distribution of the 8-oxoG oxidation products is strongly
dependent on the buffer composition and the concentration of ONOO−. For example, Oa,
Oz, and Ca are predominantly formed in the presence of a molar excess of ONOO− and
25 mM sodium bicarbonate in phosphate buffer (pH = 7.2) (20). In contrast, when
ONOO− is slowly infused over time the lesions Gh and Sp predominantly form (25). We
recently reported that these secondary lesions are an order of magnitude more mutagenic
than 8-oxoG, and that they are well-bypassed by DNA polymerase during replication in
E. coli (26-28) (Table 4.1).
In the course of our studies, we observed that Oa decomposes in aqueous solution
to form a urea (Ua) derivative, a DNA base lesion that is usually ascribed to the presence
of pyrimidine oxidation.
To our knowledge, the formation of Ua during 8-oxoG
oxidation has been reported only once (29); however, in that study a 1,3-diamine, such as
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
spermine or 1,3-diaminopropane, was required for the production of Ua. Herein, we
describe the hydrolysis of Oa to Ua in 2'-deoxynucleoside and oligonucleotide models at
physiologically relevant pH and salt concentrations (Scheme 4.1). The effect of normal
and SOS-induced cellular conditions on the mutagenic potential of Oa and Ua in DNA
was determined in an M13 bacteriophage system using the recently developed restriction
endonuclease and postlabeling (REAP) assay (Figure 4.1) (30-32). The hydrolysis and
mutation results support a novel potential pathway to mutagenesis in vivo from the
oxidation of 8-oxoG.
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
4.3. Experimental Procedures
Chemicals and Enzymes. Restriction endonucleases, T4 polynucleotide kinase, and T4
DNA ligase were purchased from New England Biolabs. M13mp7L2 was a gift from C.
W. Lawrence (University of Rochester). The E. coli cell strain used for transfection was
AB1157, and for plating was NR9050 from R.M. Schaaper (NIEHS). The E. coli strain
used for re-growth of phage was SCS110 from Stratagene. [γ-32P]ATP (6000 Ci/mmol)
was from New England Nuclear, and ATP was from Pharmacia.
Acetonitrile, β-
mercaptoethanol, and ammonium acetate were from Aldrich and ammonium phosphate
dibasic was from Mallinckrodt (AR grade). Polyethyleneimine coated TLC plates and
phosphoric acid were from J.T. Baker.
Sephadex G-25 spin columns were from
Boehringer Mannheim.
Nucleosides. Treatment of 3',5'-di-O-Ac-8-oxoGuo with ONOO− followed by HPLC
purification allowed isolation of the Oa nucleoside derivative, as previously described
(33).
Hydrolysis of Oa. The decomposition of Oa was followed using a coupled enzymatic
spectrophotometric oxalate assay (Sigma-Aldrich or Trinity Biotech), which allowed
detection of oxalate in solution through the production of an indamine dye. The oxalate
kit consisted of two reagents, A (active ingredients: 3 mM 3-(dimethylamino)-benzoic
acid and 0.22 mM 3-methyl-2-benzothiazolinone hydrazone, pH 3.1) and B (active
ingredients: 3000 U/L barley oxalate oxidase and 100,000 U/L horseradish peroxidase).
The following general procedure was used for the decomposition reactions: Oa (1.5
nmol) in 30 µL of buffer solution was incubated at 37 °C for a prescribed period of time.
The buffer solution was pH 7.4 and consisted of 150 mM potassium phosphate and,
where indicated, 5 mM magnesium sulfate and/or 20 mM sodium bicarbonate. After the
incubation, 30 µL of a 2X solution of reagent A was added, followed by 12 µL of reagent
B. The solution was mixed, incubated at room temperature for 15 minutes, and the A590
of the resulting indamine dye was determined by spectrophotometry.
The initial
concentration of Oa was adjusted such that complete reaction to form oxalate resulted in
an absorbance of 1.0 in a 1 cm path length cell.
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Electrospray Ionization Mass Spectrometry. Oa 3',5'-di-O-Ac-2'-deoxynucleoside (0.2
mg, 0.6 µmol) in 12 µL of buffer solution was incubated at 37 °C for one day. The buffer
solution was pH 7.4 and consisted of 150 mM potassium phosphate plus 5 mM
magnesium sulfate and 20 mM sodium bicarbonate.
The 3',5'-di-O-Ac-2'-
deoxynucleosides were purified by HPLC using a 250 × 4.6 mm, Hypersil ODS column
(Thermo Electron Corporation, Bellefonte, PA) (Figure 4.2). The mobile phases used
were 60 mM aqueous potassium phosphate, pH 7.4 (A) and 100% acetonitrile (B).
HPLC conditions used were: isocratic 5% B for 10 min, then 5% B to 27.4% B over 20
min. The flow rate was 1 mL/min, and products were monitored simultaneously at 213
nm and 260 nm. A Quattro MicroTM API mass spectrometer (Micromass UK Limited,
Manchester, UK) was used for electrospray ionization (ESI)-MS experiments, and spectra
were obtained in positive ion mode using a 49:40:10:1 water/methanol/acetonitrile/
ammonium hydroxide or 78:20:2 water/2-propanol/ammonium hydroxide spraying
solution (Figure 4.2).
Oligonucleotides.
Oligonucleotides were from Research Genetics (Huntsville, AL),
Integrated DNA Technologies (Coraville, IA), or synthesized on an Applied Biosystems
model 391 DNA synthesizer.
Oligonucleotides were deprotected with concentrated
NH4OH for 18 h at 55 °C. The strand containing 8-oxoG, to prevent degradation of the
lesion, required 0.25 M β-mercaptoethanol in the deprotection solution. Oligonucleotides
made by solid-phase synthesis were purified by PAGE and C18 reverse-phase HPLC
before use (~99% purity) as described (26). MALDI-TOF mass-spectrometry verified
the molecular weight of each strand after purification. The insert sequence was 5'-GCG
AAG ACC GXA GCG TCC G-3' (X = G, 8-oxoG, Oa, Ua, or a tetrahydrofuran (THF)
synthetic AP site).
−
−
ONOO Treatment of 8-OxoG in DNA. ONOO was prepared by ozonation of sodium
azide in alkaline solution (0.1 M NaOH) and stored at –80 °C (34). Concentrations of
−
ONOO were determined by spectrophotometry (ε = 1670 M-1 cm-1, λ = 302 nm in 0.1 M
158
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
−
NaOH). Reactions between ONOO and oligonucleotides were performed in buffer
containing 150 mM potassium phosphate and 25 mM sodium bicarbonate (pH = 7.2), as
described (20). The purified Oa-containing oligonucleotide was hydrolyzed to Ua under
well-defined conditions as described above for the Oa nucleoside.
HPLC Purification of Modified Oligonucleotides.
The Oa- and Ua-containing
oligonucleotides were purified by HPLC using a 250 mm x 2.1 mm, 5 µm Supelco
Supelcosil LC-18 column with UV detection at 260 nm. The mobile phase was 150 mM
aqueous ammonium acetate (A) and 100% acetonitrile (B) with a gradient of 7-12% B
over 30 min at a flow rate of 0.25 mL/min.
The quantities of each product were
calculated by UV absorbance at 260 nm using the molar absorptivity of the G-containing
oligomer (ε = 184700 M-1 cm-1) for all the modified oligonucleotides. The mass of each
oligonucleotide was determined using MALDI-TOF mass spectrometry with Oa [MW =
5843.8 (calculated), 5844.7 (observed)] and Ua [MW = 5771.8 (calculated), 5771.5
(observed)] resulting in the expected mass.
The mass of each oligonucleotide was
calibrated using oligonucleotides that bracketed the molecular ion of each analyte, which
allowed the measured mass to fall routinely within 1 atomic mass unit of the calculated
mass for each lesion-containing oligonucleotide.
Although a few percent of Ua
contamination in the Oa sample was unavoidable, sample handling during the REAP
assay either minimized or avoided additional Ua formation (26).
Since there are
relatively few products that form from 8-oxoG oxidation by ONOO−, and these products
are generally well-separated by HPLC purification, we are confident that the
oligonucleotides were 95-99% pure.
Construction of M13 Genomes Containing a Site-Specific Lesion. Single-stranded (ss)
M13 genomes containing a unique lesion were generated as previously described (Figure
4.1A) (26). Briefly, 1 pmol of ss M13mp7L2 was linearized by EcoRI and annealed to
equimolar
amounts
of
two
oligonucleotide
“scaffolds,”
which
are
partially
complementary to the 5'- and 3'-ends of the insert and the linearized genome. An
equimolar amount of the 5'-phosphorylated 19-mer insert was added and covalently
joined into the genome by incubation with T4 DNA ligase (16 °C, 1 h). The scaffolds
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
were removed by heating (50 ºC, 5 min) in the presence of a 100-fold molar excess of
“anti-scaffold” complement immediately prior to transfection.
Transfection and determination of translesion bypass efficiency. Cells, grown to an
OD600 = 0.4 in 100 mL of yeast tryptone (1X) medium (35), were made competent for
transfection by the calcium chloride method as described previously (26, 36). For SOS
induction of bypass DNA polymerases, the cells were irradiated with 254 nm light, at an
intensity of 45 J/m2 and transferred to an equivalent volume of 2×YT media and grown
for 40 min at 37 °C before being made competent. Lesion bypass was determined by
transfection with 5 ng of M13 DNA, and mutation frequency was determined using 50 ng
of M13 DNA. The number of independent transformed cells was determined by adding
2.5 mL B-broth soft agar to the transfected cells and plating onto a lawn of NR9050 E.
coli. The number of plaques on each plate was counted after incubation at 37 °C for 16 h.
As a control to estimate the number of viable viral genomes that do not have an insert, a
wild-type (WT) M13mp7L2 sample was exposed to the genome construction conditions,
and the average number of resulting plaques was subtracted from the total formed by
each insert-containing genomic construct. Transfections were performed in triplicate,
and the bypass efficiency relative to the G control was calculated to within a 95%
confidence interval of the mean for each lesion. A THF AP-site was used as a DNA
polymerase blocking control, which provided significantly fewer plaques than the Gcontaining insert control.
Mutation Frequency Determination.
The point mutations caused by each lesion in
AB1157 E. coli were determined and quantified using the restriction endonuclease and
post-labeling (REAP) assay as previously described (Figure 4.1B) (26).
Briefly,
transformed cells for determination of mutation frequency were added to 10 mL of LB
media and incubated on a roller drum for 5 h at 37 °C (growth), followed by pelleting of
the cells and decanting of the phage-containing supernatant into new 15 mL
polypropylene tubes. A 100 µL aliquot of the suspension was added to 9 mL of LB and 1
mL of mid-log SCS110 E. coli. The cells were incubated for 4 h (regrowth), pelleted,
and the phage-containing supernatant was decanted and stored at 4 °C. The growth and
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
regrowth steps served to amplify the amount of progeny phage while reducing the
relative amount of non-transfected DNA to a negligible amount (<0.1%). The mutation
frequencies were determined by performing the REAP assay on a DNA duplex produced
by PCR amplification of the insert region of the progeny phage. Each 101-mer PCR
product was cleaved by BbsI at the position in the template vector that originally
contained the lesion, affording a 55-mer whose newly formed 5'-end contained the lesion
site.
The 55-mer was dephosphorylated with shrimp alkaline phosphatase, which
exposed the lesion site at the 5'-OH for radiolabeling with PNK and [γ-32P]ATP.
Incubation with HaeIII, to yield a 19-mer, allowed PAGE purification of the radiolabeled
fragment of interest.
Digestion of the desalted 19-mer to 5'-deoxynucleotide
monophosphates (5'-dNMPs) with snake venom phosphodiesterase (SVPD) was followed
by partitioning of the mixture on a polyethyleneimine TLC plate developed in saturated
(NH4)2HPO4 (adjusted to pH = 6.1 with H3PO4). The separated radiolabeled nucleotides
were quantified by PhosphorImager analysis, which provided the point mutation
frequency and type at the lesion site.
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
4.4. Results and Discussion
4.4.1. Hydrolysis of Oa to form Ua
We investigated the hydrolytic stability and mutagenic potential of the 8-oxoG-derived
products, Oa and Ua. The specific oxidation of an 8-oxoG-containing oligonucleotide
with ONOO− followed by HPLC purification of the modified oligonucleotide was used to
obtain pure Oa. The lesion Ua was observed to form via divalent metal cation and base
catalyzed hydrolysis (Scheme 4.1), and the identity of the lesion was established from
MALDI-TOF mass spectral data as previously reported (20, 26). Incubation of Oacontaining oligonucleotide in phosphate buffered saline (pH = 7.4) at 37 °C caused no
discernable conversion to Ua (data not shown). However, addition of Mg2+, Ca2+, and
HCO3− to the solution at concentrations that are isotonic to mammalian cells, incubation
in alkaline solution, or incubation in pure water at high temperature caused appreciable
hydrolysis of the Oa oligonucleotide to form a new species, which was assigned as Ua
(Figure 4.3). The decrease in mass by 72 amu corresponds to the expected loss of oxalate
during Ua formation.
As a model system to study conveniently the hydrolysis reaction under a variety
of conditions, the 3'-5'-di-O-acetyl-2′-deoxynucleoside of Oa was synthesized from 3',5'di-O-acetyl-2'-deoxy-8-oxoguanosine (26, 33).
The Oa nucleoside was exposed to
HCO3− and to various divalent alkaline earth metal cations in order to determine if these
species can catalyze hydrolysis to Ua at or near in vivo concentrations. Exposure of the
Oa nucleoside at 37 °C to a phosphate buffered solution containing 5 mM MgSO4 and 20
mM NaHCO3 (pH = 7.4) resulted in almost complete hydrolysis of the nucleoside (halflife ~40 h), as determined by a commercially available oxalate assay (see Materials and
Methods), whereas exposure of the Oa nucleoside to similar conditions without a divalent
metal cation (half-life ~52 h) and without HCO3− (half-life ~96 h) resulted in much
slower hydrolysis of the nucleoside starting material, with the phosphate buffered control
hydrolyzing at the slowest rate (half-life ~130 h) (Figure 4.4).
The addition of 5 mM MgSO4 to an aqueous potassium phosphate buffered
solution (pH 7.4) increased the rate of hydrolysis of the Oa nucleoside (Figure 4.4),
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
implying that Mg2+ stabilizes the rate-determining transition state for the reaction. A
previous study showed that oxalate binds Mg2+ effectively (37), and Falvello et al.
recently demonstrated that the divalent metal cations Co2+, Ni2+, and Cu2+ can form a
complex with the oxalate moiety of oxalurate (38). Thus, it is plausible that Oa also
chelates Mg2+. The binding of a divalent metal cation to Oa could increase the rate of
hydrolysis by several mechanisms, either alone or in concert. By binding to the oxalate
portion of Oa, electron density could be reduced at the carbonyl of the amide, thereby
facilitating nucleophilic addition at the amide carbon. The developing negative charge on
the amide carbonyl oxygen during nucleophilic addition would also be stabilized by the
presence of Mg2+. Divalent metal cations could also stabilize the electrostatic repulsion
between the carboxylate of Oa and a negatively charged incoming nucleophile (HO− or
HCO3−). Alternatively, chelation of a divalent metal cation by the oxalate moiety could
facilitate the elimination of this group in a manner similar to the commonly known Mg2+assisted elimination of inorganic phosphate or pyrophosphate during the hydrolysis of
nucleotide triphosphates.
Surprisingly, HCO3− significantly increased the rate of hydrolysis of the Oa
nucleoside (Figure 4.4).
During the hydrolysis by water, a gem-diol tetrahedral
intermediate is presumed to form at the amide carbon. The spontaneous hydrolysis of Oa
may be explained through a comparison of the leaving group pKa values. The pKa of urea
is approximately 14 (39), whereas the pKa of water is 15.7. Assuming a minimal effect
on the Ua pKa due to the 2-deoxyribose substituent, the reaction should favor elimination
of Ua. If instead of hydroxide, HCO3− adds to the amide carbonyl of Oa, forming a
tetrahedral intermediate, fragmentation would not occur directly from this structure.
Collapse of the tetrahedral structure at this point should eliminate HCO3− (pKa H2CO3 =
6.35) about 107 fold more frequently than Ua, making the fragmentation of Oa an
unlikely event. However, the tetrahedral intermediate may follow a different reaction
coordinate. The pKa of HCO3− is 10.3; however esterification of HCO3− to produce
carbonic acid monomethyl ester (CH3OCOOH) results in a significant decrease of the
pKa of the acid to 2.92 (40). Upon addition of HCO3− to Oa, it is likely that a large
163
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
decrease of the carbonic acid pKa also results. Deprotonation of the carbonic acid could
conceivably initiate a decarboxylation event and subsequent fragmentation of Oa.
During the reaction of Oa with HCO3−, several factors may contribute to the
hydrolysis rate increase. We propose that the reaction proceeds by the addition of HCO3−
to Oa followed by decarboxylation and fragmentation (Scheme 4.2). Decarboxylation
and fragmentation may occur in a concerted fashion or in several separate steps.
Herschlag and Jencks investigated a Mg2+-catalyzed reaction in which carboxylate ions
react with phosphorylated pyridines in a nucleophilic fashion (41). They found that
HCO3− reacts much faster than CO32−, even though CO32− is more basic, because HCO3−
hydrogen bonds to a phosphoryl oxygen in the transition state, thereby avoiding
unfavorable electrostatic interactions. They also note that Mg2+ may serve as a template
for the reacting species. In the present case, intramolecular hydrogen bonding may
contribute to the ability of HCO3− to react with Oa. Given the stabilizing role Mg2+
appears to play during the hydrolysis of Oa in the absence of HCO3−, it may also play a
similar role in the addition of HCO3− to Oa.
4.4.2. In vivo mutagenesis by Oa and Ua under normal or SOS-induced conditions
The mutagenic potential of Ua was tested in vivo by inserting oligonucleotides
containing each defined lesion (8-oxoG, Oa, or Ua) into M13mp7L2 single-stranded viral
genomes and then transfecting the genomes into wild-type E. coli (Figure 4.1). Each
lesion was bypassed during replication in vivo efficiently enough to produce progeny
phage for the determination of point mutation frequency and type using the REAP assay.
Since lesions in DNA may block or inhibit replication, the relative number of
plaques formed from the immediate plating of the transformation mixture gives a
measure of the efficiency of translesion DNA synthesis. A comparison of the number of
plaques obtained after transformation of the site-specific lesion-containing phage genome
with that from an identically constructed vector that did not carry any oxidative damage
allowed calculation of the relative bypass efficiency (26, 27). As shown in Figure 4.5,
vector survival for Oa is 51 ± 3%, which is in agreement with a previously published
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
value (26). The Ua lesion bypass efficiency was 11 ± 5%, which is significantly higher
that that observed for a synthetic THF AP site employed as a control for DNA
polymerase blocking and SOS-induced bypass (26, 27, 42, 43). The bypass efficiencies
of the lesions were also determined in cells with the SOS system induced. Significant
differences were not observed compared to uninduced cells using the assay described in
Figure 4.1A, as Oa and Ua were bypassed with efficiencies of 56 ± 9% and 18 ± 4%,
respectively.
However, increased levels of insert-containing DNA after PCR
amplification during the REAP protocol were observed for Ua and THF (Figure 4.6),
which is consistent with an increased bypass efficiency. The error inherent in the bypass
assay does not allow small increases in bypass efficiency to be observed with statistical
significance. The inefficient bypass of Ua is in accord with previous studies (44).
Successful induction of the SOS operon is supported by the change in the mutation
specificity of Ua (see below) and the increased levels of insert-containing DNA after
PCR amplification of Ua and the THF lesion compared to uninduced controls during the
REAP assay (Figure 4.6).
As shown in Figure 4.7, in uninduced cells the mutation frequency and type for
both Oa and Ua was 99% with nearly all of the mutations in the form of GÆT
transversions (based on the lesions originating from G).
The mutation type and
frequency for each lesion was also determined in SOS-induced cells in order to
investigate the effect of inducible error-prone translesion DNA polymerase bypass on the
coding properties of these lesions. Compared with normal cells, SOS induction caused
Oa to have a somewhat similar mutation frequency and type, with 95% GÆT and 4%
GÆC transversions. In contrast, the more blocking Ua had a dramatically different
distribution of mutation types with 43% GÆT, 46% GÆC, and 10% GÆA mutations
and no measurable repair or correct coding. The observed insertion of G and A opposite
Ua, regardless of SOS induction, is in general agreement with previous in vitro and in
vivo studies (45-47). For example, Maccabee et al. reported that Ua produced from T
oxidation directed insertion of G and T opposite the lesion with a frequency of 62% and
26%, respectively, compared to other point mutations (45). Direct comparison of these
results to the REAP data is difficult because of sequence context variations and because
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Ua was not introduced site-specifically into the DNA, which does not allow repair or
correct coding to be scored. Studies in vitro demonstrate that Ua is excised by several
DNA glycosylases involved in base excision repair in E. coli such as Endo III, Endo VIII,
and their homologues in higher organisms (48). Additionally, another DNA glycosylase,
MutM, also known as Fpg, removes Oa in duplex DNA when the lesion is situated
opposite a C (20).
The increase in the degeneracy of the type of nucleoside inserted opposite Ua is
consistent with the larger, more flexible binding pocket generally found in Y-family
lesion-bypass polymerases (49). This binding pocket may accommodate atypical steric
configurations and unfavorable electrostatic interactions induced by the lesion and allow
promiscuous chain extension past the damaged DNA. Frameshifts and small deletions
are assumed to be negligible in this work, since there was no detectable variation in the
size of the BbsI/HaeIII restriction fragment (Figure 4.6). Clearly, the presence of Ua, like
the precursor Oa, causes a higher mutation frequency in DNA than the parent 8-oxoG.
Since well-bypassed lesions (i.e., ~50% or greater relative to guanine) have
essentially no change in bypass efficiency or mutation profile with SOS induction, they
are likely to be successfully replicated in E. coli regardless of SOS status. The change in
coding specificity observed for Ua during SOS induction is consistent with recent models
describing the initiation of a “handoff” from the replicative DNA polymerase to an errorprone bypass DNA polymerase upon encountering a lesion that stalls the replication fork
(49). Well-bypassed lesions include not only Oa, but also Ca, Iz, Oz, and Gh (Table 4.1).
The Ua lesion exists as a mixture of α- and β-anomers, so it is intriguing to hypothesize
the α-anomer may block DNA polymerase extension until β-anomerization permits
limited translesion bypass. SOS induction likely allows error-prone DNA polymerases to
bypass the lesion, perhaps regardless of the anomeric state of the sugar.
Further
experiments are needed to elucidate the structural basis of this possible effect. NMR
analysis of a duplex dodecamer containing Ua showed evidence of both anomers when
placed opposite T, and that the β-anomer was interhelical (aligned with the base stack)
while sharing at least one hydrogen bond with the opposing nucleotide (50).
166
This
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
hydrogen bonding interaction indicates that the lesion may be instructional, rather than
simply forcing the polymerase to obey the “A” rule, which states that A will be most
likely incorporated opposite a noninstructional lesion (51). A recent paper by Kroeger et
al. (52) discusses application of the REAP assay to 2'-deoxyribonolactone, an AP site
analog, and proposes that even limited hydrogen bonding contributes to the lesion being
instructional. We suggest that dA incorporation opposite Ua is, in part, attributable to
hydrogen bonding between the carbonyl and/or distal hydrogen of the urea moiety and
the respective N6-amino and/or N1 of dA. This explanation of Ua as an instructional
lesion is speculative, and alternative hydrogen bonding patterns cannot be ruled out,
particularly for SOS induced lesion bypass. Another explanation that is consistent with a
recent report by Kobayashi et al. is that nucleotide insertion opposite Ua is inhibited,
resulting in slippage and insertion of a second nucleotide opposite the base on the 3'-side
of the template Ua (53). However, the local sequence used in our work [5'-G(Ua)A-3']
only allows for GÆA mutations to occur by this mechanism, and these mutations are
only observed to a modest degree during SOS induction compared to the predominant
GÆT and GÆC mutations.
The 8-oxoG derived DNA lesions Oa and Ua are mutagenic under normal and
SOS induced conditions in vivo. If these lesions form in cells, they clearly represent a
threat to genomic integrity in the absence of an effective DNA repair mechanism.
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
4.5. Acknowledgment
Paul T. Henderson (Lawrence Livermore National Laboratory) and I were the
lead authors of this work and contributed equally. James C. Delaney (MIT), Feng Gu
(MIT), Jacquine C. Niles (MIT), Sang Soo Hah (Lawrence Livermore National
Laboratory), Steven R. Tannenbaum (MIT), and John M. Essigmann (MIT) also
contributed. We thank Dr. Kaushik Mitra for helpful discussions on the mechanism of
Oa hydrolysis. This work was supported by the National Institutes of Health (CA86489,
ES07020 and CA26731) and by the California Breast Cancer Research Program (9KB0179). This work was performed, in part, under the auspices of the U. S. Department of
Energy by the University of California, Lawrence Livermore National Laboratory under
Contract No. W-7405-Eng-48.
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Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.1. (A) Strategy for determination of translesion DNA polymerase bypass of
base damage in vivo. A site-specifically modified oligonucleotide is ligated into ss viral
M13 DNA.
Transfection into host cells followed by translesion DNA synthesis
ultimately affords infective progeny phage whose initial numbers are limited by the
lethality of the site-specific lesion. Plating of the transformation mixture onto a lawn of
indicator bacteria caused the formation of plaques (spots of infected cells), whose
numbers allowed for the calculation of the efficiency of polymerase bypass when
compared with the non-lesion control. (B) Detection of mutations using the REAP assay.
PCR amplification of the region from the progeny phage that had originally contained the
site-specific lesion was performed for each transfection. The resulting duplex DNA was
cleaved with BbsI, dephosphorylated, radiolabeled at the 5'-end, and reduced in size with
HaeIII to a 19-mer. Digestion of the radiolabeled and purified 19-mer to 5'-dNMPs,
separation by TLC, and quantification of the radioactivity by PhosphorImager analysis
allowed measurement of the fractional composition of each base to within 1%.
A. Lesion Bypass Efficiency Determination
Initial Number of Progeny
Corresponds to DNA Polymerase
Bypass Efficiency
Lesion
or Guanine
Control
M13 Genome
Construction
Oligonucleotide
Insert
N
Transform E. coli
M13 ss DNA with
Site-Specific
Modification
Progeny Phage Pool
(N = Mutation, Repair
or Correct Pairing)
B. Restriction Endonuclease and Postlabeling (REAP) Assay
N
1.PCR
2.Cleave at N with BbsI and Radiolabel (* = 32P)
3.Reduce Length with HaeIII
Progeny Phage Pool
(N = Mutation, Repair
or Correct Pairing)
Enzyme
Digest
dCMP
Elution
Mutation Frequency =
T+C+A
x 100%
T+C+A+G
*
5'-N
3'
Radiolabeled
Oligonucleotide
(19-mer)
dTMP
dGMP
dAMP
origin
169
Thin-Layer
Chromatography
*
5'-dNMP
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.2. ESI mass spectra and HPLC traces (insets) of 8-oxoG, Oa, and Ua 3',5'-di-OAc-2'-deoxynucleosides.
Abs at 260 nm
100
50
0
15
30
Time (min)
0
100
100
% Intensity
Oa
367.3 (calc.)
367.9 (found)
50
200
300
400
Mass (m/z)
Abs at 213 nm
% Intensity
8-oxoG
500
332.3 (calc.)
333.0 (found)
0
15
30
Time (min)
0
100
200
300
350
100
% Intensity
Ua
50
Abs at 213 nm
Mass (m/z)
260.2 (calc.)
261.0 (found)
0
15
30
Time (min)
0
100
200
300
Mass (m/z)
170
400
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.3. Hydrolysis of Oa to form Ua in an oligonucleotide. (Left) HPLC
chromatogram of the Oa-containing 19-mer oligonucleotide after heating in neat H2O for
20 min at 80 °C. (Right) MALDI-TOF mass spectra of the purified products.
5603.7
50 Standard
0.8
0.6
0
5300 5500
0.4
100
0.2
50
0
20
Ua
% Intensity
100
% Intensity
Absorbance (AU)
1.0
25
30
35
171
5771.5
5700
5900
Mass (m/z)
Oa
5100
6300
5844.7
5603.7
Standard
0
5300 5500
Retention Time (minutes)
6150.0
Standard
6150.0
Standard
5700
5900
Mass (m/z)
5100
6300
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.4. Oxalate formation curves for the decomposition of Oa 2'-deoxynucleoside at
37 °C in 150 mM potassium phosphate buffer, pH 7.4. Where noted, the buffer also
contained 5 mM MgSO4 and/or 20 mM NaHCO3. The error bars represent a 95%
confidence interval of the mean for three independent experiments.
Absorbance (AU)
1.0
Buffer + MgSO4 + NaHCO3
Buffer + NaHCO3
Buffer + MgSO4
Buffer Only
0.8
0.6
0.4
0.2
0.0
0
20
40
60
80
Time (h)
172
100 120
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.5. Relative bypass efficiency for Oa and Ua. Controls include G (wellbypassed) and a synthetic THF AP site (blocking). The error bars represent a 95%
confidence interval of the mean for three independent experiments.
Relative Bypass (%)
In Vivo DNA Polymerase Bypass
= - SOS
100
= + SOS
75
50
25
0
G
8-oxoG
Oa
173
Ua
AP(THF)
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.6. SOS induction allows preferential bypass of blocking lesions in vivo. PCR
amplification of the insert region in M13 progeny phage, as described for the REAP
assay in the text, yielded a binary distribution of DNA fragments; one contained the 19mer insert that had originally borne a site-specific lesion (or a G or T control nucleotide)
and another that is derived from insert-free phage DNA. Products were assayed by
agarose gel electrophoresis, which included an unmodified (WT) phage control and a no
template control (-phage). The band corresponding to a fragment ~80 bases long likely
results from deletion of the hairpin in WT M13 DNA during PCR (54, 55). Additional
controls were grouped according to bypass efficiency:
G, T, and 8-oxoG (well-
bypassed); Oa, Oz, and Ca (moderately-bypassed); whereas THF and Ua are nearly
blocking. The ratio of the two PCR products varied according to the bypass efficiency of
the site-specific lesion contained in the insert region.
THF and Ua were the most
blocking lesions, as determined by the plaque counting assay described in the text and
showed the greatest increase in insert-containing PCR product after SOS induction by
exposure to UV light. The insert-containing PCR products were digested with BbsI, 32Pradiolabeled, and digested with HaeIII to yield 19-mer oligonucleotides, which were
purified by PAGE. Note that in both cases, the signal intensity of the bands in the THF
and Ua lanes on the polyacrylamide gel correlates with findings from the agarose gel.
Insert: G
T
8-oxoG Oa
Oz
Ca
THF
Ua
115-mer
-phage WT control
101-mer (+insert)
~80-mer (-insert)
Radiolabeled
19-mers from
the REAP assay
SOS Induction
(45 J/m2 at 254 nm)
3% Agarose, 100V, 4h
THF
20% PAGE
174
Ua
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Figure 4.7. TLC showing mutation frequency and type as determined by the REAP
assay.
Markers consisting of 5'-dNMPs were derived from digestion of a 13-mer
oligonucleotide that contained a degenerate 5'-terminus.
N = 8-oxoG
SOS Induction:
- +
Oa
- +
Ua
- +
dNMPs
dCMP
Elution
dTMP
dGMP
dAMP
Mutation (%)
G → T:
7.0; 6.8
<1; 1.3
G → C:
<1; <1
G → A:
Origin
99; 95
<1; 3.8
<1; <1
175
99; 43
<1; 46
<1; 10
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Scheme 4.1. Oa hydrolyzes to from Ua and oxalate.
O
HN
R
O
N
H
O
O
Mg2+ + HCO3
(catalysts)
H2O
O
HN
R
O
NH2
O
urea
(Ua)
oxalurate
(Oa)
AcO
O
R=
or
AcO
176
O
O
DNA
oxalate
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Scheme 4.2. Proposed mechanism for Mg2+ and HCO3− catalyzed hydrolysis of Oa.
H
O
O
HN
Mg
O
N
H
2+
HO
O
HN
N
H
O
O
OH
O
H
2+
O O Mg
O
O
HN
Mg2+
O
NH2
O
O
O
O
H
O
177
CO2
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
Table 4.1. Summary of bypass efficiencies and mutation frequencies for 8-oxoG
oxidation products in wild-type E. coli.
a
property
bypass efficiency (%)
GÆT mutations (%)
GÆC mutations (%)
GÆA mutations (%)
(26).
b
(28).
c
(27).
Ca a
65 ± 8
97
<1
<1
d
Izb
60 ± 5
1
88
2
8-oxoG oxidation product
Oz a
Gh c
Sp1 c
Sp2 c
57 ± 13 75 ± 5
9±3
9±4
86
1.4
27
41
<1
98
72
57
<1
<1
<1
<1
This work.
178
Oa d
51 ± 3
99
<1
<1
Ua d
11 ± 5
99
<1
<1
Chapter 4 – Mutagenic Urea Lesion Formation From 8-OxoG Oxidation
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183
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184
CHAPTER 5
The Hydantoin Lesions Formed From Oxidation of 7,8Dihydro-8-oxoguanine are Potent Sources of Replication
Errors in Vivo
185
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.1. Abstract
Single-stranded DNA genomes have been constructed that site-specifically
contain
the
7,8-dihydro-8-oxo-2'-deoxyguanine
(8-oxoG)
oxidation
products
guanidinohydantoin (Gh) and the two stable stereoisomers of spiroiminodihydantoin (Sp1
and Sp2).
The circular viral genomes were transfected into wild-type AB1157
Escherichia coli and the efficiency of lesion bypass by DNA polymerase(s) was assessed.
Viral progeny were analyzed for mutation frequency and type using the recently
developed restriction endonuclease and post-labeling (REAP) assay. Gh was bypassed
nearly as efficiently as the parent 8-oxoG but was highly mutagenic, causing almost
exclusive GÆC transversions. The stereoisomers Sp1 and Sp2 were, in comparison,
much stronger blocks to DNA polymerase extension, and caused a mixture of GÆT and
GÆC transversions. The ratio of GÆT to GÆC mutations for each Sp lesion was
dependent on the stereochemical configuration. All observed mutation frequencies were
at least an order of magnitude higher than those caused by 8-oxoG. Were these lesions to
be formed in vivo, our data show that they are absolutely miscoding and may be
refractory to repair after translesion synthesis.
186
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.2. Introduction
Genotoxicity due to degradation of DNA by oxygen radicals is associated with
aging and carcinogenesis (1).
An understanding of the processes of spontaneous,
oxidant-induced and ionizing radiation-induced mutagenesis requires knowledge of the
exact types and biological significance of oxidative lesions formed in DNA (2). Both
endogenous and exogenous agents have been implicated in the in vivo oxidation of DNA.
The observation that oxidative damage to DNA naturally occurs in vivo was a major
discovery because it demonstrated that DNA damage could occur endogenously as a
consequence of normal metabolism (3-5). Currently, over 50 oxidative DNA lesions
have been characterized (6). The 7,8-dihydro-8-oxo-2'-deoxyguanine (8-oxoG, Figure
5.1) lesion has been studied extensively because it is a major oxidative DNA lesion (7)
and can be detected by a variety of assays (8). 8-oxoG is mutagenic because of its
propensity to mispair with A during replication by DNA polymerases. Incorporation of
A opposite 8-oxoG leads to a predominant G→T transversion mutation (9-11), which is
the second most common somatic mutation in human carcinomas (12), and is especially
frequent in the mutational spectrum of the p53 tumor suppressor gene (13).
Several studies have shown that 8-oxoG, which has a lower oxidation potential
than any of the four normal DNA bases (14), is highly reactive towards oxidizing
reagents. This suggests that 8-oxoG may also be sensitive to in vivo DNA oxidation.
−
Peroxynitrite (ONOO ), which is formed endogenously by the reaction of nitric oxide
with superoxide, is a potent oxidant that decomposes into radical species capable of
oxidizing DNA and other cellular constituents including thiols, lipids and proteins (1517). Oxidation of DNA is known to have mutagenic consequences. For example,
−
ONOO treatment of plasmid DNA containing the supF gene, followed by transfection
into mammalian or bacterial cells, induces mainly G→T, and some G→C transversions
−
(18). Reaction of ONOO with an oligonucleotide containing a portion of the supF gene
followed by incubation with hot piperidine, which cleaves DNA preferentially at oxidized
8-oxoG nucleotides, causes strand scission at sites coinciding with the mutational “hot
spots” observed after replication in bacterial cells (19). Since 8-oxoG is not a known
187
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
source of G→C transversions and 8-oxoG in DNA is refractory to cleavage by piperidine
(20, 21), these experiments suggest that additional DNA lesions are forming and that
−
these may contribute to ONOO mutagenicity.
Peroxynitrite is known to react preferentially with 8-oxoG in DNA to form
several
mutagenic 2'-deoxy-β-D-erythro-pentofuranosyl products, including cyanuric
acid, oxaluric acid and oxazalone (22), which were recently reported to cause
predominantly G→T transversion mutations in vitro using purified DNA polymerases
(23-25), and in vivo by transfection of site-specifically adducted bacteriophage DNA into
Escherichia coli (26). Determination of the in vivo mutagenic consequences of such
lesions typically have taken the following course:
(i) synthesis of the modified
oligonucleotide containing the DNA adduct of interest, (ii) insertion of the
oligonucleotide into the genome of a virus or plasmid, (iii) introduction of the sitespecifically modified genome into a bacterial or mammalian host where replication, and
possibly repair, occur, and (iv) enumeration and characterization of mutant progeny (27).
Several 8-oxoG oxidation products have yet to be analyzed for their in vivo
mutagenic potential. Recently, two major products of 8-oxoG oxidation were identified:
guanidinohydantoin (Gh) and spiroiminodihydantoin (Sp) (28-30), both of which exist as
stereoisomers as shown in Figure 5.1. Although these lesions can be made by oxidation
−
of 8-oxoG with ONOO (mimicking the chemistry of the in vivo inflammatory response)
(31) or by oxidation of guanine with singlet oxygen or other oxidants, a difficult to
separate mixture of products results when the reaction is carried out using
oligonucleotides (unpublished data).
synthesis,
Burrows
and
co-workers
Using a temperature-dependent method of
developed
selective
methods
that
yield
oligonucleotides containing Gh and Sp via oxidation of 8-oxoG using the one-electron
oxidant Na2IrCl6, a compound whose redox potential is specific for 8-oxoG as compared
with the four normal DNA bases (21, 32-34).
188
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
In view of the reported susceptibility of 8-oxoG to oxidation, the mutagenic
potential of Gh and Sp was tested in vivo by inserting oligonucleotides containing each
well-defined lesion into bacteriophage DNA, followed by transfection into wild-type E.
coli. This report describes that each lesion was bypassed efficiently enough during
replication in vivo to yield progeny phage. The viral progeny was analyzed for mutation
frequency and specificity, and it was shown that these oxidation products were much
more mutagenic than the parent 8-oxoG.
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Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.3. Experimental Procedures
Oligonucleotides. The DNA base lesion 8-oxoG and tetrahydrofuran abasic (AP) site
control 19mer oligonucleotides were made using phosphoramidite solid-phase methods
and purified as described (26).
Oligodeoxynucleotides containing Gh or Sp were
prepared by reacting 12 µM 5'-GCGAAGACCGXAGCGTCCG-3' (X = 8-oxoG) with the
oxidizing agent Na2IrC16 (175 µM final concentration) in 100 µL of 10 mM sodium
phosphate and 100 mM NaCl for 1 h. Selective formation of Gh was accomplished by
incubation of the 8-oxoG-containing oligonucleotide at pH 7.0 and 4 °C, while
incubation at pH 6.5 and 60 °C afforded the Sp lesion. The reaction mixture was
dialyzed against H2O using 2000 molecular weight cutoff dialysis tubing for 24 h. The
samples were analyzed by negative ion electrospray mass spectrometry (Micromass
Quattro II), and their purity was estimated to be ~95% on the basis of the intensities of
related molecular ions.
Although both lesions were formed as a mixture of
stereoisomers, the Sp-containing oligonucleotides were separable by anion-exchange
HPLC. The pure stereoisomer-containing strands are termed Sp1 and Sp2, although their
absolute
stereochemistry
has
not
been
assigned.
Purification
of
the
Gh
oligodeoxynucleotide was accomplished by HPLC using a Dionex DNA Pac PA-100 4 X
250mm column and an isocratic buffer system consisting of 50% solvent A (10%
acetonitrile and 90% H2O) and 50% solvent B [10% acetonitrile and 90% 1.5 M
ammonium acetate (pH 7)] (Figure 5.2). The flow rate was 1.0 mL/min and UV spectra
were recorded at 260 nm.
Oligodeoxynucleotides containing the Sp isomers were
purified in a similar manner, except that an isocratic buffer system consisting of 30%
solvent A and 70% solvent B was used (Figure 5.3). Prior to use, the HPLC purified
oligodeoxynucleotides were dialyzed against H2O for 72 h, followed by dialysis against 1
mM NaCl for 72 h, to minimize the possibility of polynucleotide kinase inhibition by
ammonium cations.
The purity of the HPLC-purified oligodeoxynucleotides was
determined by HPLC to be >99% using a Dionex DNA Pac PA-100 4 X 250mm column
and a linear gradient of 35% solvent B to 100% solvent B in 30 min (Figures 5.2 and
5.3). MALDI-TOF mass spectrometry was used to verify the molecular weight (MW) of
each strand after purification [Gh, MW = 5868.9 (calculated), 5869.2 (observed); Sp1 and
Sp2, MW = 5894.9 (calculated), 5895.7 and 5895.7 (observed), respectively] (Figures
190
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.4-5.6). The synthetic AP site 19mer control and a 20mer oligonucleotide were used to
calibrate the mass spectrometer, which allowed measurement of the MW to within 1
atomic mass unit of the calculated value. The stability of the lesions to the genome
construction protocol was verified by MALDI-TOF mass spectrometry.
Lesion-
containing oligonucleotides were exposed to all reagents and conditions, with the
omission of viral DNA and T4 polynucleotide kinase.
After the incubations, mass
spectrograms of the Gh, Sp1, and Sp2 were essentially identical to those of the purified
strands prior to genome construction (Figures 5.4-5.6).
DNA Polymerase Bypass Efficiency. Lesion bypass experiments in wild-type AB1157 E.
coli were performed as described (26, 35) in triplicate using genomes made from the
purified oligodeoxynucleotides 5'-GCGAAGACCGXAGCGTCCG-3' (X is the lesion or
guanine for the unmodified insert control), as shown in Figure 5.7. Briefly, 1 pmol of ss
M13mp7L2 was linearized by EcoRI digestion of a hairpin contained in the M13
polylinker region that interrupts the lacZ gene. Equimolar amounts of two “scaffolds”
(oligonucleotides that are complementary to and span the 5'- and 3'-ends of both the
vector and the lesion-containing insert) were annealed to the genome. Two scaffolds
were used in order to leave a ss gap around the lesion that allowed the subsequent ligation
efficiencies to be equal, regardless of the lesion employed. An equimolar amount of the
5'-phosphorylated 19mer insert was added and covalently joined into the genome by
incubation with T4 DNA ligase (16 °C, 2 h). The ligation efficiencies were confirmed to
be equal by electrophoresis of an aliquot of the ligation mixture on a 1% agarose gel,
staining of the gel with ethidium bromide, and image analysis with digital photography
(data not shown). AB1157 E. coli were made chemically competent with CaCl2 and
immediately plated after transfection using 20 ng of ss viral DNA and ~ 108 cells as
described (26). For each successful initial round of viral genome replication within each
cell, viral progeny leaves a transfected AB1157 cell and infect nearby NR9050 E. coli
(containing the F' episome required for infection). The resulting localized infection slows
the growth of the NR9050 cells, which is visible as a transparent spot (plaque)
surrounded by a dense lawn of uninfected cells. A control for the presence of viable
genomes that lacked an insert was made by constructing a genome containing a
191
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
nonphosphorylated AP-containing 19mer insert. The number of plaques arising from this
control was subtracted from the number of plaques derived from genomes containing 5'phosphorylated inserts; the latter inserts contained the lesion of interest. The number of
plaques formed by each lesion relative to that of the guanine control genome is indicated
as “relative bypass efficiency”, with the corrected guanine reference averaging 217
plaques per plate, as shown in Table 5.1.
Mutation Frequency and Type. Mutational analysis was performed using the recently
developed restriction endonuclease and postlabeling (REAP) assay with triplicate
genome constructions and transfections for each lesion (26, 35-37). Briefly, 100 µL of
cells was transformed by electroporation (2.5 kV, 129 ohms, 2mm gap cuvette) of 50 ng
(1 ng/µL) of the constructed genome, after which they were added to 10 mL of LB media
and incubated on a roller drum for 6 h at 37 °C. This amplified the ~104 initial events to
~109 progeny. The cells were pelleted, and the supernantant was decanted into 15 mL
polypropylene tubes and stored at 4 °C. To eliminate PCR artifacts due to the presence
of nontransfected DNA, 50 µL of the phage suspension was added to 9 mL of LB and 50
µL of saturated SCS110 E. coli. The cells were grown an additional 4 h producing ~3 ×
1011 phage, thus reducing the amount of nontransfected DNA template to less than 0.1%
with respect to the template from the progeny phage.
We believe this subsequent
infection will produce no change in the progeny population, since the genome was
modified with the lesion in a nonessential gene, and reconstruction experiments in which
progeny phage containing different mixtures of G and A at the lesion site retained their
initial mixture composition after subsequent infection (36).
Single-stranded viral
template DNA was prepared using 700 µL of phage supernatant (QIAPrep spin M13 kit,
Qiagen), from which 15% was used for each PCR reaction. As shown in Figure 5.8, the
region that had contained the lesion in each progeny phage sample was PCR amplified to
yield a 101 bp product whose purity was assayed by agarose gel electrophoresis. The
resulting DNA duplex was cleaved with BbsI (a type IIs restriction endonuclease that
cleaves a fixed numbers of bases away from its binding site) at the position that had
originally contained the lesion in the template vector, thus affording a 55mer whose
newly formed 5'-end contained the lesion site. The 55mer was dephosphorylated with
192
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
shrimp alkaline phosphatase, which allowed the lesion site at the 5'-overhang to be
radiolabeled using PNK and [γ-32P]ATP. Incubation with the restriction endonuclease
HaeIII yielded a 19mer, allowing for PAGE purification of the radiolabeled fragment of
interest. Digestion of the desalted 19mer to 5'-deoxynucleotide monophosphates (5'dNMPs) was followed by the partitioning of the mixture on a 20 x 20 cm
polyethyleneimine TLC plate developed in saturated (NH4)2HPO4 (adjusted to pH 6.1
with H3PO4). The separated radiolabeled nucleotides were quantified by PhosphorImager
analysis, thus providing the fractional base composition of each nucleotide at the lesion
site, from which the mutation frequency and type were determined.
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Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.4. Results
The mutagenic potential of DNA lesions is described by two key metrics: (1) the
ability of DNA polymerases to undergo translesion DNA synthesis (bypass efficiency)
and (2) the resulting mutation frequency and type. Since the viral DNA used in this study
was single-stranded, translesion DNA synthesis must have occurred in order for progeny
phage to form in the E. coli cells. A comparison of the number of plaques (slow growing
colonies of successfully transfected bacteria on a lawn of infectable cells) formed by viral
replication of lesion-bearing versus control genomes provided a measure of translesion
bypass by DNA polymerases in the cell. The REAP assay was used to determine the
identity of the base in the progeny DNA at the position that had contained the lesion,
which gave the mutation frequency and type.
Translesion Bypass Efficiency.
Since lesions in DNA may block or inhibit
replication, the relative number of plaques formed from the immediate plating of the
transformation mixture gives a measure of the efficiency of translesion DNA synthesis.
A comparison of the number of plaques obtained after transformation of the damaged
phage genome to that from an identically constructed vector that did not carry any
oxidative damage allowed calculation of the bypass efficiency for each lesion.
Transfections were performed in triplicate, and the bypass efficiency relative to the G
control was calculated to within a 95% confidence interval of the mean for each lesion.
As shown in Figure 5.7, vector survival for the 8-oxoG control was 87 ± 4% with respect
to the G control, which clearly demonstrated that 8-oxoG only weakly inhibited
replication.
This result agrees, within experimental error, with previously reported
values, both in E. coli and in mammalian cells (10, 11, 38-41). Synthesis past the
secondary oxidative lesion Gh was effective for 75 ± 5% of the vectors, which was nearly
the same as for 8-oxoG, and slightly less efficient than the G control. Sp1 and Sp2 were
clearly more blocking to DNA polymerase bypass than Gh, with survival percentages of
9 ± 3% and 9 ± 4%, respectively. All of the lesion-containing genomes afforded enough
progeny phage for mutation analysis (vide infra). As a control demonstrating DNA
replication blockage, lesion survival for a synthetic tetrahydrofuran AP site was 4 ± 2%,
which is somewhat higher than that reported for the natural aldehydic AP site placed site194
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
specifically in a nearly identical bacteriophage system (42). In contrast, the bypass
efficiency measured for the AP site is an order of magnitude lower than that reported for
transfection of a plasmid carrying the AP site in a gapped region into cultured human
cells (43), which may be due to ubiquitous expression of Y-family DNA error-prone
bypass polymerases in mammalian cells whose homologues are present at negligible
concentrations in uninduced E. coli (44). Since the 8-oxoG oxidation products are
appreciably bypassed (especially the Gh lesion) by polymerases in E. coli, they are
potential contributors to mutagenesis.
Mutation Frequency and Type. As shown in Figure 5.8, the mutation frequencies were
determined alongside an 8-oxoG control, with radiolabeled 5'-dNMPs as markers for the
relative migration of each nucleotide. The mutation frequency of 8-oxoG was modest,
providing a 3% GÆT transversion. In contrast to the low mutation frequency of 8-oxoG,
the mutation frequencies were essentially 100% for Gh, Sp1, and Sp2.
The well-
bypassed lesion Gh caused 98% GÆC and 2% GÆT mutations. Interestingly, the same
mutation frequencies were observed for the Sp stereoisomers, but the mutation types
were distributed differently, with GÆC being the predominant mutation for both Sp1 and
Sp2 stereoisomers. We observed 72% GÆC and 27% GÆT transversions for Sp1 and
57% GÆC and 41% GÆT transversions for Sp2.
195
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.5. Discussion
8-OxoG is one of the most common oxidative DNA base lesions due to the low
oxidation potential of guanine (14). The further oxidation of 8-oxoG after its formation
from guanine is reasonable, since the adduct has a lower oxidation potential than the four
normal nucleotides.
Additionally, base radical cations (the initial products of one-
electron oxidation of DNA bases) can migrate over long distances (>55 bp) in duplex
DNA to become trapped preferentially at 8-oxoG sites (45-48), resulting in oxidation of
the lesion (47). Oxidation of 8-oxoG results in the formation of the hydantoin derivatives
described in the present study, as well as a variety of products such as cyanuric acid,
oxaluric acid and a 2,5-diaminoimidazalone, which undergoes slow hydrolysis to form
oxazalone (49, 50). The hydantoin products may also be obtained directly from guanine
by use of four-electron oxidants such as singlet oxygen or type I photochemistry
involving superoxide (29, 49), which implicates them as general products of guanine
oxidation in DNA.
We investigated the mutagenic potential of three significant products of
peroxynitrite oxidized 8-oxoG in DNA: Gh, Sp1, and Sp2. Convenient synthesis of each
defined lesion was enabled by reaction of an 8-oxoG-containing oligonucleotide with the
oxidant Na2IrCl6 under conditions that favored nearly exclusive formation of either Gh or
Sp from 8-oxoG. Each lesion was placed site-specifically into a bacteriophage genome
and assayed for bypass efficiency and mutation type following transfection into E. coli.
A modest mutation frequency (3%) was observed for 8-oxoG, as expected on the basis of
literature values (10, 11, 26, 39, 41). The predominant mutation for this lesion was
G→T. It is known that 8-oxoG, placed site-specifically in a ss DNA vector, is subject to
repair in E. coli, since the mutation frequency increases approximately 10-fold in cells
deficient in the base-excision repair glycosylases MutM (which catalyzes cleavage of a
variety of oxidized guanine lesions from duplex DNA) and MutY (predominantly
removes A opposite 8-oxoG) (11, 51). Presumably, the repair occurs after the first round
of replication, since both MutM and MutY require duplex substrates (11, 52-54). In E.
coli, it appears that no mechanism exists to restore the original G⋅C pair from the
secondary oxidation products of 8-oxoG after translesion DNA synthesis, since there
196
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
were negligible amounts of G in the position of the bacteriophage that had contained Gh
or Sp, as measured by the REAP assay. Additionally, the lesions Gh, Sp1, and Sp2 are
well-bypassed by the E. coli DNA polymerase(s).
This combination of translesion
bypass and miscoding properties apparently contributes to potent mutagenicity of the
guanine oxidation products presented in this report. The ~100% mutation frequencies
observed may implicate these lesions collectively as a primary source for the frequently
observed GÆT and GÆC transversions caused by the endogenous oxidation of DNA (2,
13, 18, 51, 55).
It is not known if the sequence context surrounding the lesions influences the
mutagenesis of the adducts nor if nontargeted mutations can arise from Gh and Sp. Such
a sequence context effect on mutation frequency is observed for 8-oxoG (56, 57). Studies
of context effects on Gh and Sp mutagenesis and toxicity would be useful. In this regard,
the present work may shed light on the context-dependent mutagenesis induced by
methylene blue plus UV light in studies by Loeb and co-workers (58). They observed
frequent GÆC mutations under conditions now known to produce abundant amounts of
Sp (31).
Significant frequencies of small deletions or insertions would have been detected
by the REAP assay, since the BbsI-HaeIII cleavage products would vary detectably in
length. The homogeneity of the resulting 19mer, as assayed by PAGE, indicated that
point, rather than frame-shift, mutations predominated in the sequence context evaluated
(data not shown).
Lesions derived from 8-oxoG have yet to be detected in cells, but there is indirect
evidence that their formation in DNA is relevant in cell-based mutagenesis studies.
Juedes and Wogan investigated the mutagenicity of peroxynitrite using the supF gene of
the pSP189 shuttle vector as a mutation target in bacterial and mammalian cells (18).
−
Exposure to ONOO caused the mutation frequency to increase 21-fold in the vector
when replicated in E. coli MBL50 cells, and 9-fold upon replication in human Ad293
cells compared to untreated vector. In both systems, mutations occurred at G⋅C base
197
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
pairs, predominantly involving GÆT transversions (65% when replication was in
bacteria and 63% when in human cells). However, GÆC transversions were observed
but at a lower frequency (28% in MBL50 and 11% in Ad293 cells). In a related study,
Termini and co-workers reported that, upon transfection of peroxy radical exposed
bacteriophage DNA into E. coli (55), approximately 88% of the of the mutant progeny
consisted of nearly equal numbers of GÆT and GÆC transversions. The authors could
not detect 8-oxoG by HPLC/electrochemical analysis, suggesting that the GÆT
transversions were not caused by this base lesion. The work presented here provides at
least a partial explanation for the mutation spectrum observed in the supF and
bacteriophage mutagenesis studies. Other lesions that may be responsible for GÆC
transversions include the 2-amino-5-[(2'-deoxy-β-D-erythro-pento-furanosyl)amino]-4Himidazol-4-one derivative (Iz), which is formed from the oxidation of guanine and 8oxoG (59). Iz is known to pair preferentially with G during primer extension with Pol I
and, therefore, may be a contributor to G→C transversions in vivo (60).
Single nucleotide primer extension studies carried out using the Klenow fragment
(exo−) recently showed that both Gh- and Sp-containing DNA templates directed
insertion of dAMP and dGMP opposite the lesions (61). Both Gh and Sp showed nearly
the same pattern of insertion of nucleotides opposite the lesions, always with an
approximately 2:1 preference for dAMP over dGMP. It is noteworthy that, in vivo,
nearly exclusive dGMP incorporation occurs opposite Gh, while Sp lesions exhibit a
mixture of dAMP and dGMP insertion opposite the lesion, but with an opposite
preference of dGMP incorporation with respect to the in vitro primer extension work.
Perhaps there is an unknown repair pathway operating on Gh⋅A or Sp⋅A base pairs, since
recent studies in the David group have indicated that the adenine opposite the lesions is
not a substrate for MutY (34). An alternative explanation is that the replicative DNA
polymerase(s) in E. coli has (have) different dNTP incorporation preferences opposite Gh
and Sp than the Klenow fragment. It would be interesting to determine the molecular
basis for the dGMP nucleotide insertion preference for the hydantoin lesions with respect
to the many oxidative lesions such as Oa, Ca, and Oz, which prefer dAMP incorporation
(nearly exclusively for Oa and Ca) in vivo (26).
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Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Guanine oxidation results predominantly in the formation of 8-oxoG, Gh and Sp,
as well as a variety of other products. Indeed, oxidation of 8-oxoG has been shown to be
a chemically favorable event, and its oxidation products also include Gh and Sp. We
report that the hydantoin products Gh, Sp1, and Sp2 are much more mutagenic in vivo
than 8-oxoG, thus demonstrating that lesions arising from facile oxidation of dG and 8oxoG may contribute to the observed GÆT and GÆC transversions upon replication of
oxidized DNA in E. coli and mammalian cells.
199
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.6. Acknowledgment
Paul T. Henderson (MIT) was the lead author of this work. James C. Delaney
(MIT), James G. Muller (U. of Utah), Steven R. Tannenbaum (MIT), Cynthia J. Burrows
(U. of Utah), and John M. Essigmann (MIT) also contributed. Financial support was
provided by grants from the NIH (CA86489, CA80024, CA90689, ES07020, CA26731,
CA55861, and ES04705).
200
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.1. Structures of 8-oxoG, Gh, Sp1, Sp2 and the oligonucleotide sequence used
for genome construction.
O
H
N
NH
O
N
dR
O
NH
H2N
N
H
NH2
OO
OO
H
N
HN
O
N
dR
N
HN
NH
N N
dR
H2N
O
H2N
Sp1
+
Sp2
Gh
NH
N N
dR
X
5'-GCGAAGACCG
O
P O
O
O
O
AGCGTCCG-3'
X = 8-oxoG, Gh, Sp1, and Sp2
19mer insert
201
O
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.2. HPLC chromatogram for crude (A) vs. purified (B) oligodeoxynucleotide 5’GCG AAG ACC GXA GCG TCC G-3’ (X = Gh).
202
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.3. HPLC chromatogram for crude (A) vs. purified (B) oligodeoxynucleotide 5’GCG AAG ACC GXA GCG TCC G-3’ (X = Sp1 or Sp2).
203
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.4. MALDI-TOF mass spectra for purified (A) vs. exposure to mock genome
construction conditions (B) of oligodeoxynucleotide 5’-GCG AAG ACC GXA GCG
TCC G-3’ (X = Gh).
204
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.5. MALDI-TOF mass spectra for purified (A) vs. exposure to mock genome
construction conditions (B) of oligodeoxynucleotide 5’-GCG AAG ACC GXA GCG
TCC G-3’ (X = Sp1).
205
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.6. MALDI-TOF mass spectra for purified (A) vs. exposure to mock genome
construction conditions (B) of oligodeoxynucleotide 5’-GCG AAG ACC GXA GCG
TCC G-3’ (X = Sp2).
206
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.7. (A) Strategy for determination of in vivo translesion DNA polymerase bypass of site-specific lesions in a ss bacteriophage
DNA (M13mp7L2). A site-specifically adducted ss viral DNA was transfected into the host cell, followed by translesion DNA
synthesis to afford a ds replicative form intermediate. Expression of gene products from the ds intermediate enabled the formation of
infective progeny phage. Plating of the transformation mixture onto a lawn of generic bacteria immediately after transfection caused
formation of plaques, whose number allowed for the calculation of the efficiency of polymerase bypass for each lesion relative to a
guanine control. The base composition (N) at the site that had contained the lesion prior to transfection gave the mutation frequency
and type. (B) Relative bypass efficiency data for Gh, Sp1, and Sp2. Controls for DNA polymerase bypass efficiency include G and 8oxoG (well-bypassed) and a synthetic THF AP site (blocking). The error bars represent a 95% confidence interval of the mean based
N
Relative Bypass Efficiency (%)
on individual transfections from three independently constructed genomes.
N
N
N
N
N
100
80
60
40
20
0
N
N
N
207
G
8-oxoG
Gh
Sp1
Sp2 AP(THF)
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Figure 5.8. (A) Detection of mutations using the REAP assay. (B) TLC showing mutation frequency analysis by the REAP assay for
Gh, Sp1, and Sp2. Nucleotide insertion frequencies were determined in triplicate experiments, except for Sp2, which was performed
in duplicate. The 8-oxoG lesion was included as a control of known mutation frequency and type, and 5'-dNMP markers were derived
from digestion of a 13-mer oligonucleotide that contained a radiolabeled degenerate 5'-end.
N
Progeny Phage
Mutant Pool
B N
1. BbsI/
BbsI site
Phosphatase
(cleaves 5' to N) HaeIII site
32
2. *[ P]-ATP/
Kinase
5'
N
3'
PCR
3. HaeIII
Duplex DNA
4. PAGE
(101 bp)
Base Mutation Frequency
T, C, or A
x 100%
T+C+A+G
Elution
dCMP
Frequency (% ± s.d.)
C:
–
98(0.2) 72(0.9) 57(1)
T: 2.7(0.1) 1.4(0.2) 27(1) 41(2)
G: 97(0.3)
–
–
–
A:
–
–
–
–
*
5'-N
3'
Radiolabeled
19-mer
Enzyme
Digest
dTMP
dGMP
dAMP
Thin-Layer
Chromatography
dCMP
*
5'-dNMP
Mutation
dTMP
Elution
A
dGMP
dAMP
8-oxoG
208
Gh
Sp1
Sp2
Markers
Repair or
Correct
Coding
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
Table 5.1. Bypass data for G, Gh, Sp1, Sp2, and AP. The number of plaques per plate
produced from a cut control (using a nonphosphorylated AP-containing insert that cannot
ligate) was subtracted from each measurement to correct for the presence of noninsertcontaining circular ss DNA). The assay was performed in triplicate (one construct was
transfected into three pools of cells). The G-control was defined as 100% bypass. The
square root of the sum of the variances for the lesion (or G control) and the cut control
divided by the average number of plaques for the G control was used to calculate the
standard deviation.
Ga
Gb
Gc
plaques
250
243
258
8-oxoGa
8-oxoGb
8-oxoGc
220
228
216
Gha
Ghb
Ghc
198
205
188
Sp1a
Sp1b
Sp1c
57
52
48
Sp2a
Sp2b
Sp2c
61
54
45
APa
APb
APc
40
45
38
cuta
cutb
cutc
34
36
30
sum
avg
sd
avg-cut
avg-cut
sd
%MAX
overall
95% C.I.
751
250.3
7.5
217.0
8.1
100
4
664
221.3
6.1
188.0
6.8
87
4
591
197.0
8.5
163.7
9.1
75
5
157
52.3
4.5
19.0
5.4
9
3
160
53.3
8.0
20.0
8.6
9
4
123
41.0
3.6
7.7
4.7
4
2
100
33.3
3.1
0.0
4.3
0.0
2
209
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
5.7. References
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Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
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spiroiminodihydantoin and guanidinohydantoin and DNA synthesis past
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212
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
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214
Chapter 5 – The Hydantoin Lesions Produce G-to-T and G-to-C Transversions in Vivo
lack of involvement of 8-oxo-dG1 and/or abasic site formation.
Biochemistry 37, 7030-7038.
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215
CHAPTER 6
Effect of Escherichia coli Polymerases II, IV, and V on the
Bypass Efficiencies and Mutational Signatures of Guanine
Oxidation Products
216
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
6.1. Abstract
The role of the E. coli SOS inducible polymerases II, IV, and V in the translesion
synthesis of guanine oxidation products in vivo was determined. M13 viral genomes
containing
7,8,-dihydro-8-oxoguanine
(8-oxoG),
guanidinohydantoin
(Gh),
spiroiminodihydantoin (Sp) diastereomers, oxaluric acid (Oa), urea (Ua), and 5guanidino-4-nitroimidazole (NI) were constructed and replicated in wild-type, pol II
deficient, pol IV deficient, pol V deficient, and pol II / pol IV / pol V deficient AB1157
E. coli. The bypass efficiency and mutational properties of each lesion was determined in
each strain under uninduced and SOS induced conditions. In uninduced wild-type cells,
Gh, Sp, Ua, and NI were poorly bypassed. They were well-bypassed, however, in SOS
induced wild-type cells. Elimination of pol V completely abrogated the increase in
bypass efficiency upon SOS induction demonstrating the importance of this polymerase
in alleviating the genotoxicity of these lesions. Removal of pol II and pol IV activity
only affected the bypass efficiency of Sp1. With the exception of 8-oxoG, all of the
lesions studied were potent sources of mutations in all strains. Pol V had a strong effect
on the identity of the bases inserted opposite Sp2, Ua, and NI. Additionally, pol II and
pol V were shown to be involved in the translesion replication of NI, and this lesion was
the only damage product for which multiple polymerases were involved in translesion
synthesis.
217
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
6.2. Introduction
Induction of the SOS system in E. coli occurs in response to DNA damage and
causes the upregulation of at least 43 genes (1). Three of the protein products of these
genes have been identified as DNA polymerases and are known as polB (pol II) (2, 3),
dinB (pol IV) (4), and umuD2’C (pol V) (5-7). Extensive research on the structure and
function of these polymerases and their homologues suggests that these polymerases
allow replication to progress in the presence of DNA lesions that are strongly inhibitory
to the replicative polymerase, pol III.
Pol II is a B-family DNA polymerase and possesses a 3’Æ5’ exonuclease activity.
Consequently, this enzyme replicates normal DNA with high fidelity and has an error rate
less than 10−6 (8). Pol II participates in error-free replication restart (9, 10) and can also
carry out translesion synthesis of lesions such as abasic sites (2, 11), 3,N4-ethenocytosine
(12), and N-2-acetylaminofluorene (13). Pol IV and pol V are members of the Y-family
of DNA polymerases, which have less sterically restrictive active sites than normal
replicative polymerases (14, 15) and lack exonuclease activity (16, 17). The error rates
of these polymerases are much higher than the other E. coli polymerases and are on the
order of 10−3 to 10−4 for pol V (18, 19) and 10−4 to 10−5 for pol IV (19) in vitro. Pol IV
does not appear to affect the chromosomal mutation rate but rather operates mostly on
extrachromosomal DNA (20) and functions in adaptive mutagenesis by inducing −1
frameshifts (21). Pol IV can also participate in translesion synthesis but does so in a pol
V dependent manner (22, 23). Pol V is the major translesion synthesis polymerase (24)
and has been shown to bypass lesions such as abasic sites and UV photoproducts of DNA
(5, 19, 25) and also is involved in untargeted mutagenesis (18).
In an effort to explore potential mechanisms of carcinogenesis as a result of
inflammation, we have examined the role of the SOS-inducible E. coli polymerases in the
bypass of DNA lesions generated by peroxynitrite (ONOO−) oxidation of DNA.
Activation of the immune response causes the secretion of nitric oxide (•NO) and
superoxide (O2•−) by macrophages (26) and neutrophils (27) (two types of inflammatory
218
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
cells). These radicals combine in a diffusion-limited reaction to form ONOO− (28), a
powerful oxidizing and nitrating agent capable of damaging DNA (29). Possessing the
lowest redox potential of the four DNA nucleobases (E7 = 1.27 V vs. NHE) (30), guanine
(G) is preferentially oxidized by agents such as ONOO− as compared to the other natural
nucleobases (31). A variety of DNA oxidation products results from guanine including
guanidinohydantoin (Gh), spiroiminodihydantoin diastereomers (Sp1 and Sp2), cyanuric
acid (Ca), oxaluric acid (Oa), urea (Ua), 2-aminoimidazolone (Iz), oxazalone (Z), and 5guanidino-4-nitroimidazole (NI) (32), and these oxidation products are potently
mutagenic in vivo as determined by M13 viral-based mutagenesis studies (33-36).
Previous studies in our laboratory showed that several of the ONOO−-derived
DNA lesions are substrates for the SOS response system in E. coli (35, 36). These
findings strongly suggested involvement of the E. coli polymerases II, IV, and/or V and
prompted us to investigate here the roles of these polymerases in the bypass of these
lesions. In the present work, we have examined the effect of the SOS inducible DNA
polymerases on the bypass and mutagenicity of Gh, Sp1, Sp2, Oa, Ua, and NI. The
results of these experiments will serve to define further the properties of these
polymerases while examining in greater detail the mechanisms at the molecular level for
how ONOO− causes genotoxicity and mutagenesis.
219
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
6.3. Experimental Procedures
Oligodeoxynucleotides. DNA synthesis reagents were purchased from Glen Research.
Unmodified ODNs were purchased from IDT, Inc. and were purified by polyacrylamide
gel electrophoresis (PAGE). The 16mer ODN sequence used was 5’-GAA GAC CTX
GGC GTC C-3’, where X is G, 8-oxoG, Gh, Sp1, Sp2, Oa, Ua, NI, or a tetrahydrofuran
(THF) abasic site. The 16mers containing 8-oxoG or the THF abasic site analog were
prepared on an Applied Biosystems 391 DNA synthesizer and purified by PAGE (HPLC
and MALDI-TOF data for 8-oxoG are given in Figure 6.2). The 16mers containing 8oxoG, Gh, Sp1, Sp2, Oa, Ua, and NI used for genome construction were purified by
anion exchange HPLC on a Dionex NucleoPac PA-100 (4 × 250 mm) analytical column
using 10% CH3CN in water (solvent A) and aqueous 1.5 M NH4OAc (solvent B). A flow
rate of 1.0 mL/min was used and solvent B was increased from 10% to 50% over 5 min,
then 50% to 75% over 25 min. The effluent was monitored at 254 nm. The purified
oligonucleotides were desalted twice with NAP-10 (Amersham Biosciences) columns
(eluting with H2O) and lyophilized. ODNs were characterized by MALDI-TOF mass
spectrometry on a PerSeptive Biosystems (Framingham, MA) Voyager-DE STR
spectrometer. The instrument was operated in the negative ion linear mode with the
accelerating voltage set at 25,000 V, the grid voltage set at 95%, and the guide wire set at
0.28%. A delayed extraction time between 110 and 220 ns and a manual laser intensity
between 1983 and 2083 were used to collect data. The lesion-containing ODN (15-30
pmol) was mixed with 4 pmol of 5’-GAA GAC TGG GCG TC-3’ ([M−H+]: 4327.9) and
1.2 pmol 5’-T18-3’ ([M−H+]: 5412.6). These additional unmodified ODNs serve as
internal standards for instrument calibration and have masses that bracket the mass of the
lesion-containing ODN. A C18-ZipTip (Millipore) was used for sample preparation to
exchange metal cations for triethylammonium cations, and the DNA was eluted with
approximately 2 µL of matrix, which was prepared as described (37).
The 16mer containing Gh was prepared as previously described (38) using 5 nmol
of 8-oxoG-containing 16mer in a reaction volume of 416 µL and purified by anion
220
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
exchange HPLC as described above. [M−H+] (calculated): 4912.2; [M−H+] (observed):
4912.5 (Figure 6.3).
The 16mer containing Sp1 or Sp2 was prepared as previously described (38)
using 5 nmol of 8-oxoG-containing 16mer in a reaction volume of 416 µL. Following
the reaction, the Sp isomers were purified and separated by anion exchange HPLC as
described above. Sp1 [M−H+] (calculated): 4938.2; [M−H+] (observed): 4939.0 Sp2
[M−H+] (calculated): 4938.2; [M−H+] (observed): 4939.0 (Figures 6.4 and 6.5).
The 16mer containing Oa was prepared as previously described (39) by mixing
2.5 nmol of 8-oxoG-containing 16mer with ~130 nmol of ONOO− (a gift of Prof. P.C.
Dedon, MIT) in 50 µL of buffer (125 mM KH2PO4, 25 mM NaHCO3, pH 7.2).
Following the addition of ONOO−, the solution was incubated for 18 h at 37 °C and then
purified by anion exchange HPLC as described above. [M−H+] (calculated): 4887.2;
[M−H+] (observed): 4887.5 (Figure 6.6).
The 16mer containing Ua was prepared by hydrolysis of the unpurified 16mer
containing Oa. Following incubation for 18 h at 37 °C, the 50 µL reaction solution was
adjusted to 750 µL and 0.1 M NaOH, incubated for 6.5 hr at 37 °C, and purified by anion
exchange HPLC as described above. [M−H+] (calculated): 4815.1; [M−H+] (observed):
4815.7 (Figure 6.7).
The 16mer containing NI was prepared as previously described (40) and purified
by anion exchange HPLC as described above. [M−H+] (calculated): 4925.2; [M−H+]
(observed): 4926.0 (Figure 6.8).
Genome Construction. Genomes were constructed on a 50 pmol scale. Single-stranded
M13 DNA (0.5 pmol/µL) was linearized by cleavage with EcoRI (1 U/µL, 23 °C for 8 h)
(New England Biolabs) at a hairpin containing a single EcoRI site (41). The linearized
M13 DNA was annealed with 1.25 equivalents each of scaffolds 5'-GGT CTT CCA CTG
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
AAT CAT GGT CAT AGC-3' and 5'-AAA ACG ACG GCC AGT GAA TTG GAC GC3' (sequences that are partially complementary to the 5’ and 3’ sides of the insert and the
genomic DNA termini) by heating the mixture to 50 °C for 5 min and cooling linearly to
0 °C over 50 min (41). Insert ODNs (75 pmol) were phosphorylated on the 5’-terminus
with 1 U/µL T4 polynucleotide kinase (U.S. Biochemical) in the supplier’s buffer
supplemented with 1 mM ATP and 5 mM DTT (final volume 75 µL, 15 min at room
temperature). The phosphorylated ODN (75 pmol in 75 µL) and the annealed scaffoldM13 mixture (50 pmol M13 in 100 µL) were mixed, and the solution was adjusted to 1
mM ATP, 10 mM DTT, 32 µg/mL BSA, and 22.5 U/µL T4 DNA ligase (New England
Biolabs) (final volume 190 µL) and incubated for 2 h at 16 °C to recircularize the
genome (33, 34, 41). The scaffold DNA was removed using the exonuclease activity of
T4 DNA polymerase (New England Biolabs) (42, 43). The ligation solution was adjusted
to 9.85 mM MgCl2, 50 µg/mL BSA, and 0.25 U/µL T4 DNA polymerase (final volume
210 µL) and incubated for 1 h at 16 °C. The genome constructs were reduced to ~115 µL
in vacuo, extracted with 100 µL of phenol:chloroform:isoamyl alcohol (25:24:1)
(Invitrogen), and desalted with Sephadex G50 fine resin (Amersham Biosciences). The
desalting procedure also removes residual organics.
Normalization of the Genomes. The amount of circular, 16mer insert-containing genome
in each sample was quantified by PAGE and phosphorimagery by the procedure detailed
below.
The desalted, insert-containing genomes were diluted to ~100 µL and the
concentration of genome in each sample was estimated by spectrophotometry using the
conversion factor 14 pmol/mL = 1 A260 unit. Approximately 0.5 pmol of genome (5 µL)
was mixed with 3.5 pmol each of the scaffolds used for genome construction and
incubated with 1.2 U/µL HinF1 (New England Biolabs) and 0.12 U/µL shrimp alkaline
phosphatase (Roche) in 1X Buffer 2 (50 mM NaCl, 10 mM Tris-HCl, 10 mM MgCl2, 1
mM DTT, pH 7.9) (New England Biolabs) for 1 h at 37 °C, followed by 5 min at 80 °C
and cooling to 20 °C at 0.2 °C/s (final volume 8.12 µL). This reaction linearizes the
genome 5’ of the insert and exposes an M13 5’-terminus for radiolabeling.
The
genome/scaffold mixture was supplemented with 5.6 mM DTT, 14 µM ATP, 0.16 µM γ222
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
32
P-ATP (6,000 Ci/mmol, PerkinElmer), 0.47 U/µL T4 polynucleotide kinase, and 0.94
U/µL HaeIII (New England Biolabs) (final volume 10.7 µL) and incubated for 2 h at 37
°C.
This reaction releases a radiolabeled 34mer fragment from the M13 genomes
constructed with a 16mer insert and a 37mer fragment from the M13 genome constructed
with a 19mer insert (referred to as the “+3 control”). The reactions were quenched by the
addition of 10 µL of 2X formamide/EDTA gel loading buffer, and 20 µL of each solution
was analyzed by 20% denaturing PAGE. Dilutions corresponding to 5, 10, 20, 30, 40,
and 50% of the +3 control were also loaded in order to assess how much of the +3 control
to add as an internal standard to the normalized genomes (see Translesion Bypass
Efficiency). The gel was run at 550 V for 2 hr and visualized by phosphorimagery. The
bands corresponding to the 34mer fragments were quantified and the solutions
normalized such that each contained identical amounts of the 34mer fragment. An R2
value of 0.99 was achieved for the dilutions of +3 control (Figure 6.9).
Cell Strains. The cell strains used in this study were gifts of D.F. Jarosz and G.C. Walker
(MIT) (Table 6.1). All strains were characterized by sequencing of PCR products of the
gene regions of interest. E. coli genomes were isolated from 1 mL of an overnight LB
culture grown in the presence of 25 µg/mL chloramphenicol (pol IV− and GW8017
strains), 70 µg/mL spectinomycin (pol II− strain), or both chloramphenicol and
spectinomycin (triple knockout strain). Cells were pelleted by centrifugation at 13,200
RPM for 3 min and resuspended in 570 µL TE buffer, pH 8.0. To this solution was
added 30 µL of 10% SDS and 3 µL of proteinase K (Roche), and the mixture was
incubated for 1 h at 37 °C. Following the incubation, 100 µL of 5 M NaCl and 80 µL of
10% CTAC/0.7 M NaCl were added and the solution was mixed gently. After incubation
at 65 °C for 10 min, the solution was extracted twice with 750 µL
phenol:chloroform:isoamyl alcohol (25:24:1) (Invitrogen). Approximately 700 µL of
aqueous solution remained after extraction. Isopropanol was added (420 µL), and the
solution was mixed thoroughly and then centrifuged for 5 min at 13,200 RPM. The
supernatant was removed and 70% ethanol (250 µL) was added. The solution was
centrifuged for 5 min at 13,200 RPM and the supernatant removed. The DNA pellet was
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
dried in vacuo for 10 min and resuspended in 100 µL TE, pH 8.0. PCR amplification of
the E. coli genomic DNA was carried out using Taq DNA polymerase (Invitrogen). The
50 µL PCR mix for wild-type polB, dinB, and umuDC gene regions and ∆dinB and
∆umuDC gene regions included 25 mM KCl, 10 mM Tris-HCl (pH 8.3), 3.5 mM MgCl2,
2% DMSO, 0.5 µM each primer, 0.05 mM each dNTP, 0.05 unit/µL Taq, and 2 µg of
genomic DNA. The amplification cycle consisted of denaturation at 94 °C for 0.5 min,
annealing at 56 °C for 0.5 min, and extension at 72 °C for 2 min. After 30 cycles,
samples were incubated at 72 °C for 10 min and stored at 4 °C until further use. The 50
µL PCR mix for the polB∆1 gene region included 25 mM KCl, 10 mM Tris-HCl (pH
8.3), 1.5 mM MgCl2, 2% DMSO, 0.5 µM each primer, 0.05 mM each dNTP, 0.05
unit/µL Taq, and 0.5 µg of genomic DNA.
The amplification cycle consisted of
denaturation at 94 °C for 0.5 min, annealing at 56 °C for 0.5 min, and extension at 72 °C
for 3.5 min. After 30 cycles, samples were incubated at 72 °C for 10 min and stored at 4
°C until further use. The primers used amplification of the polB and polB∆1 regions
were 5’-CTG GAA CGA AGT GTA TTA CGG GTT TCG-3’ and 5’-TAT GGT ACT
GGA TGG CAA AGC ATT CG-3’ The primers used amplification of the dinB region
were 5’-CAT GGG GAT AAA GTG GTG CAG C-3’ and 5’-CTG GCA CTT AAG AGA
TAT CCT GCG G-3’. The primers used amplification of the umuDC region were 5’CCA CGT GAG CC AAG ATA AGA GAA CG-3’ and 5’-TAC CTG ATT GTC GCA
GTG CTG G-3’. The PCR products were analyzed by 1% agarose gel electrophoresis in
1X TBE buffer. To 5 µL of each PCR product solution was added 5 µL of H2O and 2 µL
of 6X Ficoll loading buffer (25% Ficoll), and 10 µL were loaded into each well. The gel
was run for 130 V for 1:40 h and stained with 1X SYBR Gold (Molecular Probes) in 1X
TBE for 1 h. The PCR products were prepared for DNA sequencing using a Qiagen PCR
Purification Kit, eluting with a 1:10 dilution of buffer EB (final concentration 1 mM TrisHCl, pH 8.5). For strains containing wild-type polB, 5’-TAT CGA TGC AGC GCA
AAT GCC-3’, 5’-CGA TCA TCC GCA CCT TTC TGA TTG-3’, 5’-CAA TCA GAA
AGG TGC GGA TGA TCG-3’, and 5’-TAT GGT ACT GGA TGG CAA AGC ATT
CG-3’ were used as primers for DNA sequencing. For strains containing polB∆1, 5’CTG GAA CGA AGT GTA TTA CGG GTT TCG-3’ was used in place of 5’-TAT CGA
TGC AGC GCA AAT GCC-3’. For strains containing wild-type dinB and ∆dinB, the
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
same primers used for PCR were used for DNA sequencing. For strains containing wildtype umuDC and ∆umuDC, 5’-GAA AGT TGG AAC CTC TTA CGT GCC-3’ and the
same primers used for PCR were used for DNA sequencing.
Preparation of Electrocompetent Cells. LB media (4 × 150 mL) was inoculated with 1.5
mL each from separate overnight cultures of E. coli. The culture was grown on a shaker
at 37 °C to an A600 of approximately 0.4. The overnight cultures were grown in the
presence of antibiotics (25 µg/mL chloramphenicol and/or 70 µg/mL spectinomycin, see
Cell Strains). Each culture was centrifuged, resuspended in 25 mL of 10 mM MgSO4,
and transferred to a large (150 × 15 mm) Petri dish. The SOS system was induced in two
of the cultures by irradiating with 254 nm light (45 J/m2 of energy), immediately
transferring the cells to 2 × 125 mL of 2×YT media, and growing for 40 min at 37 °C
with shaking. Uninduced cells were treated identically, except without exposure to UV
light. Each set of 2×YT cultures was centrifuged, combined, and washed with 2 × 175
mL of deionized water. Each set of electrocompetent cells was resuspended in 4 mL of a
10% solution of glycerol in water, stored at 4 °C and used the following day.
Production and Isolation of Phage Populations. The normalized genomes were split into
three portions (for triplicate analysis) and +3 control was mixed with each genome
solution to serve as an internal standard for the bypass assay.
An aliquot of
genome/internal standard solution was added to 100 µL of electrocompetent cells. The
cell/genome mixtures were electroporated (2.5 kV, 129 ohms in a 0.2-cm cuvette) and
transferred to 10 mL of LB, generating at least 103 independent, transformed cells
(“infective centers”) as determined by plating of an aliquot onto agar plates immediately
following transformation. The remaining balance of the cultures was incubated at room
temperature for at least 30 min, and then incubated on a roller drum for 4 h at 37 °C to
amplify the progeny phage. The cells were spun down and the progeny phage-containing
supernatant retained. In this system, successful replication of the genomes by the E. coli
host leads to the production of progeny phage. Amplified progeny phage (100 µL) and
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
10 µL of an overnight growth of SCS110 E. coli were added to 10 mL of LB media and
grown for 6 h at 37 °C on a roller drum. The cultures were centrifuged and the phagecontaining supernatant retained. Single-stranded phage DNA was isolated from 700 µL
of each sample using a QIAPrep Spin M13 kit (Qiagen) and used for the determination of
translesion bypass efficiency and mutation frequency.
PCR Amplification. PCR amplification of the isolated M13 DNA was carried out using
Turbo Pfu polymerase (Stratagene). The 25 µL PCR mix included 10 mM KCl, 10 mM
(NH4)2SO4, 20 mM Tris-HCl (pH 8.8), 2 mM MgSO4, 0.1% Triton X-100, 100 µg/mL
BSA, 1.0 µM each primer, 0.2 mM each dNTP, 0.05 unit/µL Pfu, and 10 µL of Qiagen
isolated M13 DNA. The amplification cycle consisted of denaturation at 94°C for 0.5
min, annealing at 67°C for 1 min, and extension at 72°C for 1 min. After 30 cycles,
samples were incubated at 72°C for 5 min and stored at 4°C until further use. The
primers used to generate the 84 bp (from 16mer-insert-containing genomes) or 87 bp
(from the internal standard 19mer-insert-containing genome) PCR products for
determination of translesion bypass efficiency were 5’-Y CAG CTA TGA CCA TGA
TTC AGT GGA AGA C-3’ and 5’-Y CAG GGT TTT CCC AGT CAC GAC GTT GTA
A-3’ [Y is an aminoethoxyethyl ether group from a “5’-amino-modifier 5”
phosphoramidite (Glen Research, Sterling, VA) or an aminohexyl group from a “5’amino-modifier C6” phosphoramidite (IDT, Inc.), which prevent
32
P-labeling of the 5’-
termini]. The primers used to generate the 61 bp PCR products for determination of
mutation frequency were 5’-Y CAG CTA TGA CCA TGA TTC AGT GGA AGA C-3’
and 5’-Y TGT AAA ACG ACG GCC AGT GAA TTG GAC G-3’. Following PCR, each
reaction
was
diluted
to
115
µl
with
H2O,
extracted
with
100
µl
of
phenol:chloroform:isoamyl alcohol (25:24:1), and desalted with Sephadex G50 fine resin.
Restriction Endonuclease Digestion.
An aliquot of the PCR product (10 µL) was
incubated in 20 µL of final volume containing 1X Buffer 2, 5 U BbsI (New England
Biolabs), and 1 U of shrimp alkaline phosphatase (Roche) for 4 h at 37 °C, followed by 5
min at 80 °C and cooling to 20°C at 0.2°C/s. The cut, dephosphorylated PCR product (4
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
µL) was incubated in a 6 µL final volume containing 1X Buffer 2, 10 mM DTT, 3.33 µM
cold ATP, 0.278 µM γ-32P-ATP (6,000 Ci/mmol, PerkinElmer), and 5 U of T4
polynucleotide kinase for 15 min at 37°C, followed by 20 min at 65°C and cooling to
23°C at 0.1°C/s. To the radiolabeled PCR product was added 4 µL of 1X Buffer 2
containing 10 units of HaeIII. The resultant solution was incubated for 2 h at 37 °C, and
the reaction was quenched by the addition of 10 µL of 2X formamide/EDTA loading
buffer.
Translesion Bypass Efficiency.
The samples (20 µL) were analyzed by 20% denaturing
PAGE (550 V for 3-3.5 h) and visualized by phosphorimagery. Two bands of interest
appear on the gel. The faster migrating band results from the genome that contains a
16mer insert, and the slower migrating band results from the internal standard genome
that contains a 19mer insert.
The translesion bypass efficiency was calculated as
described (36, 44).
Mutation Type and Frequency. The REAP assay was used to determine the identity of
the base at the site formerly occupied by the lesion (33, 34, 41, 45).
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
6.4 Results
6.4.1. Translesion bypass efficiency
Given the number of samples to be analyzed in this study, an improved version of
the bypass assay described in Chapter 3 was used (44). Genomes were constructed as
described by inserting a 16mer G- or lesion-containing oligonucleotide into M13mp7L2
single-stranded DNA.
In the improved assay, restriction endonucleases and
polynucleotide kinase were employed to excise and radiolabel a 34mer fragment from the
genome construct. Because the 34mer includes the 16mer insert, quantification of the
fragments by PAGE and phosphorimagery allows by deduction the amount of circular,
16mer insert-containing genome to be determined. A genome containing a 19mer insert
was also constructed for use as an internal standard in the bypass assay. When subjected
to the same quantification procedure, a 37mer oligonucleotide results and quantification
of this band allows specification of the proper ratio of 16mer insert-containing genome
and internal standard. To this end, the quantified 16mer insert-containing genomes were
normalized to each other and mixed with the internal standard to produce an approximate
ratio of 75% 16mer insert-containing genome to 25% 19mer insert-containing genome.
Passage of these mixtures through E. coli causes a change in the ratio that reflects the
efficiency of bypass of a DNA lesion since genomes containing poorly bypassed lesions
exhibit poor survivability and genomes containing well-bypassed lesions exhibit high
survivability relative to the internal standard genome. The harvested progeny phage are
PCR amplified, digested and radiolabeled, and the resultant DNA fragments analyzed by
PAGE and phosphorimagery. An 18mer oligonucleotide results from the 16mer insertcontaining genome, whereas a 21mer oligonucleotide results from the internal standard.
The values of the quantified bands were translated into percent bypass efficiency as
described (44). The bypass efficiency of a lesion is scaled relative to guanine, which is
defined as having a bypass efficiency of 100%. A limitation of this assay is that +3
frameshifts overlap with the signal from the internal standard.
All increases and
decreases noted below are statistically significant.
The bypass efficiencies of 8-oxoG, Oa, Gh, Sp1, Sp2, Ua, and NI were
determined in wild-type, pol II deficient, pol IV deficient, pol V deficient, and pol II / pol
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
IV / pol V deficient E. coli strains under normal and SOS-induced conditions for a total
of ten cellular experimental systems. The bypass efficiencies of 8-oxoG and Oa were
identical within experimental error to G (100%) and did not differ among the ten cellular
systems. The remaining bypass results therefore will focus exclusively on the remaining
lesions. In the wild-type strain, these lesions exhibited poor bypass efficiencies and were
strong substrates for the SOS system (Figure 6.10). The Sp diastereomers showed a
significant difference in bypass efficiency under both normal and SOS-induced
conditions, with the Sp1 diastereomer bypassed more efficiently.
As a control
demonstrating a blocking lesion and showing induction of the SOS system, the bypass
efficiency of a tetrahydrofuran (THF) synthetic abasic site was determined and found to
increase from 1.0 ± 0.4% to 19 ± 9%, or about 20 fold. In contrast, the bypass efficiency
increased about 3 fold from 19 ± 4% to 65 ± 8% for Gh, increased about 4 fold from 16 ±
2% to 69 ± 5% for Sp1, increased about 6 fold from 8.2 ± 2.3 to 50 ± 8 for Sp2, increased
about 6 fold from 5.2 ± 1.7% to 34 ± 10% for Ua, and increased about 8.5 fold from 4.5 ±
1.8% to 38 ± 7% for NI.
Deletion of pol II had no significant effect on the bypass efficiencies of Gh, Sp2,
Ua, NI, and THF, but caused a reduction in the bypass efficiency of Sp1 from 16 ± 2% to
8.2 ± 2.3% (Figure 6.11). No significant differences were observed upon SOS induction
as compared to SOS induced wild-type cells. In comparison to results in wild-type cells,
deletion of pol IV produced a significant effect only for Sp1, decreasing the bypass
efficiency from 16 ± 2% to 10 ± 1% and 69 ± 5% to 60 ± 1% in uninduced and SOSinduced cells, respectively (Figure 6.12).
Deletion of pol V produced dramatic changes in the bypass efficiencies (Figure
6.13). With the exception of 8-oxoG and Oa, which exhibited no differences upon
inactivation of pol V, the bypass efficiencies were lower in uninduced pol V deficient
cells as compared to uninduced wild-type cells and essentially did not increase for any of
the lesions upon induction of the SOS system. In the pol V deficient strain, Gh was
bypassed with efficiencies of 8.7 ± 3.0% and 9.7 ± 4.0% in uninduced and SOS-induced
pol V deficient cells, respectively. The bypass efficiency of Sp1 decreased to 0.44 ±
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
0.16% and 0.39 ± 0.16% in uninduced and SOS-induced cells, respectively, whereas
essentially no bypass was observed for Sp2. In uninduced cells, the bypass efficiency of
Ua decreased to 2.3 ± 0.6% and in SOS induced cells decreased to 1.9 ± 0.2%. The
bypass efficiency of NI decreased to 1.7 ± 0.5% and 2.7 ± 0.4% in normal and SOSinduced cells, respectively. Results from the pol II / pol IV / pol V deficient strain
mirrored those from the pol V deficient strain (Figure 6.14).
6.4.2. Mutation type and frequency
The mutational signature of each lesion in WT AB1157 E. coli was determined
using the restriction endonuclease and post-labeling analysis of mutation frequency
(REAP) assay (46). As for the bypass assay results, all increases and decreases noted
below are statistically significant.
Frameshift mutations. In wild type cells, both uninduced and SOS-induced, frameshift
mutations were negligible (less than 1%). Frameshifts (−1) were observed in the pol V
deficient strain and the triple knockout strain for Sp1 and Sp2 at levels of about 2% and
7.5%, respectively, in uninduced cells. Sp1 also generated roughly 1.5% −2 frameshifts,
whereas Sp2 produced about 6%. When these strains were SOS induced, Sp1 generated
about 15% and 4% −1 frameshifts in the pol V deficient and triple knockout strains,
respectively. Sp2 produced about 10% and 12% −1 frameshifts in the SOS induced pol V
deficient and triple knockout strains, respectively. In uninduced cells, NI produced about
2% −1 frameshifts in the pol II and pol IV deficient strains, about 7% in the pol V
deficient strain, and about 10% in the triple knockout strain. A low amount (~1%) of −2
frameshifts were also observed in the pol V deficient strain. When the SOS system was
induced, NI caused about 6% −1 frameshifts in the pol V deficient strain and about 16%
in the triple knockout strain.
Point mutations in wild-type cells (Figure 6.15). In uninduced cells, replication of 8oxoG was 1.72% mutagenic, and the mutations were primarily GÆT. Gh, Sp1, Sp2, Oa,
and Ua were all about 99% mutagenic, but the mutational signatures differed greatly
among this group of lesions. Gh induced 37% GÆT and 59% GÆC mutations, whereas
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Sp1 caused 83% GÆT and 15% GÆC mutations and Sp2 caused 43% GÆT and 54%
GÆC mutations. Oa generated 99% GÆT mutations, and Ua produced 9% GÆA, 55%
GÆT, and 35% GÆC mutations. NI was 64% mutagenic and induced 16% GÆA, 30%
GÆT, and 18% GÆC mutations. In SOS-induced cells, a small increase in GÆA
mutations was observed for Gh (1.9% to 3.6%). No difference in mutational signature
was observed for Sp1, but GÆT mutations induced by Sp2 increased to 58% and GÆC
mutations decreased to 39%. The mutational signature of Oa changed slightly and the
lesion induced 1.0% GÆA, 97% GÆT, and 1.7% GÆC mutations.
The mutation
frequency and signature of Ua was essentially unchanged. The mutation frequency of NI
decreased to 48% and the mutational signature shifted to fewer GÆA mutations (9.0%)
and GÆC mutations (4.7%).
Point mutagenesis in polymerase deficient cells.
Figure 6.16 shows the mutation
frequency of 8-oxoG as a function of cell type under both uninduced and SOS-induced
conditions.
The mutation frequency remains constant within experimental error in
uninduced cells. Comparison of the mutation frequencies in SOS induced cells reveals a
small decrease in the amount GÆT mutations in pol V deficient and triple knockout
strains as compared to wild-type.
The results for Gh are shown in Figure 6.17. In uninduced cells, statistically
significant differences were only observed for GÆA mutations, which decreased from 12% in wild-type, pol II deficient, and pol IV deficient cells to about 0.1% in pol V
deficient and triple knockout cells. Induction of the SOS system caused a decrease in
GÆC mutations from 62% to 52% in pol II deficient cells. GÆA mutations increased to
3-5% in pol II and pol IV deficient cells upon SOS induction. In the pol V deficient
strain, SOS induction decreased the GÆA mutations to 0.13%. Relative to SOS induced
wild-type cells, GÆT mutations decreased to 35%, whereas GÆC mutations increased to
64% in SOS induced pol V deficient cells.
The general mutational signature for Sp1 did not differ among the pol V
proficient strains; however, small differences were noted for the specific mutations
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
observed (Figure 6.18). In the pol II deficient strain, SOS induction caused an increase
from 79% to 87% in GÆT mutations and a decrease from 19% to 11% in GÆC
mutations. Induction of the SOS system in the pol IV deficient strain increased GÆT
mutations from 80% to 85% and decreased GÆC mutations from 18% to 13%. The
overall mutation frequency decreased from 99% in uninduced and SOS induced pol V
proficient cells to 93% in uninduced and 94% in SOS induced pol V deficient cells.
GÆT mutations increased from 79-83% in uninduced and 85-87% in SOS induced pol V
proficient cells to 88% in uninduced and 91% in SOS induced pol V deficient cells.
Fewer GÆC mutations resulted and were 5% in uninduced and 3% in SOS induced pol V
deficient cells versus 15-19% in uninduced and 11-13% in pol V proficient cells.
Comparison of the results for wild-type and pol II deficient strains indicate pol II
is not involved in Sp2 induced mutagenesis (Figure 6.19). Elimination of pol IV does not
affect the mutational signature of Sp2 uninduced cells. In SOS-induced cells, elimination
of this polymerase reduces the amount of GÆT mutations from 57% to 49% and the
amount of GÆC mutations from 38% to 48% as compared to SOS induced pol II
deficient cells. These differences may also apply in comparison to SOS induced wildtype cells, but statistical error precludes this determination. Dramatic differences in
mutational signature were observed upon elimination of pol V. In both uninduced and
SOS induced cells, the overall mutation frequency decreased from 99% to 86%. GÆT
mutations increased from 43-46% to 79% and from 49-58% to 74% in uninduced and
SOS induced cells, respectively. GÆC mutations decreased from 51-54% to 6% in
uninduced cells and 38-48% to 10% in SOS induced cells.
The mutation frequency of Oa remained constant at greater than 99% across all
strains and under both uninduced and SOS induced conditions (Figure 6.20). By far, the
predominant mutation induced was GÆT. In SOS induced pol II and pol IV deficient
strains, GÆC mutations were higher than in uninduced wild-type cells, and removal of
pol V lowered the GÆC mutations from 1.6-2.6% to less than 0.5% in SOS induced
cells.
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
The mutational signature of Ua was similar in wild-type, pol II deficient, and pol
IV deficient strains and was drastically different in the pol V deficient strains under both
uninduced and SOS induced conditions (Figure 6.21). GÆT mutations were the major
mutations in the pol V deficient strains. These mutations increased from 55-69% to 93%
in uninduced cells and from 47-53% to 94% in SOS induced cells. GÆC transversions
accounted for the remaining mutations.
Arguably the most interesting results came from NI (Figure 6.22). In uninduced
cells, the overall mutation frequency of the lesion increased from 64% to 81% upon
removal of pol II, but decreased to 57% in the pol IV deficient strain, 40% in the pol V
deficient strain, and 51% in the triple knockout strain. In SOS induced cells, the mutation
frequency increased to 82% from 48% when pol II was removed, increased slightly to
49% when pol IV was removed, decreased to 21% when pol V was removed and
decreased slightly to 45% when all three SOS inducible polymerases were removed.
These results suggest the involvement of both pol II and pol V in the bypass of NI, and
the differences in mutational signature among the cell strains tested support this
conclusion. Removal of pol II did not affect the amount of GÆA and GÆC mutations
induced but caused an increase in the amount of GÆT mutations to 45% and 62% in
uninduced and SOS induced cells, respectively, with a concomitant increase in the overall
point mutation frequency. This result suggests pol II preferentially inserts a C opposite
the lesion. Removal of pol IV did not affect the mutational signature of the lesion,
whereas removal of pol V caused a large decrease in the amount of GÆT mutations with
a corresponding decrease in mutation frequency. In uninduced cells, the amount of GÆT
mutations decreased from 30% to 7% and in SOS induced cells decreased from 35% to
4% relative to wild-type cells. This result shows involvement of pol V in generating
GÆT mutations from NI. Results from the triple knockout strain resemble those from
the pol V single knockout strain.
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
6.5. Discussion
All of the lesions studied in this work were previously examined in wild type E.
coli cells in the sequence context GXA, where X is the site of the lesion (34-36). Results
from the present TXG sequence context differ dramatically in some cases. For example,
in the GXA context Gh is bypassed with 75% efficiency and induces 98% GÆC
mutations. In the context TXG, bypass of Gh occurred with only 19% efficiency, and the
mutational signature shifted to 59% GÆC and 37% GÆT. In the case of Sp1, the bypass
efficiency in the TXG context was 16% versus 9% for the GXA context, and the
mutational signature changed from 72% GÆC and 27% GÆT to 15% GÆC and 83%
GÆT. The bypass efficiency and mutational signature of Sp2 did not differ between the
two sequence contexts. The bypass properties of the hydantoin lesions, Gh and Sp, have
shown sequence context dependence previously. Burrows and coworkers found that the
bypass of Gh and Sp by Klenow fragment occurs more readily in the sequence context
XGG as compared to XAA and proposed the differences were due either to primer
misalignment or to stabilization of the primer/template complex by the GG pair 3’ of the
lesion (38).
In the first case, simple primer misalignment should yield frameshift
mutations in our system; however, these mutations are not observed upon replication in
wild-type cells. A complex primer misalignment in which a bulge in the template forms
initially to allow synthesis beyond the lesion followed by slippage of the template to
remove the bulge could account for the large increase in GÆT mutations observed by us
for Gh and Sp1 in the TXG context without concomitant frameshift mutations. As for the
argument about the sequence 3’ of the template, the present sequence is TXGG, whereas
the sequence we used previously was GXAG. According to the hypothesis put forth by
Burrows and coworkers, the TXGG sequence should produce a more stable duplex
structure prior to translesion synthesis as a result of the more stable GG pair and facilitate
lesion bypass. Whereas the bypass results for Sp1 are as predicted by this hypothesis, the
results for Gh disagree.
Significant differences in mutational signature were also
observed for Ua. In the GXA context, Ua induces 99% GÆT mutations in uninduced
cells; however, in the present study, Ua induced 9% GÆA, 55% GÆT and 35% GÆC
mutations. The mutational signature of NI also changed although the overall mutation
pattern remained consistent. Clearly, the sequence context effect is complicated, and
234
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
further experiments are needed to determine the reason for the large changes observed for
the bypass and coding properties of these DNA lesions.
Comparison of the data in Figures 6.10 and 6.13 shows that pol V was responsible
for the vast majority of SOS dependent translesion synthesis for the oxidation products in
this study. Pol II and pol IV appeared to have a small role in the bypass efficiency of
Sp1. By contrast, pol V was essential for bypass. The small effect of pol IV existed in
both uninduced and SOS-induced cells, whereas the reduction in bypass efficiency as a
result of pol II deficiency was rescued in SOS-induced cells. In its simplest form, the
bypass of a DNA lesion consists of two parts, the first part is insertion of a nucleotide
opposite the lesion and the second part is extension of the nascent strand (47). One
possible explanation for the Sp1 bypass observations is that pol V is required for the
extension step and pol II and/or pol IV participate in, but are not required for, the
insertion step. A second explanation is that pol II and/or pol IV may participate in the
extension step with pol V being required for the insertion step. As a third possibility, pol
II and pol IV may increase the rate of translesion synthesis without the use of their
inherent polymerase activity, for instance by interacting with the replication complex or
with pol V.
The bypass results from wild type, pol V deficient, and triple knockout cells
demonstrated that pol I and/or pol III can bypass Gh, Ua, and NI to a small extent and
Sp1 and Sp2 to a negligible (but still measurable in terms of determining mutation
frequency) extent. Experiments with Klenow fragment suggest that the extension step is
rate limiting for Gh (38, 48, 49), Ua (50), and NI (51). Thus, it is possible that the role of
pol V in the bypass of these lesions is more for extension rather than insertion. However,
pol V undoubtedly has a significant role in the insertion step for Ua and NI based on the
large differences in mutational signature that occurred with these lesions upon
inactivation of pol V.
NI was the only lesion for which involvement of more than one SOS inducible
polymerase could be concluded from the mutagenesis data. Inactivation of pol II caused
235
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
a significant increase in the amount of A inserted opposite the lesion and a significant
decrease in the amount of C incorporated opposite the lesion, while causing no decrease
in the bypass efficiency. This result suggests pol II is involved principally during the
insertion step of translesion synthesis and preferably inserts C opposite the lesion, which
leads to no mutation. Inactivation of pol V causes a large increase in the amount of C
inserted opposite the lesion and a significant decrease in the amount of A inserted
opposite the lesion. This result demonstrates that pol V preferentially inserts A opposite
the lesion leading to GÆT mutations. Since the wild type mutational signature consists
of roughly equivalent amounts of no mutation and GÆT mutations, pol II and pol V
appear to have comparable rates of insertion opposite NI. This result coupled with the
strong dependence of NI bypass efficiency on pol V suggests pol V is responsible for
extension of the primer after incorporation of a nucleotide opposite the lesion by pol II or
pol V.
The involvment of two polymerases for the bypass of a lesion has been
demonstrated previously in the case of benzo[a]pyrene and N-2-acetylaminofluorene (22,
52, 53).
The Sp diastereomers exhibited markedly different mutational signatures and
responded differently to the knockout of pol V. Sp1 induced more than 85% GÆT and
about 10% GÆC mutations in SOS induced pol V proficient strains. The elimination of
pol V activity caused a small decrease in the amount of GÆC mutations and a
corresponding increase in the amount of GÆT mutations. This result contrasts with that
from Sp2 where the inactivation of pol V caused the mutational signature to shift from
similar amounts of GÆT and GÆC mutations to primarily GÆT mutations. This result
indicates that in the case of Sp2, pol V induces principally GÆC mutations. As with the
other lesions studied, it is not possible to conclude from these data in the case of Sp1 if
pol V is more important for extension or insertion. For Sp2, pol V has a significant role
in insertion; however, the significance of this polymerase for extension after the insertion
step is unclear. The stereochemical difference between Sp1 and Sp2 is the logical
explanation for the observed differences in bypass and mutational signature. Jia et al.
performed a computational study of duplexes containing the Sp diastereomers (54). The
authors note that the orientation in the duplex of one isomer is opposite that of the other
236
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
isomer and that the duplex containing the R isomer is always lower in energy than the
duplex containing the S isomer. Furthermore, the A ring of both isomers mimics the
hydrogen bonding face of thymine, whereas the R isomer forms a more stable base pair
with G than the S isomer.
While these results do not allow prediction of bypass
efficiencies or specific coding properties, they do set a precedent for differences based on
stereochemistry.
In this study, we have examined the role of the E. coli SOS inducible polymerases
in the bypass and mutagenicity of selected G oxidation products. Our results demonstrate
a clear requirement of the umuDC gene product for the bypass of Sp, Ua, and NI, and
strong involvement of this gene product in the bypass of Gh. These results expand the
substrate repertoire of pol V to include ring opened G oxidation products and provide
mechanistic details for how these oxidation products may cause mutations and ultimately
cancer in inflamed tissues.
237
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
6.6. Acknowledgment
Sarah Delaney (MIT), Yuriy O. Alekseyev (MIT), James C. Delaney (MIT), and John M.
Essigmann (MIT) contributed to this work. We thank Daniel Jarosz and Graham Walker
(MIT) for providing the cell strains used in this work. We also thank Peter T. Rye for
assistance with cell strain characterization.
We acknowledge the MIT Biological
Engineering Division Mass Spectrometry Laboratory and the MIT Biopolymers
Laboratory for their services. Financial support was provided by NIH awards CA26731
and CA080024 and by NSF awards DBI-9729592 and CHE-9808061.
238
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.1. Products of ONOO− oxidation and nitration of G.
O
O
N
HN
H2N
N
G
HN
N
R
H2N
H
N
O
N
N
R
H2N
HO
O
O
N
H
Oa
N
H
N
R
O
O
NH
R
H2N
HN
H2N
Ua
239
NH
R
H2N
O
N
NH
N
O R
Sp
Gh
8-oxoG
O
O
NH
H
N
O2N
NH
N
N
H
N
R
NI
O
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.2. (Top) HPLC trace of purified 8-oxoG oligonucleotide. (Bottom) MALDITOF spectrum of purified 8-oxoG oligonucleotide.
7.5
Absorbance (mAU)
5.0
2.5
0.0
-2.5
15.0
17.5
20.0
22.5
25.0
27.5
30.0
Retention Time (Minutes)
100
4922.0
8-oxoG
% Intensity
80
60
40
20
0
4000
14mer MW standard
4327.9
4400
18mer MW standard
5412.6
4800
5200
Mass (m/z)
240
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.3. (Top) HPLC trace of purified Gh oligonucleotide. (Bottom) MALDI-TOF
spectrum of purified Gh oligonucleotide.
15
Absorbance (mAU)
10
5
0
15.0
17.5
20.0
22.5
25.0
27.5
30.0
Retention Time (Minutes)
100
4912.5
Gh
% Intensity
80
60
40
20
0
4000
14mer MW standard
4327.9
4400
18mer MW standard
5412.6
4800
Mass (m/z)
241
5200
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.4. (Top) HPLC trace of purified Sp1 oligonucleotide. (Bottom) MALDI-TOF
spectrum of purified Sp1 oligonucleotide.
Absorbance (mAU)
10
5
0
-5
15.0
17.5
20.0
22.5
25.0
27.5
30.0
Retention Time (Minutes)
100
4939.0
Sp1
% Intensity
80
60
40
20
0
4000
14mer MW standard
4327.9
4400
18mer MW standard
5412.6
4800
5200
Mass (m/z)
242
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.5. (Top) HPLC trace of purified Sp2 oligonucleotide. (Bottom) MALDI-TOF
spectrum of purified Sp2 oligonucleotide.
7.5
Absorbance (mAU)
5.0
2.5
0.0
-2.5
15.0
17.5
20.0
22.5
25.0
27.5
30.0
Retention Time (Minutes)
100
4939.0
Sp2
% Intensity
80
60
40
20
0
4000
14mer MW standard
4327.9
4400
18mer MW standard
5412.6
4800
5200
Mass (m/z)
243
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.6. (Top) HPLC trace of purified Oa oligonucleotide. (Bottom) MALDI-TOF
spectrum of purified Oa oligonucleotide.
Absorbance (mAU)
30
20
10
0
15.0
100
17.5
20.0
22.5
25.0
Retention Time (Minutes)
27.5
30.0
4887.5
Oa
% Intensity
80
60
40
20
0
4000
+ K+
14mer MW standard
4327.9
4400
18mer MW standard
5412.6
4800
5200
Mass (m/z)
244
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.7. (Top) HPLC trace of purified Ua oligonucleotide. (Bottom) MALDI-TOF
spectrum of purified Ua oligonucleotide.
Absorbance (mAU)
30
20
10
0
15.0
17.5
20.0
22.5
25.0
27.5
30.0
Retention Time (Minutes)
100
4815.7
Ua
% Intensity
80
60
40
20
0
4000
14mer MW standard
4327.9
4400
18mer MW standard
5412.6
4800
5200
Mass (m/z)
245
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.8. (Top) HPLC trace of purified NI oligonucleotide. (Bottom) MALDI-TOF
spectrum of purified NI oligonucleotide.
12.5
10.0
Absorbance (mAU)
7.5
5.0
2.5
0.0
-2.5
15.0
17.5
20.0
22.5
25.0
27.5
30.0
Retention Time (Minutes)
100
4909.9
NI
4894.4
% Intensity
80
60
40
20
0
4000
4926.0
14mer MW standard
4327.9
4400
4800
18mer MW standard
5412.6
5200
Mass (m/z)
246
5600
6000
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.9. Genome quantification gel and standard curve (see Experimental Procedures for details). (Left) PAGE of digested and
radiolabeled competitor and genome constructs (lanes 1-11). G (lane 1), 8-oxoG (lane 2), Gh (lane 3), Sp1 (lane 4), Sp2 (lane 5), Oa
(lane 6), Ua (lane 7), NI (lane 8), THF (lane 9). The unlabeled lanes are not relevant to the present work. (Right) Plot of quantified
competitor lanes, R2 = 0.99.
1 2 3 4 5 6 7 8
9
M13 Competitor (37mer Fragment) Standard Curve
PhosphorImager Volume
6000000
37mer
34mer
5000000
4000000
3000000
2000000
1000000
0
0
10
20
30
40
Theoretical Amount (%)
% Competitor:
5 10 20 30 40 50
247
50
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.10. Bypass efficiency of DNA lesions relative to G in wild-type cells.
140
Bypass Efficiency (%)
120
100
80
60
40
20
G
8-oxoG
Gh
Sp1
Sp2
Lesion
248
Oa
Ua
NI
THF
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.11. Bypass efficiency of DNA lesions relative to G in pol II deficient cells.
140
Bypass Efficiency (%)
120
100
80
60
40
20
0
G
8-oxoG
Gh
Sp1
Sp2
Lesion
249
Oa
Ua
NI
THF
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.12. Bypass efficiency of DNA lesions relative to G in pol IV deficient cells.
140
Bypass Efficiency (%)
120
100
80
60
40
20
0
G
8-oxoG
Gh
Sp1
Sp2
Lesion
250
Oa
Ua
NI
THF
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.13. Bypass efficiency of DNA lesions relative to G in pol V deficient cells.
140
Bypass Efficiency (%)
120
100
80
60
40
20
0
G
8-oxoG
Gh
Sp1
Sp2
Lesion
251
Oa
Ua
NI
THF
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.14. Bypass efficiency of DNA lesions relative to G in triple knockout cells.
140
Bypass Efficiency (%)
120
100
80
60
40
20
0
G
8-oxoG
Gh
Sp1
Sp2
Lesion
252
Oa
Ua
NI
THF
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.15. (A) REAP assay results for uninduced wild-type cells. (B) REAP assay
results for SOS-induced wild-type cells.
A 100
uninduced wild-type
A
G
T
C
Mutation Frequency (%)
80
60
40
20
0
G
GO
Gh
Sp1
Sp2
Oa
Ua
NI
Lesion
B 100
A
G
T
C
SOS-induced wild-type
Mutation Frequency (%)
80
60
40
20
0
G
GO
Gh
Sp1
Sp2
Lesion
253
Oa
Ua
NI
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.16. (A) REAP assay results for 8-oxoG in uninduced cells. (B) REAP assay
results for 8-oxoG in SOS-induced cells.
8-oxoG uninduced
100
A
G
T
C
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
8-oxoG SOS induced
100
A
G
T
C
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
254
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.17. (A) REAP assay results for Gh in uninduced cells. (B) REAP assay results
for Gh in SOS-induced cells.
100
A
G
T
C
Gh uninduced
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
100
Gh SOS-induced
Mutation Frequency (%)
80
A
G
T
C
60
40
20
0
Wild type
Pol II minus
Pol IV minus
Cell Strain
255
Pol V minus
Triple mutant
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.18. (A) REAP assay results for Sp1 in uninduced cells. (B) REAP assay results
for Sp1 in SOS-induced cells.
100
Sp1 uninduced
A
G
T
C
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
100
A
G
T
C
Sp1 SOS-induced
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus
Pol IV minus
Cell Strain
256
Pol V minus
Triple mutant
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.19. (A) REAP assay results for Sp2 in uninduced cells. (B) REAP assay results
for Sp2 in SOS-induced cells.
100
Sp2 uninduced
Mutation Frequency (%)
80
A
G
T
C
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
100
Sp2 SOS-induced
Mutation Frequency (%)
80
A
G
T
C
60
40
20
0
Wild type
Pol II minus
Pol IV minus
Cell Strain
257
Pol V minus
Triple mutant
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.20. (A) REAP assay results for Oa in uninduced cells. (B) REAP assay results
for Oa in SOS-induced cells.
Oa uninduced
100
A
G
T
C
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
Oa SOS-induced
100
A
G
T
C
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus
Pol IV minus
Cell Strain
258
Pol V minus
Triple mutant
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.21. (A) REAP assay results for Ua in uninduced cells. (B) REAP assay results
for Ua in SOS-induced cells.
100
Ua uninduced
Mutation Frequency (%)
80
A
G
T
C
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
100
Ua SOS-induced
Mutation Frequency (%)
80
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
259
A
G
T
C
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Figure 6.22. (A) REAP assay results for NI in uninduced cells. (B) REAP assay results
for NI in SOS-induced cells.
100
NI uninduced
Mutation Frequency (%)
80
A
G
T
C
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
100
NI SOS-induced
Mutation Frequency (%)
80
A
G
T
C
60
40
20
0
Wild type
Pol II minus Pol IV minus Pol V minus Triple mutant
Cell Strain
260
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
Table 6.1. E. coli strains used in this work.
Strain
AB1157
Genotype
F thr-1 araC14 leuB6(Am) ∆(gpt-proA)62 lacY1 tsx-33
supE44(AS) galK2(Oc) hisG4(Oc) rfbD1 mgl-51
rpoS396(Am) rpsL31(Strr) kdgK51 xylA5 mtl-1 argE3(Oc)
thi-1
−
pol II
As AB1157 but polB∆1::Ω Sm-Sp
pol IV− As AB1157 but dinB::cat
GW8017 As AB1157 but umuDC595::cat
triple k.o. As AB1157 but polB∆1::Ω Sm-Sp ∆dinB umuDC595::cat
–
261
Source
G. Walker
G. Walker
G. Walker
G. Walker
G. Walker
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
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Escherichia coli. DNA Repair (Amst) 2, 1227-1238.
(24) Livneh, Z. (2001) DNA damage control by novel DNA polymerases: translesion
replication and mutagenesis. J. Biol. Chem. 276, 25639-25642.
(25) Maor-Shoshani, A., Ben Ari, V., and Livneh, Z. (2003) Lesion bypass DNA
polymerases replicate across non-DNA segments. Proc. Natl. Acad. Sci.
U. S. A. 100, 14760-14765.
(26) Xia, Y., and Zweier, J. L. (1997) Superoxide and peroxynitrite generation from
inducible nitric oxide synthase in macrophages. Proc. Natl. Acad. Sci. U.
S. A. 94, 6954-6958.
(27) Carreras, M. C., Pargament, G. A., Catz, S. D., Poderoso, J. J., and Boveris, A.
(1994) Kinetics of nitric oxide and hydrogen peroxide production and
formation of peroxynitrite during the respiratory burst of human
neutrophils. FEBS Lett. 341, 65-68.
(28) Burney, S., Caulfield, J. L., Niles, J. C., Wishnok, J. S., and Tannenbaum, S. R.
(1999) The chemistry of DNA damage from nitric oxide and peroxynitrite.
Mutat. Res. 424, 37-49.
(29) Juedes, M. J., and Wogan, G. N. (1996) Peroxynitrite-induced mutation spectra
of pSP189 following replication in bacteria and in human cells. Mutat.
Res. 349, 51-61.
(30) Steenken, S., and Jovanovic, S. V. (1997) How easily oxidizable is DNA? Oneelectron reduction potentials of adenosine and guanosine radicals in
aqueous solution. J. Am. Chem. Soc. 119, 617-618.
(31) Burney, S., Niles, J. C., Dedon, P. C., and Tannenbaum, S. R. (1999) DNA
damage in deoxynucleosides and oligonucleotides treated with
peroxynitrite. Chem. Res. Toxicol. 12, 513-520.
(32) Dedon, P. C., and Tannenbaum, S. R. (2004) Reactive nitrogen species in the
chemical biology of inflammation. Arch. Biochem. Biophys. 423, 12-22.
(33) Henderson, P. T., Delaney, J. C., Gu, F., Tannenbaum, S. R., and Essigmann, J.
M. (2002) Oxidation of 7,8-dihydro-8-oxoguanine affords lesions that are
potent sources of replication errors in vivo. Biochemistry 41, 914-921.
(34) Henderson, P. T., Delaney, J. C., Muller, J. G., Neeley, W. L., Tannenbaum, S.
R., Burrows, C. J., and Essigmann, J. M. (2003) The hydantoin lesions
formed from oxidation of 7,8-dihydro-8-oxoguanine are potent sources of
replication errors in vivo. Biochemistry 42, 9257-9262.
264
Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
(35) Henderson, P. T., Neeley, W. L., Delaney, J. C., Gu, F., Niles, J. C., Hah, S. S.,
Tannenbaum, S. R., and Essigmann, J. M. (2005) Urea lesion formation
in DNA as a consequence of 7,8-dihydro-8-oxoguanine oxidation and
hydrolysis provides a potent source of point mutations. Chem. Res.
Toxicol. 18, 12-18.
(36) Neeley, W. L., Delaney, J. C., Henderson, P. T., and Essigmann, J. M. (2004) In
vivo bypass efficiencies and mutational signatures of the guanine
oxidation products 2-aminoimidazolone and 5-guanidino-4nitroimidazole. J. Biol. Chem. 279, 43568-43573.
(37) Zhang, L. K., and Gross, M. L. (2000) Matrix-assisted laser desorption/ionization
mass spectrometry methods for oligodeoxynucleotides: improvements in
matrix, detection limits, quantification, and sequencing. J. Am. Soc. Mass
Spectrom. 11, 854-865.
(38) Kornyushyna, O., Berges, A. M., Muller, J. G., and Burrows, C. J. (2002) In vitro
nucleotide misinsertion opposite the oxidized guanosine lesions
spiroiminodihydantoin and guanidinohydantoin and DNA synthesis past
the lesions using Escherichia coli DNA polymerase I (Klenow fragment).
Biochemistry 41, 15304-15314.
(39) Tretyakova, N. Y., Niles, J. C., Burney, S., Wishnok, J. S., and Tannenbaum, S.
R. (1999) Peroxynitrite-induced reactions of synthetic oligonucleotides
containing 8-oxoguanine. Chem. Res. Toxicol. 12, 459-466.
(40) Neeley, W. L., Henderson, P. T., and Essigmann, J. M. (2004) Efficient synthesis
of DNA containing the guanine oxidation-nitration product 5-guanidino-4nitroimidazole: Generation by a postsynthetic substitution reaction. Org.
Lett. 6, 245-248.
(41) Delaney, J. C., and Essigmann, J. M. (1999) Context-dependent mutagenesis by
DNA lesions. Chem. Biol. 6, 743-753.
(42) Moriya, M. (1993) Single-stranded shuttle phagemid for mutagenesis studies in
mammalian cells: 8-oxoguanine in DNA induces targeted G.C-->T.A
transversions in simian kidney cells. Proc. Natl. Acad. Sci. U. S. A. 90,
1122-1126.
(43) Moriya, M., Zhang, W., Johnson, F., and Grollman, A. P. (1994) Mutagenic
potency of exocyclic DNA adducts: marked differences between
Escherichia coli and simian kidney cells. Proc. Natl. Acad. Sci. U. S. A.
91, 11899-11903.
(44) Delaney, J. C., Smeester, L., Wong, C., Frick, L. E., Taghizadeh, K., Wishnok, J.
S., Drennan, C. L., Samson, L. D., and Essigmann, J. M. (2005) AlkB
reverses etheno DNA lesions caused by lipid oxidation in vitro and in
vivo. Nat. Struct. Mol. Biol. 12, 855-860.
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Chapter 6 – Effects of Bypass Polymerases on Guanine Oxidation Products
(45) Kroeger, K. M., Jiang, Y. L., Kow, Y. W., Goodman, M. F., and Greenberg, M.
M. (2004) Mutagenic effects of 2-deoxyribonolactone in Escherichia coli.
An abasic lesion that disobeys the A-rule. Biochemistry 43, 6723-6733.
(46) Delaney, J. C., and Essigmann, J. M. (2004) Mutagenesis, genotoxicity, and
repair of 1-methyladenine, 3-alkylcytosines, 1-methylguanine, and 3methylthymine in alkB Escherichia coli. Proc. Natl. Acad. Sci. U. S. A.
101, 14051-14056.
(47) Bridges, B. (2001) The two-step model for translesion synthesis: then and now.
Mutat. Res. 485, 61-67.
(48) Duarte, V., Muller, J. G., and Burrows, C. J. (1999) Insertion of dGMP and
dAMP during in vitro DNA synthesis opposite an oxidized form of 7,8dihydro-8-oxoguanine. Nucleic Acids Res. 27, 496-502.
(49) Sugden, K. D., and Martin, B. D. (2002) Guanine and 7,8-dihydro-8-oxoguanine-specific oxidation in DNA by chromium(V). Environ. Health
Perspect. 110 Suppl 5, 725-728.
(50) McNulty, J. M., Jerkovic, B., Bolton, P. H., and Basu, A. K. (1998) Replication
inhibition and miscoding properties of DNA templates containing a sitespecific cis-thymine glycol or urea residue. Chem. Res. Toxicol. 11, 666673.
(51) Gu, F., Stillwell, W. G., Wishnok, J. S., Shallop, A. J., Jones, R. A., and
Tannenbaum, S. R. (2002) Peroxynitrite-induced reactions of synthetic
oligo 2'-deoxynucleotides and DNA containing guanine: formation and
stability of a 5-guanidino-4-nitroimidazole lesion. Biochemistry 41, 75087518.
(52) Fuchs, R. P., Koffel-Schwartz, N., Pelet, S., Janel-Bintz, R., Napolitano, R.,
Becherel, O. J., Broschard, T. H., Burnouf, D. Y., and Wagner, J. (2001)
DNA polymerases II and V mediate respectively mutagenic (-2
frameshift) and error-free bypass of a single N-2-acetylaminofluorene
adduct. Biochem. Soc. Trans. 29, 191-195.
(53) Wagner, J., Etienne, H., Janel-Bintz, R., and Fuchs, R. P. (2002) Genetics of
mutagenesis in E. coli: various combinations of translesion polymerases
(Pol II, IV and V) deal with lesion/sequence context diversity. DNA
Repair (Amst) 1, 159-167.
(54) Jia, L., Shafirovich, V., Shapiro, R., Geacintov, N. E., and Broyde, S. (2005)
Structural and thermodynamic features of spiroiminodihydantoin damaged
DNA duplexes. Biochemistry 44, 13342-13353.
266
FUTURE DIRECTIONS
267
Future Directions
Introduction
The work in this thesis describes the genotoxic and mutational properties for the
majority of the end products of guanine oxidation by peroxynitrite. Additionally, the
significant involvement of DNA polymerase V in the replication of several of these
products was demonstrated. Potential future studies are outlined below.
Mutational Properties of Other Guanine Oxidation Products
Thus far, the mutagenesis studies of guanine oxidation products have focused on
the most chemically stable products due to the availability of procedures for the
preparation of these lesions and their ability to survive current genome construction
conditions. Several compounds discussed in Chapter 1, notably 8-nitroguanine (8-NO2G), dehydroguanidinohydantoin (DGh), and 4-hydroxy-2,5-dioxo-imidazolidine-4carboxylic acid (HICA) are of particular interest. Suzuki et al. have developed a method
for the preparation of 8-NO2-G in oligonucleotides and found that the lesion is more
stable (t1/2 = 31 h, 25 °C) than previously reported (1). The stability exhibited by this
lesion is sufficient for use in our M13 system. The stability of DGh is too low for current
genome construction conditions; however, the conditions could be improved to allow
study of this lesion. For example, the ligation step is currently 2 h at 16 °C, but ligation
reactions may also be carried out at 4 °C. Also, differences in stability as a result of
sequence context have been noted for 8-NO2-G, perhaps in the appropriate sequence
context the stability of DGh will improve. Study of HICA is precluded by the lack of a
method for preparing oligonucleotides containing the lesion in amounts sufficient for
study. At this point, we have successfully synthesized a protected 2’-deoxynucleoside
precursor of HICA. Completion of the synthesis is dependent on successful removal of
the protecting groups.
Structural Studies
Little is known about the mechanisms by which polymerases bypass DNA
lesions. NI and Ca both have the stability necessary for X-ray crystal studies, and a study
of a translesion synthesis polymerase cocrystallized with a primer/template containing
268
Future Directions
one of these lesions would provide valuable mechanistic data.
We are currently
collaborating with Dinshaw Patel at Memorial Sloan-Kettering to obtain a structure of a
primer/template containing NI complexed with the bypass polymerase Dpo4.
Repair of Guanine Oxidation Products
Quantitative in vitro repair studies of the lesions discussed in this thesis are
lacking and only one in vivo study has been reported (2). The effect of repair enzymes
and their complexes on the in vivo mutational properties of guanine oxidation products
are unknown.
Given the availability of methods for preparing oligonucleotides
containing guanine oxidation products and for purifying many repair enzymes, in vitro
studies easily could be completed. Use of a double-stranded vector containing a sitespecific DNA lesion could allow determination of the effect of repair enzymes in vivo.
In Vivo Detection
The relevance of guanine oxidation products ultimately depends on the detection of these
lesions in vivo. So far, 8-oxoG, 8-NO2-G, and Sp have been detected in cells. The recent
discovery of Sp was facilitated by knocking out the repair enzyme, Nei (2). Discovery of
other guanine oxidation products may be dependent on more sensitive detection methods
or on the identification of enzymes that show activity for specific lesions. The detection
of Sp proves that at least some of the chemistry responsible for guanine oxidation product
formation does occur in vivo. Strictly from a chemical perspective, the formation of the
other lesions in vivo is plausible. Given the results presented in this thesis, an efficient
repair system for these oxidation products may exist to combat the toxic and mutagenic
effects and likely would need to be inactivated to allow detection and quantification of
the DNA lesions.
References
(1) Suzuki, N., Yasui, M., Geacintov, N. E., Shafirovich, V., and Shibutani, S. (2005)
Miscoding events during DNA synthesis past the nitration-damaged base 8nitroguanine. Biochemistry 44, 9238-9245.
269
Future Directions
(2) Hailer, M. K., Slade, P. G., Martin, B. D., and Sudgen, K. D. (2005) Nei Deficient
Escherichia coli Are Sensitive to Chromate and Accumulate the Oxidized
Guanine Lesion Spiroiminodihydantoin. Chem. Res. Toxicol. 18, 1378-1383.
270
WILLIAM L. NEELEY
wlneeley@alum.mit.edu
EDUCATION
Ph.D., Biological Chemistry, Massachusetts Institute of Technology, Cambridge, MA, February 2006
Advisor: Professor John M. Essigmann
A.B., Chemistry (summa cum laude), Washington University, St. Louis, MO, May 2001
Advisor: Professor John-Stephen Taylor
Diploma, Bellarmine College Preparatory, San Jose, CA, May 1997
AWARDS AND HONORS
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German Exchange Program, Northeastern Section of the American Chemical Society Younger
Chemists Committee and the Gesellschaft Deutcher Chemiker Jungchemikerforum (2005)
Kluwer Poster Prize, Northeastern Section of the American Chemical Society 6th Annual
Northeastern Student Chemistry Research Conference (2004)
Massachusetts Institute of Technology Presidential Graduate Fellow (2001 – 2002)
John C. Sowden Prize in Chemistry for the Outstanding Graduating Senior, Washington U. (2001)
Sigma Xi, Associate Member, Washington U. Chapter (2001)
Hypercube Scholar “For scholastic excellence in chemistry” (2001)
Howard Hughes Medical Institute/Washington U. Undergraduate Research Fellow (2000)
American Chemical Society Outstanding Junior Chemistry Student (2000)
RESEARCH EXPERIENCE
2002 – Present
Graduate Research Assistant, Department of Chemistry, MIT, Cambridge, MA
Advisor: Professor John M. Essigmann
• Developing chemical methods for the synthesis and incorporation into DNA
of nucleosides produced by peroxynitrite oxidation.
• Evaluating the genotoxicity and mutagenic potential of DNA lesions in vivo
resulting from peroxynitrite oxidation.
• Determining the effect of SOS inducible DNA polymerases on the bypass
and mutational signature of oxidized DNA lesions.
2000 – 2001
Undergraduate Research Assistant, Department of Chemistry, Washington
University, St. Louis, MO
Advisor: Professor John-Stephen Taylor
• Prepared four novel photoproduct analogs within oligodeoxynucleotides.
• Probed the mechanism of T7 DNA polymerase bypass of DNA
photoproducts. Results confirmed the hypothesis that the 5’ moiety of DNA
(6-4) and Dewar photoproducts are within the active site of the polymerase
during primer extension opposite the photoproducts.
RELEVANT SKILLS
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Laboratory: multi-step organic synthesis, 1D and 2D NMR spectroscopy, UV-vis spectroscopy,
IR spectroscopy, MALDI-TOF mass spectrometry, HPLC, agarose and polyacrylamide gel
electrophoresis, autoradiography and phosphorimagery, nucleic acid synthesis and purification,
protein purification, bacteria and bacteriophage culture, molecular biology
Computer: computational chemistry (molecular mechanics, semi-empirical, and ab initio
calculations)
Machine shop: mill and lathe operations
PROFESSIONAL SOCIETIES
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•
American Chemical Society (Chemical Toxicology Division)
Society of Presidential Fellows (Massachusetts Institute of Technology)
271
PROFESSIONAL ACTIVITIES
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Graduate Student Representative, Institute Committee on Assessment of Biohazards, MIT
(2004 – 2006)
Member, Biological Engineering Division Environment, Health, and Safety Committee, MIT
(2003 – present)
Member, Department of Chemistry Environment, Health, and Safety Committee, MIT
(2002 – present)
Environment, Health, and Safety Representative, Essigmann Laboratory, MIT (2002 – present)
Teaching Assistant Trainer, Department of Chemistry, MIT (2002 – 2005)
TEACHING EXPERIENCE
Department of Chemistry, Massachusetts Institute of Technology, Cambridge, MA
• Teaching Assistant, Biological Chemistry Laboratory. Supervised and instructed students in the
laboratory component of the course. (February 2002 – May 2002)
• Teaching Assistant, Biological Chemistry. Instructor for two recitation sections. (August 2001 –
December 2001)
PUBLICATIONS
1. Henderson, P. T. *; Neeley, W. L.*; Delaney, J. C.; Gu, F.; Niles, J. C.; Hah, S. S.; Tannenbaum,
S. R.; Essigmann, J. M. Urea Lesion Formation in DNA as a Consequence of 7,8-Dihydro-8oxoguanine Oxidation and Hydrolysis Provides a Potent Source of Point Mutations. Chemical
Research in Toxicology, 2005, 18, 12-18. (* These authors contributed equally to the manuscript)
2. Neeley, W. L.; Delaney, J. C.; Henderson, P. T.; Essigmann, J. M. In Vivo Bypass Efficiencies
and Mutational Signatures of the Guanine Oxidation Products 2-Aminoimidazolone and 5Guanidino-4-nitroimidazole. Journal of Biological Chemistry, 2004, 279, 43568-43573.
3. Neeley, W. L.; Henderson, P. T.; Essigmann, J. M. Efficient Synthesis of the Guanine OxidationNitration Product 5-Guanidino-4-nitroimidazole: Generation by a Post-Synthetic Substitution
Reaction. Organic Letters, 2004, 6, 245-248.
4. Henderson, P. T.; Delaney, J. C.; Muller, J. G.; Neeley, W. L.; Tannenbaum, S. R.; Burrows, C.
J.; Essigmann, J. M. The Hydantoin Lesions Formed From Oxidation of 7,8-Dihydro-8oxoguanine are Potent Sources of Replication Errors in Vivo. Biochemistry, 2003, 42, 92579262.
SELECTED ORAL PRESENTATIONS
1. Neeley, W. L.; Henderson, P. T.; Delaney, J. C.; Gu, F.; Niles, J. C.; Hah, S. S.; Tannenbaum, S.
R.; Essigmann, J. M. Formation of 2-Aminoimidazolone and Urea in DNA as a Consequence of
7,8-Dihydro-8-oxoguanine Oxidation Provides Potent Sources of Point Mutations in Vivo.
Frühjahrssymposium 2005. April 7, 2005, Berlin, Germany.
2. Neeley, W. L.; Henderson, P. T.; Delaney, J. C.; Essigmann, J. M. Synthesis and in Vivo
Mutational Properties of the Guanine Oxidation-Nitration Product 5-Guanidino-4-nitroimidazole.
228th American Chemical Society National Meeting. August 22, 2004, Philadelphia, PA.
3. Neeley, W. L.; Taylor, J.-S. Preparation and Characterization of UV Photoproduct Analogs NMethyl-T[6-4]T, 5-Methyl-C[6-4]T, and Their Respective Dewar Isomers as Probes for the
Mechanism of UV-Induced Mutagenesis. Pew Midstates Science and Mathematics Consortium
Undergraduate Research Symposium in the Physical Sciences and Mathematics. November 5,
2000, Washington U., St. Louis, MO.
SELECTED POSTER PRESENTATIONS
1. Neeley, W. L.; Henderson, P. T.; Delaney, J. C.; Essigmann, J. M. In Vivo Bypass Efficiencies
and Mutational Signatures of the Guanine Oxidation Products 2-Aminoimidazolone and 5Guanidino-4-nitroimidazole. American Society for Biochemistry and Molecular Biology Annual
Meeting and 8th International Union for Biochemistry and Molecular Biology Conference. June
12-16, 2004, Boston, MA.
2. Neeley, W. L.; Henderson, P. T.; Delaney, J. C.; Essigmann, J. M. Synthesis and in Vivo
Mutational Properties of the Guanine Oxidation-Nitration Product 5-Guanidino-4-nitroimidazole.
Northeastern Section of the American Chemical Society 6th Annual Northeastern Student
Chemistry Research Conference. April 24, 2004, Boston University, Boston, MA.
272