Response of DNA repair and replication systems to
exocyclic nucleic acid base damage
by
Nidhi Shrivastav
B.Tech., M.Tech., Biochemical Engineering and Biotechnology (2005)
Indian Institute of Technology, Delhi
Submitted to the Department of Biological Engineering
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biological Engineering
ARCH[VES
at theASSACHUSETTS
Massachusetts Institute of Technology
INSTITUTE
APR r 2 2012
February 2012
UBRARIES
@ 2012 Massachusetts Institute of Technology
All rights reserved
Signature of Author
Department of Biological Engineering
February, 2012
Certified by
John M. Essigmann
William R.and Betsy P. Leitch Professor of Toxicology, Chemistry, and Biological Engineering
Associate Head, Department of Chemistry
Thesis supervisor
Accepted by
Forest M. White
Co-chairman, Department Committee on Graduate Students
Associate Professor of Biological Engineering
This thesis has been examined by a thesis advisory committee consisting of the following
members:
Peter C.Dedon
Underwood Prescott Professor of Toxicology and Biological Engineering
Deputy Director, MIT Center for Environmental Health Sciences
Committee Chair
Bevin P. Engelward
Associate Professor of Biological Engineering
Edward L. Loechler
Professor of Biology, Boston University
V
John M. Essigmann
William R.and Betsy P.teit/ch Professor of-oxicology, Chemistry, and Biological Engineering
Associate Head, Department of Chemistry
Thesis supervisor
Response of DNA repair and replication systems to
exocyclic nucleic acid base damage
By Nidhi Shrivastav
January 13, 2012, in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biological Engineering
Abstract
Genomes experience an often hostile environment that creates a vast array of damages that
can give rise to myriad biological outcomes. Fortunately, cells are equipped with networks such
as direct reversal, base excision repair, nucleotide excision repair, homologous recombination,
and translesion synthesis that help preserve informational integrity.
The first part of this dissertation focuses on whether or not bulky alkyl lesions at the N2 atom of
guanine are addressed in vivo by the DinB bypass polymerase. In the work described herein, a
collection of N2 -guanine lesions was inserted in single-stranded M13 genomes and evaluated in
strains possessing or lacking DinB via the competitive replication and adduct bypass (CRAB) and
restriction endonuclease and postlabeling (REAP) assays. It was found that DinB could in fact
bypass the N2 -furfuryl-guanine lesion and its saturated homolog in vivo.
The second part of this work describes how we systematically investigated the role that the
distance from an origin of replication may have in the mutagenesis of an adduct. Our
hypothesis was that a lesion farther from the origin of replication would be less mutagenic
since it would be afforded more time for detection and removal before the replicative
polymerase traversed it, fixing the mutation. We inserted 0-methylguanine in single-stranded
M13 genomes at different distances from the origin of replication and analyzed progeny phage
by the REAP assay. Our findings were in contrast with the hypothesis; a higher mutation
frequency was obtained at the site distal from the origin of replication. Alternative hypotheses
and future experiments are discussed as part of this work.
The third part of this dissertation seeks to expand the spectrum of known substrates for the
enzyme AIkB, which mediates direct reversal of DNA damage. AlkB is an iron- and CCketoglutarate-dependent dioxygenase that is part of the adaptive response in E. coli, and has
homologs in many species. On basis of in vitro data we created the hypothesis that the N2
guanine lesions as well as 6-methyladenine would be substrates for the enzyme AlkB in vivo.
We found, however, in this case the in vitro results did not predict the biology observed in cells.
Thesis supervisor: John M. Essigmann
Title: William R. and Betsy P. Leitch Professor of Toxicology, Chemistry, & Biological Engineering
Associate Head, Department of Chemistry
Acknowledgements
"'It takes a village to raise a child"
whose book was inspired by this proverD.
The person instrumental to the completion of this work is my advisor, Dr. John Essigmann.
Thank you John, for giving me the opportunity to perform research on several projects in your
laboratory. John has given me the freedom to do independent work, while providing valuable
insight and input as and when needed. He has been a great mentor, keeping in mind not just
the research experience a graduate student should gain, but also the education a graduate
student should receive that would enable her to succeed in further endeavors, be that a
teaching opportunity, a mentoring opportunity in the form of a UROP, or presenting one's
research in public forums. John has been a great boss to work for, an act that I will surely find
tough to follow as I step out of student life.
I would like to express my gratitude to my committee members, Dr. Peter Dedon, Dr. Bevin
Engelward, and Dr. Edward Loechler for their time and effort towards the work described in this
thesis. They have analyzed my work with a keen and critical view at every step of my research.
This work would not have taken the shape it has without their in-depth advice. A special thank
you to Dr. Loechler for making several bike trips across the river for my meetings.
The Essigmann laboratory has been a great place to work in. The people have made my five
years in the lab a very pleasant, fun, and collaborative experience. A heartfelt thank you goes
out to Jim Delaney, for taking me under his tutelage and teaching me the intricacies of
experimental lab work. You have truly showed me how to critically prepare, plan, and analyze a
project from start to finish in the best possible manner. I feel honored to have your unfailing
mentorship during my stay at MIT. Deyu and Bogdan, you have been an inspiration. I have
learned from you not just experimental techniques and new research insights, but also what
motivation and passion towards science can achieve. Beth and Sarah worked with me on two
different projects and hopefully I was able to impart a level of knowledge that would compare
to what I learned from them. Nicole, Leslie, Bob, Vipender, and Steven - thank you for making
the lab a fun place to be in!
In my five-year stay in the Essigmann laboratory, I have had the fortune of spending time with a
generation of researchers, without whom I would not have completed this journey. Sreeja,
thank you for being my unofficial MIT mentor even before I stepped foot on campus. Charles
and Lauren, thank you for all your advice over the years, fun conversations, and help with
editing my thesis. I would also like to thank Jeannette, Kyle, Peter, Will, Francis, Sarah, Eunsuk,
Pei-sze, Alfio, John, Tang, and Ann for your support over the years.
A graduate student's work is often incomplete without collaboration. I would like to thank Dr.
Jamie Foti from Dr. Graham Walker's laboratory for help in the creation of DinB strains. I would
also like to thank Dr. Daniel Jarosz for answering my persistent questions on the DinB project.
Lastly, I would like to thank all members of the Engelward laboratory for technical help,
ramaraderie. and several interestin2 conversations over lunch.
been amle to spena nailT a uecaue dL IVIiI WILIiUUL Uman01I1I5 a3uppunl
III i Iny
family back home. To Mom and Dad, for bringing me up with the belief that I could achieve
anything in life, and for instilling in me the love of learning and of science at a very young age.
Thank you Urvi, my sister, for your loving company and for sportingly being at the receiving end
of many of my jokes. A warm thank you to my parents-in-law, who have always showered me
with their blessings and are extremely proud of what I have now achieved.
I woula not nave
The one person who has been with me through this period day in and day out is Saurabh, my
husband. No words can thank you enough for your emotional support through what has been a
roller-coaster journey. I have always had your encouragement and your belief in my abilities to
show me through obstacles in my work. Thank you for being here with me.
My life in the United States and at MIT would have been a very different experience had it not
been for the many close friends I made here. Abhinav, thank you for many, many, many
discussions on everything under the sun, and for your unfailing support and presence. Thank
you Bonnie for being a very understanding and accommodating roommate and friend. Alex,
James, Brandon, Ta, Karen, Sharon, and Asha, thank you for more entertaining conversations
than I can remember, for your selfless kindness, and for memories that will last a lifetime. Last
but not the least, I would like to thank Sonia, my first friend when I came to the U.S.A. I hope
our friendship and the good times last through the years.
I would like to thank several admin personnel, Dalia Fares, Mary Files, Kim-Bond Schaefer, Aran
Parillo, and the International Students Office for making my transition and stay at MIT a smooth
and comfortable one. Lastly, I would like to thank the NIH for funding this work.
Table of Contents
Committee Page
Abstract
Abbreviations
List of Figures
List of Tables
Chapter 1: Introduction
1.1 Introduction
1.2 The SOS response
1.3 Transcription-coupled translesion synthesis
1.4 References
Chapter 2: Chemical biology of mutagenesis and DNA repair: cellular responses to DNA
alkylation
2.1 Abstract
2.2 Introduction
2.3 0 6-Methylguanine and 0 6-ethylguanine
2.4 04-Methylthymine
2.5 02-Methylcytosine and 0 2-methylthymine
2.6 Methylphosphotriesters
2.7 N1-Methyladenine and N1-ethyladenine
2.8 N3-Methyladenine
2.9 N7-Methyladenine
2.10 N1-Methylguanine
2.11 N3-Methylguanine
2.12 N7-Methylguanine and its degradation products
2.13 N3-Methylcytosine and N3-ethylcytosine
2.14 N3-Methylthymine
2.15 8-Methylguanine
2.16 1,N6-Ethenoadenine and 1,N -ethanoadenine
2.17 1,N2-Ethenoguanine and 3,N2 -ethenoguanine
2.18 3,N4-Ethenocytosine
2.19 Perspective
2.20 Acknowledgements
2.21 Figures
2.22 Tables
2.23 References
Chapter 3:
3.1
3.2
3.3
DinB bypasses N2 -dG lesions in vivo
Introduction
Materials and methods
Results
R R 1 Crpitinn nf DinR- rll -strains
3.3.3 KEAP assay
3.4 Discussion
3.5 Figures
3.6 Tables
3.7 References
Chapter 4: DNA lesion placement relative to the origin of replication and polymerase
obstacles as variables in 06-methylguanine mutagenesis in vivo
4.1 Introduction
4.2 Materials and methods
4.3 Results
4.3.1 GATC controls
4.3.2 Test of recombination between the proximal and distal sites
4.3.3 Effect of distance from the origin of replication on the mutation
frequency of O6MeG
4.3.4 Effect of genome lesion load on the mutation frequency of 06MeG
4.3.5 Effect of a blocking lesion at the proximal site on the mutation
frequency of O6 MeG at the distal site
4.4 Discussion and future work
4.5 Figures
4.6 Tables
4.7 References
78
79
84
93
93
95
100
104
105
111
112
121
132
132
133
134
136
137
138
149
161
165
2
Chapter 5: Investigating the role of AIkB in the repair of N -dG lesions and the
demethylation of m6A in vivo
5.1 Introduction
5.2 Materials and methods
5.3 Results
3.3.1 CRAB assay
3.3.2 REAP assay
5.4 Discussion
5.5 Figures
5.6 Tables
5.7 References
173
Appendix A: Publication: Delaney,J.C., Gao,J., Liu,H., Shrivastav,N., Essigmann,J.M., and
KoolE.T. (2009) Efficient replication bypass of size-expanded DNA base pairs in bacterial
cells. Angewandte Chemie International Edition, 48, 4524-4527
201
174
177
186
186
187
187
191
195
197
Abbreviations
4-NQO: 4-nitroquinoline-1-oxide
8-oxo-dG: 8-oxo-7,8-dihydro-2'-deoxyguanosine
AP:
APNG:
APOBEC:
ATL:
ATP:
BaP:
BCNU:
BER:
BSA:
Ci:
cm:
CRAB:
DMF:
DMS:
DNA:
dNTP:
DTT:
E. coli:
e2G:
ENU:
FF:
FTO:
g:
h:
kb:
kV:
LB:
M:
mIG:
m2G:
m3C:
m3T:
MCS:
MePT:
mg:
MGMT:
Min:
Apurinic or apyrimidinic site; abasic site
Alkylpurine-DNA-N-glycosylases
Apolipoprotein Bediting complex
Alkyltransferase-like protein
Adenosine triphosphate
Benzo[a]pyrene
Bis-Chloroethylnitrosourea
Base excision repair
Bovine serum albumin
Curies
Centimeter
Competitive replication of adduct bypass assay
Dimethylformamide
Dimethylsulfate
Deoxyribonucleic acid
Deoxyribonucleotide triphosphate
Dithiothreitol
Escherichia coli
2-Ethylguanine
N-ethyl-N-nitrosourea
2-Furan-2-yl-methylguanine or N2 -furfuryl-dG
Fat mass and obesity associated gene
Grams
Hours
Kilobases
Kilovolts
Luria broth
Molar
1-Methylguanine
2-Methylguanine
3-Methylcytosine
3-Methylthymine
Multiple cloning site
Methylphosphotriesters
Milligrams
06-methylguanine-DNA-methyltransferase
Minutes
ml:
mm:
mM:
mmol:
Milliliter
Millimeters
Millimolar
Millimoles
MMR-
MAi-match rpnair
MNNG:
MNU:
MUG:
MW:
2
N -CEdG:
N2-dG:
NER:
NFZ:
0 4 MeT:
o MeG:
or:
PAGE:
PCR:
PEG:
pmol:
PNK:
REAP:
RF:
RNA:
ROS:
rpm:
SAM:
SSB:
TCR:
TE:
THF:
TLS:
U:
uCi:
ug:
ul:
uM:
V:
N-methyl-N-nitro-N-nitrosoguania ine
N-methyl-N-nitrosourea
Mammalian uracil-DNA-glycosylase
Molecular weight
N2-(1-carboxyethyl)-2'-deoxyguanosine
N2 group of 2'-deoxyguanosine
Nucleotide excision repair
Nitrofurazone
04 -Methylthymine
06-Methylguanine
Origin of replication
Polyacrylamide gel electrophoresis
Polymerase chain reaction
Polyethylene glycol
Picomoles
Polynucleotide kinase
Restriction endonuclease and postlabeling assay
Replicative form
Ribonucleic acid
Reactive oxygen species
Rotations per minute
S-adenosyl-L-methionine
Single-stranded binding protein
Transcription-coupled repair
Tris-EDTA buffer
2-Tetrahydrofuran-2-yl-methylguanine
Translesion synthesis
Enzyme unit
Microcuries
Micrograms
Microliter
Micromolar
Volts
wt: Wild-type
Y: Aminoethoxyethyl ether group
Q: Ohms
EC: 3,N4 -Ethenocytosine
List of Figures
Figure 2.1:
Pathways by which DNA damaging agents induce biologically relevant events.
58
Figure 2.3:
Structures of DNA alkylation lesions.
Structures of lesions used in the DinB study.
bU
Overview of the REAP and CRAB assay.
Bypass efficiencies of N2-dG lesions in DinB+ and DinB- cells as determined by
the CRAB assay.
Mutagenicity of the N2 -dG lesions as determined by the REAP assay.
101
149
Figure 4.2:
Experimental Outline.
Map of M13mp7(L2) and M13EH.
Figure 4.3:
REAP assay.
151
Figure 4.4:
152
Figure 4.5:
Structure of hairpin inserted in the modified M13 vector.
Complete sequence of M13EH.
Figure 4.6:
GATC controls.
155
Figure 4.7:
Test of possible recombination between the distal and proximal sites.
Graphical representation of mutation frequency data.
Mutation frequency of O6MeG as a function of distance from the ori in an
Ada+ Ogt- repair background in GXG and AXG nearest neighbor sequence
contexts.
Effect of lesion load on mutation frequency in an Ada+ Ogt- repair background
in GXG and AXG nearest neighbor sequence contexts.
Mutation frequency of O6MeG (m6G) placed at the distal site when a known
replication block (m3T, THF) is placed at the proximal site.
156
Structure of lesions in the AlkB study.
Bypass efficiencies in AlkB+ and AlkB- cells as determined by the CRAB assay.
Mutagenicity results in AlkB+/- SOS+/- cells as determined by the REAP assay.
Bypass efficiencies of N2-dG lesions in AlkB+ and AlkB- (both DinB-) cells as
determined by the CRAB assay.
191
Figure 3.1:
Figure 3.2:
Figure 3.3:
Figure 3.4:
Figure 4.1:
Figure 4.8:
Figure 4.9:
Figure 4.10:
Figure 4.11:
Figure 5.1:
Figure 5.2:
Figure 5.3:
Figure 5.4:
100
102
103
150
153
157
158
159
160
192
193
194
List of Tables
Table 2.1:
Table 3.1:
Table 3.2:
Table 4.1:
Table 4.2:
Table 4.3:
Table 4.4:
Table 5.1:
Table 5.2:
Table 5.3:
Mutagenicity, genotoxicity, and repairability of DNA alkylation lesions.
Bypass efficiencies of N2 -dG lesions in DinB+ and DinB- cells as determined by
the CRAB assay.
Mutagenicity of the N2-dG lesions as determined by the REAP assay.
61
M13 genes and their functions.
Possible candidates for replication-blocking lesions to be placed at the
proximal site.
Mutation frequency data in a GXG nearest neighbor sequence context.
Mutation frequency data in an AXG nearest neighbor sequence context.
161
Bypass efficiencies of N2-dG lesions in AlkB+ and AlkB- cells (both DinB+) as
determined by the CRAB assay.
Bypass efficiencies of N2-dG lesions in AlkB+ and AlkB- cells (both DinB-) as
determined by the CRAB assay.
Mutagenicity results in (a)AlkB+/SOS-, (b) AlkB-/SOS-, and (c) AlkB-/SOS+ cells
as determined by the REAP assay.
195
104
104
162
163
164
195
196
Chapter 1:
Introduction
1.1 Introduction
Genomes, both RNA and DNA, experience an often hostile environment. Reactive oxygen
species, reactive nitrogen species, organic electrophiles, reactive metals, ionizing and nonionizing radiations impinge upon genetic material creating a vast array of damages that can give
rise to myriad biological outcomes. These types of DNA and RNA damage could influence
replicative fidelity, affect the ability of a cell to avoid acute toxicity from strand breaks, alter
patterns of epigenetic modification, and disrupt transcriptional programs by interfering with
normal transcription factor - DNA interactions [1]. Although DNA damage and mutation has the
benefit of providing a driving force for evolution, it is also the basis of many genetic diseases.
Fortunately, cells are equipped with molecular networks that help preserve informational
integrity. Foremost amongst these networks are the hundreds of repair proteins, most of
which are enzymes, which reverse DNA damage and restore the structural and informational
integrity of the genome. Nearly all of these repair factors act upon DNA genomes, although
recent exciting data suggest that RNA molecules may also be substrates [2;3]. Repair systems
fall into three general categories. The first involves direct reversal, the second involves base
excision repair, and the third involves nucleotide excision repair. These repair systems are
complemented by recombinational networks that allow tolerance of the damage until such
time as the damage can be physically removed from the genome.
A third general
genoprotective strategy cells use to counter the effects of DNA damaging agents is to sequester
the electrophilic derivatives of classical DNA and RNA damaging agents; glutathione
transferases, epoxide hydrolases, and other phase 11systems exemplify this tactic.
When DNA damage, in particular, occurs, it is well established that a plethora of structurally
different lesions forms within the genome. Although it is known that nucleic acid damaging
agents cause mutations, until recently it was difficult to dissect the relative involvement of any
individual lesion to the spectrum of mutations that occurs in cells. Similarly, it has been difficult
to assign genotoxic properties to specific lesions in the genome. The approach taken in this
laboratory and described in this dissertation involves the synthesis of easily characterizable
oligonucleotides containing single DNA lesions. By using recombinant DNA techniques, the
genome of the M13 virus is reengineered to situate the oligonucleotide at a specific site. The
biological properties of the lesion can be studied in any sequence context at many different
sites in the overall viral genome. In order to study these properties, the genome is introduced
into an E. coli cell under conditions such that only one genome containing a single lesion is
introduced per cell. The viral genome is allowed to replicate in a milieu containing the native
polymerases and repair systems of the host cell.
Progeny are isolated and analyzed by a
technique in which the area of the genome among progeny that originally contained the adduct
is interrogated.
Mutants are enumerated, providing a chemical 'rulebook' for any genetic
change that occurred within the cell [4]. If the genome is introduced into a cell containing an
altered repair system, one is able to define the precise role that repair protein has in protecting
against that specific type of damage. Similarly, replication of a lesion-containing genome in a
cell with altered polymerase content allows one to evaluate the role that specific polymerases
play in the bypass or mutagenesis of specific DNA lesions. Taken together, the aforementioned
technology helps to create a contribution to a biochemical 'periodic table' that defines the
properties of specific DNA lesions [5].
The first major topic addressed in this thesis research involves the role of bypass polymerases in
the tolerance of genotoxic damage. When E. coli is challenged with many DNA damaging
agents, some of the lesions formed block DNA replication resulting in the formation of
daughter-strand gaps downstream of the replication fork. RecA binds to this single-stranded
DNA, creating a RecA nucleofilament that cleaves LexA, a transcriptional repressor. Thus,
cleavage of LexA induces several genes, including the damage inducible genes [6]. One of them
is DinB or pol IV in E. coli (Dpo4 in Sulfolobus solfataricus, pol K in mammals). Interestingly
every life form contains a homolog of this protein [7]. A substantial advance occurred in 2006
with the discovery that DinB could bypass, in vitro, the N2 -furfuryl-guanine lesion more
accurately and efficiently than an unmodified guanine [8]. An unanswered question is whether
or not these bulky lesions at the N2 atom of guanine are addressed in vivo by the DinB bypass
polymerase. The site specific mutagenesis and adduct bypass technology described above is
well suited to answer this question. In the work described herein, a collection of N -guanine
lesions is evaluated in strains possessing or lacking DinB.
The data from in vivo studies
corresponds well with previous work in vitro.
The second part of this work explores the effect of the distance from the origin of replication on
the mutagenesis of a lesion. Earlier work by Delaney et al. showed that a certain repair protein,
Ada, showed significant sequence context dependence in its ability to counter the effects of O6_
methylguanine in a viral genome [9;10]. These data and others [11-13] showed clearly that
there were context effects that influenced mutagenic spectra. Another variable that has not as
yet been systematically investigated is the role that the distance from an origin of replication
may play in the qualitative and quantitative features of mutagenesis of an adduct. If, for
example, an adduct is close to an origin, one would expect that repair enzymes would have
little opportunity to be able to reverse the damage before the polymerase traverses it, fixing
the mutation.
A more distal lesion, by contrast, might be less mutagenic because repair
proteins would be more effective in detecting and removing the damage. It is also possible that
repair proteins may move in concert with polymerases helping to clear DNA of lesions
immediately ahead of the replication fork. In this scenario, one might imagine that repair
proteins are consumed or otherwise derailed with the result that a distal lesion may be more
mutagenic than a proximal one.
One objective of the work described herein is to try to
determine whether the mutation frequency of a lesion, in identical sequence contexts but at
different distances from an origin of replication, differs. This is an important question as it
helps to define the mutational landscape called a mutational spectrum.
Finally, this laboratory and others have had a long standing interest in the iron- and alphaketoglutarate-dependent dioxygenases. As with Ada described above, this group of enzymes
effectuates a direct reversal of damage to DNA and RNA. For example, a 1-methyladenine will
be oxidized to a transient 1-hydroxymethyladenine that then decomposes to adenine plus
formaldehyde [14]. As a second example, an ethenoadenine is epoxidized at the etheno bridge
and then the epoxide is removed as the dialdehyde glyoxal, again liberating unmodified
adenine [15]. As part of a systematic effort to understand the substrate specificity of this
remarkable group of enzymes, the third part of this dissertation focuses on a number of N2_
guanine lesions as well as 6-methyladenine. On basis of in vitro data we created the hypothesis
that the lesions would be substrates for the enzyme AlkB in vivo. We found however in this
case the in vitro results did not predict the biology observed in cells.
None of the
aforementioned types of damage can be included in the set of lesions now known to be
addressed by the AlkB class of repair proteins.
The common thread among the three projects described above is the assay, which is an engine
for characterizing important biological effects of hazardous agents. Our assay allows us to
study the effect of lesions at a low dose of one lesion per cell, as compared to in vitro systems
in which extreme conditions and doses are often used. While in vitro experiments are great for
characterizing chemical mechanisms, our assay enables one to observe the effect of a lesion in
living cells without overwhelming the cells with exposure to damaging agents. This thesis does
exactly that. While most of the literature pertinent to the individual studies is reviewed in
subsequent chapters, two DNA damage responses that are major players in the work described
herein are briefly summarized below.
1.2 The SOS response
The first experiment that hinted at the existence of what is now known as the SOS response
was done by Weigle in 1953. He discovered that UV-irradiated A phage could be rescued by
infection into UV-irradiated E. coli [16].
A second study by Witkin, over a decade later,
speculated that the rescue of UV-irradiated A phage, along with a concomitant increase in
point mutations, was the result of a damage-induced DNA repair system in bacterial cells [17].
This "mutation-prone" replication mechanism involving the lexA and recA gene functions that
could account for UV-induced mutations of Aphage and E. coli was subsequently named "SOS
repair" by Radman [18].
DNA damage caused by exposure to UV-irradiation or to chemicals can trigger the SOS response
[19;20]. The primary aim of the SOS response is to rescue cells from death caused by the
blocking of replication fork progression by unrepaired lesions or lesions undergoing repair
[21;22].
This system, regulated by both the LexA transcriptional repressor and the RecA
recombinase, controls the induction of over 40 genes [23;24]. A majority of these genes are
involved in error-free DNA repair pathways such as base excision repair (BER), nucleotide
excision repair (NER), and recombinational DNA repair [22]. However, if these pathways fail to
complete repair and restart replication, the error-prone tolerance mechanism of SOS, a process
called translesion DNA synthesis (TLS) [25], is triggered [20;26]. TLS is mediated by error-prone
polymerases, and is responsible for the large increase in mutations (~100-fold [22]) originally
observed by Weigle.
Interruption of the replication fork progression by unrepaired lesions leads to the accumulation
of regions of single-stranded DNA, which induces the SOS response. The RecA protein has a
strong tendency to form nucleoprotein filaments on single-stranded DNA [27;28], the formation
of which proceeds in the 5'43' direction at a ratio of 1 molecule RecA per 3 DNA bases. The
formation process requires dATP or ATP, but no ATPase activity; however, the disassembly
needs hydrolysis of ATP to ADP and is slower. This nucleoprotein has a coprotease activity
(RecA*), which facilitates the self-cleavage of LexA protein resulting in derepression of SOSregulated genes. The regulation is achieved by the presence of a specific 20-nucleotide-long
"SOS-box" (or LexA-box) near the promoter/operator site of each SOS-induced damage
inducible (din) or sos gene. The LexA repressor protein is bound to this box as a dimer [29],
preventing RNA polymerase binding and gene expression [20;28;301. RecA* adds an additional
layer of control to the expression of one of the SOS-induced genes, pol V (UmuD' 2C). It is
required for the conversion of UmuD to UmuD' by nicking UmuD at the Cys24-Gly25 site
[31;32], which is a prerequisite for the assembly of the final product, UmuD' 2 C. Induction of the
SOS response continues until 45-60 min after treatment of bacteria with SOS inducing agents,
with the timing of the derepression of individual din genes depending on the strength of the
LexA repressor binding with the SOS-box and on the ease with the LexA repressor is detached
from a particular SOS-box. This window of time is usually sufficient to repair most lesions.
There are three SOS-induced genes that are relevant to the work described in this manuscript,
namely, the three translesion DNA polymerases of E. coli [25]. Translesion DNA polymerases
are a class of DNA polymerases which enable tolerance of lesions that block the highly accurate
replicative DNA polymerases, until such time as the damage can be physically removed from
the genome. The process by which they do so is termed translesion synthesis (TLS). Although
these polymerases have a reduced fidelity on undamaged templates, they can replicate across
cognate lesions with great proficiency [25;33].
E. coli has three SOS-induced translesion DNA
polymerases, namely, pol II, pol IV and pol V, which are conserved across almost all domains of
life. In a wild-type E. coli cell, it is estimated that there are 30-50 molecules of pol II, 250 of pol
IV, and 15 of pol V [34;35]. Upon SOS induction, this number jumps to 350 for pol II, 2500 for
pol IV, and to 200 for pol V [35;36]. Pol 11,a B-family polymerase discovered in 1970 [37;38],
has 3' -> 5' exonuclease activity [39], and synthesizes DNA accurately [40]. Cells deficient in pol
Il do not show adverse phenotypic effects except when pol V is also missing [41]. The role of
pol 11is speculated to be in replication restart in an error-free manner. Also, double mutants of
pols 11and V are more sensitive to UV compared to cells lacking pol V alone [41]. It is also
involved in AAF-induced -2 frameshift mutagenesis [42]. Pol V is the product of the UmuDC
genes, and lacks 3'
->
5' proofreading exonuclease activity. As mentioned in the previous
section, a RecA* nucleofilament is needed for the activation of pol V TLS. Pol V is involved in
the bypass of benzo[a]pyrene (BaP), thymine-thymine cyclobutane dimers, abasic sites, and [64] photoproducts [42;43] . Pol IV is reviewed in detail in Chapter 3. These polymerases come
into play not only via the SOS response, but also when transcription is blocked due to the
presence of an unrepaired lesion as discussed below.
1.3 Transcription-coupled translesion synthesis
Transcription-coupled repair, or TCR, is known to occur when RNA polymerase is stalled at a
lesion. This stalling can trigger the activation of the NER repair pathway, as well as the TLS
pathway of lesion tolerance.
NER is divided into global-genomic NER (GG-NER) and
transcription-coupled NER (TC-NER) that differ primarily in DNA damage recognition but have
subsequent overlapping steps. In E. coli TC-NER, the transcription repair coupling factor Mfd
combines with the UvrABC exinuclease system to effect the removal of the offending lesion.
Briefly, Mfd binds to and displaces the RNA polymerase to enable the recruitment of UvrA at
the site. The subsequent steps involve lesion verification by UvrB, DNA strand dual incision by
UvrC, excision by UvrD, repair by DNA polymerase I, and ligation (see [44] for a review). It has
been recently proposed that NusA can also recruit the TC-NER pathway, specifically in the case
when transcription is blocked by N2 -guanine lesions [45]. The existence of this second coupling
factor between NER and transcription would explain why cells lacking Mfd are only mildly
sensitive to UV damage. Whether or not there exists a specific transcription-coupled BER
pathway is unclear; it is possible that the lesions that would normally be addressed by BER are
shunted through the TC-NER pathway when encountered by a blocked RNA polymerase [46].
The other response that is relevant for the work described in this dissertation is transcriptioncoupled translesion synthesis (TC-TLS).
This pathway involves the recruitment of the TLS
polymerases, described in the previous section, by the nusA protein that is associated with the
E. coli RNA polymerase [47]. NusA is thought to be associated with the RNA polymerase
throughout the elongation and termination steps of transcription, and has been shown to
interact with pol IV [47].
This role of NusA as a coupling protein of transcription and
translesion synthesis was proposed when it was found that catalytically active pol IV or pol V
were needed to suppress the temperature sensitivity of the nusAll(ts) strain. The translesion
synthesis abilities of pol IV were particularly required for this suppression. It was additionally
found that the nusAll mutant strains were highly sensitive to nitrofurazone (NFZ) and 4nitroquinolone-1-oxide (4-NQO) but not to ultraviolet (UV) radiation or methylmethane
sulfonate (MMS) at the permissive temperature. The model proposed by the authors suggests
that the N -guanine lesions generated by NFZ and NQO are efficiently bypassed by pol IV during
replication, but then go on to disrupt transcription, where pol IV is once again recruited through
a NusA mediated TC-TLS pathway [45]. The TC-TLS pathway, particularly in the context of the
N 2-guanine lesions, is relevant to the work done in Chapter 3. The recent discovery of the role
of NusA in TC-NER as well as TC-TLS demonstrates that the role these pathways play in the cell
are still in the process of being explored and understood.
1.4 References
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mutagenesis. ASM Press..
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3. Jia,G., Yang,C.G., Yang,S., Jian,X., Yi,C., Zhou,Z., and He,C. (2008) Oxidative
demethylation of 3-methylthymine and 3-methyluracil in single-stranded DNA and RNA
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4. Delaney,J.C. and Essigmann,J.M. (2006) Assays for determining lesion bypass efficiency
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7. Ohmori,H., Friedberg,E.C., Fuchs,R.P.P., Goodman,M.F., Hanaoka,F., Hinkle,D.,
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Walker,G.C., Wang,Z., and Woodgate,R. (2001) The Y-family of DNA polymerases.
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8. Jarosz,D.F., Godoy,V.G., Delaney,J.C., Essigmann,J.M., and Walker,G.C. (2006) A single
amino acid governs enhanced activity of DinB DNA polymerases on damaged templates.
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9. Delaney,J.C. and Essigmann,J.M. (1999) Context-dependent mutagenesis by DNA
lesions. Chem. Biol., 6, 743-753.
10. Delaney,J.C. and Essigmann,J.M. (2001) Effect of sequence context on 0 6-methylguanine
repair and replication in vivo. Biochemistry, 40, 14968-14975.
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21. SETLOWR.B., SWENSON,P.A., and CARRIER,W.L. (1963) Thymine dimers and inhibition
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23. Fern ndez de Henestrosa,A.R., Ogi,T., Aoyagi,S., Chafin,D., Hayes,J.J., Ohmori,H., and
Woodgate,R. (2000) Identification of additional genes belonging to the LexA regulon in
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24. Courcelle,J., Khodursky,A., Peter,B., Brown,P.O., and Hanawalt,P.C. (2001) Comparative
gene expression profiles following UV exposure in wild-type and SOS-deficient
Escherichia coli. Genetics, 158, 41-64.
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26. Echols,H. and Goodman,M.F. (1990) Mutation induced by DNA damage: a many protein
affair. Mutat.Res., 236, 301-311.
27. Arenson,T.A., Tsodikov,O.V., and Cox,M.M. (1999) Quantitative analysis of the kinetics
of end-dependent disassembly of RecA filaments from single-stranded DNA. J Mol Biol,
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28. Schlacher,K., Pham,P., Cox,M.M., and Goodman,M.F. (2006) Roles of DNA polymerase V
and RecA protein in SOS damage-induced mutation. Chem Rev, 106, 406-419.
29. Thliveris,A.T., Little,J.W., and Mount,D.W. (1991) Repression of the E.coli recA gene
requires at least two LexA protein monomers. Biochimie, 73, 449-456.
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cleaved by the RecA protein. Cell, 27, 515-522.
31. Burckhardt,S.E., Woodgate,R., ScheuermannR.H., and Echols,H. (1988) UmuD
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RecA. Proceedings of the National Academy of Sciences, 85, 1811-1815.
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cleavage of UmuD protein and SOS mutagenesis. Proc.Natl.Acad.Sci.U.S.A, 85, 18061810.
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Chapter 2:
Chemical biology of mutagenesis and DNA repair: cellular responses
to DNA alkylation
A version of this chapter has been previously published and is reprinted here with the
permission of the publisher:
Shrivastav,N., Li,D., and Essigmann,J.M. (2010) Chemical biology of mutagenesis and DNA
repair: cellular responses to DNA alkylation. Carcinogenesis, 31, 59-70.
2.1 Abstract
The reaction of DNA damaging agents with the genome results in a plethora of lesions,
commonly referred to as adducts. Adducts may cause DNA to mutate, they may represent the
chemical precursors of lethal events, and they can disrupt expression of genes. Determination
of which adduct is responsible for each of these biological endpoints is difficult, but this task
has been accomplished for some carcinogenic DNA damaging agents. Here, we describe the
respective contributions of specific DNA lesions to the biological effects of low molecular
weight alkylating agents.
2.2 Introduction
DNA damage can be caused by radiation, by organic and inorganic chemical agents and by
enzymes that have the roles of promoting natural methylation and deamination, such as
members of the S-adenosylmethionine - dependent methyltransferases [1], the activation
induced deaminase (AID) and the apolipoprotein B editing complex (APOBEC) [2;3]. Because
DNA is abundantly equipped with nucleophilic sites, reaction with extracellularly generated and
endogenously produced electrophiles results in an amazingly diverse array of covalent
chemical-DNA adducts. These lesions compromise cellular welfare in three major ways (Figure
2.1).
First, misreplication or misrepair of the lesions triggers mutations, which can be the
initiating lesions of genetic diseases, including cancer. Second, the lesions can jeopardize the
epigenetic program imprinted by natural enzymatic DNA modifications. Finally, the lesions can
block RNA and DNA polymerases and can lead directly or indirectly to DNA strand breaks, which
tend to be lethal in most cells. The biological importance of DNA damage is evidenced by the
28
large commitment of the genome to protection of informational integrity; such genoprotective
networks include electrophile scavengers, recombination complexes that permit DNA lesion
tolerance, specialized polymerases that afford lesion bypass, and a large battery of DNA repair
proteins. Loss of one or more of these networks results in loss of informational integrity and,
ultimately, the onset of disease [2].
Once it was appreciated that DNA lesions cause mutagenic and toxic events, researchers sought
to understand the relationships between the structure of each lesion in DNA and the biological
endpoints indicated above [4].
For example, discovery of the mutagenic lesion of a
carcinogenic DNA damaging agent might lead to strategies to reduce the level of that lesion in
DNA, and hence reduce the likelihood of carcinogenesis.
Studies on the DNA adducts of
aflatoxin Bi led to intervention strategies at the population level that offer promise of reducing
liver cancer burden [5]. As a second example, knowledge of the relationship between the
structures of DNA adducts of anticancer drugs and cytotoxicity endpoints can aid drug
development efforts in clinical pharmacology.
While it is obvious that establishing the
relationships between DNA adducts and their biological endpoints is important, it proved very
difficult to develop an experimental strategy to address the problem. Even a single simple DNA
damaging agent such as the aforementioned aflatoxin results in nearly a dozen DNA adducts,
which frustrated early attempts to determine which adducts are biologically important [6].
Dissection of the relative biological importance of individual DNA lesions proved to be a
tractable problem with the advent of methodology whereby investigators could place one
lesion at a time into synthetic DNA (Figure 2.2). In early in vitro studies, the oligonucleotides
with adducts at known sites were acted upon with purified polymerases (Figure 2.2A) and
repair proteins, which gave results that helped predict the biological relevance of a lesion and
helped define the cellular repair systems that might protect against it. A second step involved
the use of shuttle vectors that were globally modified by a DNA damaging agent (Figure 2.2B).
Chemical or enzymatic tools allowed the mapping of some (but not all) lesion sites along a
stretch of DNA. The damage spectrum was then compared to the spectrum of mutations that
arose when the modified vector was replicated within cells. Often multiple types of mutation
were observed at a single site and it was impossible to ascertain if a single lesion gave rise to
multiple mutations at, for example, a guanine site, or whether there were several distinct
guanine adducts each of which had its own signature and singular mutation. Nevertheless, this
approach was and continues to be a cornerstone of mutation research.
The fusion of chemistry and biology, termed "chemical biology," gave rise to a more advanced
technology in which synthetic oligonucleotides containing well-characterized single DNA lesions
were genetically engineered into the genomes of viruses or plasmids, which could be
introduced into bacterial or mammalian cells (Figure 2.2C). Within the cell, the lesion would
encounter the host repair and replication systems much in the same way that the lesion would
be treated if it had formed endogenously. Lethal endpoints could be measured as a decrease in
viral or plasmid progeny. Mutagenic outcomes could be determined by interrogating the vector
genomes in the vicinity of the genomic site that originally contained the adduct. The relative
importance of various DNA repair and polymerase systems to deal with or process the adduct
could be determined by introduction of the vector into cell strains with known defects in repair
or replication. In time, the quantitative and qualitative features of mutagenesis and toxicity of
a wide array of DNA damaging agents were profiled by this new technology.
This review examines in detail the application of a variety of experimental systems, primarily
the use of site-specifically modified vector genomes, to categorize the mutagenic and toxic
properties of DNA alkylating agents. Such agents are common environmental carcinogens,
some are formed endogenously and cause spontaneous DNA damage, and some have found
use as cancer chemotherapeutic agents. The paper specifically reviews current knowledge of
the biological properties of each of the lesions formed by low molecular weight alkylating
agents. The structures of the relevant lesions are shown in Figure 2.3. By compiling data on
lesion mutagenicity, genotoxicity and repairability, we develop a biological "fingerprint" for
each lesion (Table 1). It is noteworthy that some lesions have mutagenicities at or approaching
100%, whereas others display comparatively weak mutagenic properties; however, it must be
kept in mind that a lesion with a mutagenicity of only 0.1% creates mutations at a rate that is
five orders of magnitude greater than the basal or spontaneous rate of mutagenesis. In the
review, exocyclic mono-adducts are covered first, followed by adducts in which endocyclic
atoms are the points of attachment to the alkyl residue. The final sections of the review cover
small cyclic adducts. To keep the manuscript of a manageable size, we have limited our
attention to adducts of one or two carbon residues, avoiding larger adducts and some of the
lipid-derived adducts that have been reviewed elsewhere [7].
2.3 06-Methylguanine and 0 6-ethylguanine
0 6-Methylguanine (O6MeG), which causes G-->A transitions [8], is the primary mutagenic lesion
under most conditions of alkylation damage to the genome [9]. 0 MeG is formed from both
endogenous [10;11] and exogenous sources [12], and studies have correlated its persistence to
organ-specific tumorigenicity in rats [13]. 0 6-Ethylguanine (O6EtG) is the major mutagenic
lesion formed by ethylating agents [14] and also primarily causes G+A transitions [15].
Escherichia coli has two 06-methylguanine-DNA-methyltransferases that can repair the adduct the constitutive Ogt protein and the inducible Ada protein, which directly reverse methylation
damage by transferring the alkyl group to one of the internal cysteine residues on each repair
protein. This transfer irreversibly inactivates the repair proteins, making the non-enzymatic
stoichiometric reaction "suicidal" [16].
Ada is part of the adaptive response, which was
discovered when E. coli treated with a low dose of a methylating agent acquired resistance to
the mutagenicity and toxicity of subsequent higher doses [17]. The alkyl groups from 0 6AlkGua
and 04AlkThy are transferred to Cys 321 at the C-terminus, while those from a third substrate,
methylphosphotriesters (MePT), are transferred to the N-terminus of Ada.
It was initially
believed that the methyl group from MePT was transferred to Cys-69 on the protein [18] but
recent evidence identifies Cys-38 as the acceptor residue [19]. Methylation of Cys-38 of Ada
converts it to a transcriptional activator of the genes encoding the "adaptive response" to
alkylating agents, namely, ada, alkA, alkB and aidB. This isthe most nucleophilic of all available
cysteine residues in Ada since it is not part of a network of hydrogen bonds. Methylation at this
site reduces the overall negative charge on Ada. Reduction in charge density is important for
the role of Ada as a transcription factor as it enhances its interaction with negatively charged
DNA by 1000-fold [20]. The number of Ada molecules is estimated to rise from 1-2 molecules in
an unadapted state to ~3000 molecules in a fully adapted cell [21;22]. It was initially found that
Ada preferentially repairs 06MeG as compared to 04 -methylthymine (04MeT) [23] but recent
evidence suggests it repairs both lesions with equal efficiency [24].
The second DNA methyltransferase, Ogt, was discovered by deletion of the ada operon [25;26].
Unlike Ada, Ogt is constitutively expressed in E. coli, shows a preference for repair of 04MeT
and larger alkyl adducts, and does not repair MePT [26]. It is estimated that there are ~30
molecules of Ogt in wild type E. coli [22]. The mammalian homolog of Ogt and Ada is MGMT
(also referred to as AGT). This enzyme works in a similar suicidal fashion but is not inducible,
and it shows a 35-fold higher preference for repairing 0 MeG over 04MeT [24]. Human MGMT
can be silenced by epigenetic modifications [27].
This silencing plays a dual role in
carcinogenesis as tumors not expressing MGMT acquire a mutator phenotype but also become
more susceptible to killing by alkylating agents [28].
Ogt is speculated to provide protection at low levels of sporadic exposure to alkylating agents,
whereas the adaptive response becomes more important against higher chronic exposures or
acute exposures that trigger the transcriptional switch of the adaptive response operon.
In
addition to the methyltransferases, the UvrABC nucleotide excision repair (NER) pathway can
also repair 06 MeG. Excision of 06MeG on duplex substrates has been shown to occur in vitro
[29] and in vivo [30]. When O6MeG is present in a single-stranded context in vivo, NER does not
affect mutation frequency of the lesion; the mutation frequencies in E. coli uvrB~ada ogt cells
are very similar to those found in uvrB-adaogt cells [31]. Interestingly, Chambers et al. found
a 40-fold decrease in the G-A transition caused by an O6 MeG lesion introduced on a singlestranded <PX174 genome in an NER deficient (uvrA) cell strain vs. wild type [32]. The authors
suggest a shielding mechanism by which UvrA binds to the lesion and protects it from repair by
Ada or Ogt, leading to elevated mutation frequencies. There is some evidence of the NER
pathway playing a role in repair of 0 MeG in Drosophila melanogaster [33], and of 06 EtG in D.
melanogaster [34] and mammalian cells [35].
The mismatch repair (MMR) pathway has also been implicated in the cellular response to
o
MeG [36]. 06MeG can be processed by post-replicative mismatch repair in E. coli in a double-
stranded context, but in a single-stranded context (a gapped plasmid) the mutation frequencies
in wild type and mutS cells are the same [37]. Using an M13 single-stranded system containing
an 06 MeG lesion, Rye et al. showed that the Dam and MutH proteins did not impact the repair
of O6 MeG by Ada or Ogt, but cells lacking MutS and MutL were less efficient at repairing 06MeG
by Ada or Ogt [38]. The study also suggested that the MutS and MutL MMR proteins could aid
the repair of O6 MeG by identifying and presenting the lesion in a manner that facilitated the
activity of methyltransferases on the lesion. While early work suggested that O6 EtG is not
repaired by alkyltransferases or MMR in E. coli [39], more recent studies suggest that it is
repaired by the same machinery that repairs O6MeG in mammalian cells [40]. Nevertheless, in
rat mammary cells, 0 EtG is repaired 20 times faster than 0 MeG by an unknown, MGMTindependent, mechanism [41].
In line with expectations based upon this finding, a G4A
mutation is not seen as a frequent event at codon 12 of the H-ras gene in tumors initiated by Nethyl-N-nitrosourea (ENU) compared to tumors initiated by N-methyl-N-nitrosourea (MNU).
The toxicity of O6MeG has been established by several studies, and it appears that abortive
MMR or inhibition of replication systems may play roles in converting the adduct into lethal
intermediates. Evidence that O6MeG is potently toxic in mammalian cells come from a number
of studies, including those in MGMT knock-out mice, which display hypersensitivity to the lethal
effects of alkylating agents that generate O6MeG [42]. There are two proposed mechanisms by
which this lesion contributes to the toxicity generated by alkylating agents. The first suggests
that the lesion reduces the efficiency of replication by polymerases. This phenomenon has
been studied using in vitro systems.
The rates of replication by T4 and T5 phage DNA
polymerases and E. coli polymerase I decrease linearly with increasing proportion of O6 MeG in
the synthetic oligonucleotide used as a template [43]. Also, human polymerase P, subcloned in
an E. coli plasmid, is blocked by O6MeG present on a single-stranded DNA template [44]. The
second mechanism leading to toxicity is that of futile cycling of the mismatch repair system at
an 0 MeG:T pair [45;46].
The model proposes recognition of this base pair by the MMR
enzymes, which results in the removal of the newly incorporated thymine from the nascent
strand opposite the lesion. On re-replication, O6MeG preferentially pairs once again with an
incoming thymine [8], reinitiating the repair and replication cycle. This persistent iteration of
excision and synthesis is thought to result in a stabilized nick or small gap in one strand of DNA,
which may activate damage signaling pathways [47].
The recursive cycling mechanism is
thought to be of practical significance in that it may explain the lethal effects of the anticancer
drug, temozolomide [48]. In E. coli, 06EtG is more toxic than 0 MeG [39] but the mechanism
underlying this differential toxicity is unknown.
06MeG is known to be highly mutagenic. To study the mutations formed in vivo, Loechler et al.
constructed single-stranded M13mp8 DNA containing 06 MeG at a specific position and
transfected the same into E. coli. It was found that the predominant mutation generated by
this lesion was a G->A transition. In wild type E. coli, the lesion was weakly mutagenic, but
challenging the Ada and Ogt repair systems of the cell by treatment with N-methyl-N'-nitro-Nnitrosoguanidine (MNNG; which forms alkyl adducts in the host genome) resulted in a robust,
dose-dependent demonstration of the mutagenic power of this adduct [8]. This early study
showed how significant even a few molecules per cell of a DNA repair protein could be as a
protection against DNA damage. The ethyl homolog of O6 MeG, 06EtG, introduced at a specific
position in OX174 and transfected into E. coli produces higher mutation frequencies compared
to 06 MeG in the same system [49;50].
0
MeG and O6 EtG also have been site-specifically
incorporated in Chinese hamster ovary cells and are shown to have a mutation frequency of
19% and 11%, respectively, in cells lacking 0 6 -alkylguanine-DNA alkyltransferase [51].
A recent study used site-specific mutagenesis to generate single-stranded M13mp7 genomes
containing 06 MeG in all sixteen possible permutations and combinations of nearest neighbor
sequence contexts. These genomes were then introduced into E. coli mutants of different
repair backgrounds and the mutation frequencies were determined by a novel and very
sensitive assay. It was found that 06 MeG went from being 10% mutagenic in repair-proficient
cells to 100% mutagenic in repair-deficient cells [31]. Moreover, it was found that DNA repair
in vivo is sequence context-dependent.
With regard to effects on gene expression, 0 MeG can inhibit carbon-5 methylation of
cytosines in CpG
motifs by interfering with the binding of 5-methylcytosine DNA
methyltransferases; eventually this interference with natural methylation can lead to genome
hypomethylation. The pairing of O6MeG with thymine can also lead to DNA hypomethylation
[52]. By these mechanisms, the formation of this adduct could affect the epigenetic program of
mammalian cells.
2.4 0 4-Methylthymine
0 4MeT is one of the mutagenic lesions formed concurrently with
06 MeG when DNA is exposed
to alkylating agents that react with DNA by an SN1 mechanism. 04MeT is formed at a much
lower level than 0 MeG; for example, the methylated thymine was detected at a level 126
times lower than that of O6MeG in calf thymus DNA treated with MNU [53]. Although it is not
an abundant lesion, 0 4 MeT can be very mutagenic. Using site-specific mutagenesis tools, it was
shown that 04MeT incorporated in single-stranded M13mp19 had a mutation frequency of 12%
in repair-proficient E. coli. 0 MeG gave a mutation frequency of less than 2% in the same
repair-proficient system. Pretreatment with MNNG to deplete or occupy endogenous repair
enzymes doubled this mutation frequency [54]. Similar results were obtained using doublestranded and gapped plasmids in E. coli (mutation frequency of 45% for 04MeT vs. 6% for
0 MeG) leading to the conclusion that 04MeT is much more mutagenic than O6MeG [39] on a
mole-per-mole basis under normal conditions of DNA repair proficiency in cells. 04MeT mimics
cytosine in structure and generates an overwhelming majority of T4C transitions [55]; it can
also cause a small number of T-A transversions in MMR deficient cells [40]. 04MeT has been
examined as a site-specific adduct in mammalian vectors and again appears to be more
mutagenic than 06MeG in both repair-proficient and repair-deficient backgrounds [40;56]. In E.
coli, 04MeT is toxic but less so than 06MeG and 06EtG [39]. 04MeT has been shown to be toxic
to mammalian cells deficient in NER capability [57], suggesting a role for this repair pathway in
the cellular defense against this adduct.
In E. coli, 04MeT is repaired by the same alkyltransferases that repair 06MeG. The Ogt protein
from E. coli seems to have a preference for repair of 04 MeT over 06 MeG [26]. Ada repairs
0 MeG and 04MeT with equal efficiency but human and rat alkyltransferases show a
preference for 0 MeG repair [24;58].
Studies in mammalian cells have shown that the
mutation frequency of 04MeT does not vary significantly in the presence or absence of
alkyltransferase, indicating that it is probably not repaired by MGMT [40;56].
In fact,
mammalian alkyltransferases may actually inhibit repair of 04MeT by the NER pathway by
binding to and shielding the lesion, as evidenced in E. coli in studies using plasmids expressing
human and mouse methyltransferases [59].
A study done in human cells lines using site-
specifically modified plasmids containing 04MeT shows that repair is not influenced by the
levels of alkyltransferase and that NER seems to be the most significant repair system for this
lesion [57]. With regard to repair by MMR, one in vitro study found that E. coli MutS (a DNA
mismatch repair binding protein) does not bind to oligonucleotide-duplexes containing a site-
specifically incorporated 04MeT:A base pair [36], while another shows that hMutSa, a protein
of the MMR pathway in humans, recognizes and binds to a 04MeT:A base pair quite well but
very poorly to an 04MeT:G base pair [60].
2.5 0 2-Methylcytosine and 0 2-methylthymine
o
-methylcytosine (O2 MeC) and 0 2-methylthymine (O2MeT) are minor reaction products
formed by treatment of DNA with alkylating agents such as MNU or MNNG. Both lesions are
repaired in vitro by E. coli AlkA [61]. 02MeC and O2MeT are predicted to interfere with minor
groove contacts, yet there have been very few studies of these modifications, making the
lesions good candidates for future study.
2.6 Methylphosphotriesters
Methylation
damage
can
occur on the
methylphosphotriesters (MePT).
DNA
sugar-phosphate
backbone to
form
The physical accessibility and negative charge of the
phosphate oxygens makes them a favorable site for chemical reaction. When double-stranded
DNA is treated with MNU, 17% of the total methylation occurs on the backbone to yield
methylphosphotriesters [14]. These adducts react with water and other nucleophiles much
faster than the common diester form of phosphate linking adjacent nucleosides, leading to
facile cleavage of the backbone.
Of the two diastereomers formed, only the Sp-MePT is
repaired by the Cys-38 residue in the N-terminal domain of Ada (N-Ada). This selective repair
results because the oxygen atom on the phosphate in the Sp diastereomer is only 3.5A away
from the acceptor cysteine residue, vs. 5A in the R, configuration [19]. As discussed earlier, N39
Ada has an inherent electrostatic switch that works in a methylation-dependent fashion to
modulate its affinity for DNA and ability to act as a transcription activator. There is no known
homolog of N-Ada in eukaryotes, thus making the repair of MePT in mammalian cells uncertain.
In vivo studies using wild-type Ada and truncated Ada (lacking MePT repair capability)
transfected into HeLa cells showed the same extent of resistance to the cytotoxic effects of
alkylating agents, similar sister-chromatid exchange induction, as well as host-cell reactivation
of adenovirus [62]. This observation suggests that MePT may not have cytotoxic effects in cells.
The role of MePT seems to be a chemosensor for detection of methylation damage and
induction of the adaptive response in E.coli, but their role, if any, in eukaryotes is unknown.
2.7 N1-Methyladenine and N1-ethyladenine
N1-Methyladenine (1MeA) is formed by alkylating agents mainly in single-stranded DNA and
has been
detected in vitro [63-69] and in vivo [65;70-731.
SN2
agents, such as
methylmethanesulfonate (MMS) and the naturally occurring methyl halides can generate 1MeA
[16]; similarly, the ethyl homolog, N1-ethyladenine (1EtA), is formed by ethylating agents both
in vitro and in vivo [65]. The preference for formation in single-stranded DNA is owed to
location of the N1 atom of adenine at a site usually protected by base pairing in doublestranded DNA [74].
1MeA is cytotoxic because it disturbs DNA replication [16].
Methyldeoxyadenosine is known to be unstable due to a base-catalyzed
1-
Dimroth
rearrangement, a complex mechanism, the net result of which is the migration of the NI
methyl group to the exocyclic N6 position of adenine [75].
A specialized DNA repair system protects cells from Ni-substituted DNA lesions. The AlkB
enzyme of the adaptive response repairs 1MeA both in vitro and in vivo [76] in an oxidative
reaction that liberates formaldehyde from the methylated base, affording complete reversal of
the damage. The role of AIkB in the repair of 1MeA seems to be the prevention of genotoxicity,
because this very toxic adduct is only weakly mutagenic in cells. AIkB and its human homologs,
ABH2 and ABH3, also repair 1EtA residues in DNA, with the release of acetaldehyde as the
repair product [77].
Studies of 1MeA in vivo reveal that the lesion severely blocks DNA
replication, but the replication block can be partially overcome by the induction of SOS bypass
polymerases. The 1MeA blockade is completely removed in AlkB-proficient cells [76],
underscoring the physiological relevance of the AIkB system for countering the toxicity of this
base. While very toxic, as indicated above, 1MeA is at best weakly mutagenic. To the extent
that it is mutagenic, 1MeA induces A to T mutations, which are enhanced following induction of
the SOS polymerases. The base composition for A vs. T was respectively 99% vs. 0.61% in SOS
/AIkB cells, 99.7% vs. 0.06% in SOS/AIkBt cells, and 98.6% vs. 1.0% in SOS*/AlkB cells [76].
While the AlkB protein can repair the 1EtA lesion, it cannot repair 3-ethyladenine damage,
which parallels AIkB's activity on 1MeA but not on 3-methyladenine [77]. AlkB repairs 1EtA
somewhat less well than 1MeA.
2.8 N3-Methyladenine
N3-Methyladenine (3MeA) can be formed in DNA by methylating agents as well as nonenzymatically by intracellular SAM. In a mammalian cell, SAM or some other methylating agent
reacts with DNA to generate an estimated 600 3MeA per day [78]. The half life of 3MeA in vivo
is estimated to be between 4-24 h [79]. While 3MeA is not particularly mutagenic, it is a
cytotoxic DNA lesion by virtue of its ability to block replication or by virtue of its ability to give
rise to a chemically- or enzymatically-generated abasic/apurinic (AP) site.
With regard to
replication inhibition, it is thought that the methyl at the N3-position of purines sterically
interferes with the required contact between the polymerase and minor groove on DNA [80].
This property makes it essential for the cell to have in place defenses against this form of
damage. 3MeA-DNA-glycosylases have evolved in both prokaryotic and eukaryotic systems to
afford the efficient repair this lesion. The prokaryotic system includes the highly selective and
constitutive TAG protein, and the inducible AlkA glycosylase with a broader specificity. The
eukaryotic system is comprised of alkylpurine-DNA-N-glycosylases (APNG) and human 3MeADNA-glycosylase (AAG/ MPG). AlkA and TAG repair 3MeA with equal efficiency on doublestranded DNA, but AlkA is 10-20 fold more efficient on single-stranded DNA [81]. There is also
evidence that UvrA, an ATPase and DNA-binding protein of the NER pathway, may be able to
mitigate the cytotoxic effects of this lesion. One study used a neutral DNA equilibrium binding
agent, Me-lex (N-methylpyrrolecarboxamide dipeptide (lex) modified with an 0-methyl
sulfonate ester functionality), to introduce selectively 3MeA lesions in the minor groove of
DNA. It was shown that this agent shows increasing toxicity to E.coli mutants lacking one base
excision repair (BER) repair enzyme (AlkA), two BER enzymes (AlkA and TAG), or both BER and
NER repair capabilities (AlkA, TAG and UvrA), in that order [82].
3MeA is not considered to be a seriously promutagenic lesion based upon work done in
bacterial and in yeast systems.
In 3MeA-DNA-glycosylase I (tag) deficient E. coli mutants,
treatment with MNU leads to a 5-fold increase in mutation frequency only under SOS-induced
conditions. Furthermore, in repair-proficient cells, removal of 3MeA from the DNA does not
show a significant difference in mutagenesis in SOS-induced vs. SOS-uninduced cells [83]. To
study the mutational profile of 3MeA in eukaryotic cells, the p53 gene cDNA on a yeast
expression vector was treated with Me-lex in vitro and transfected into a yeast strain containing
the p53-dependent reporter ADE2 gene. The results show that Me-lex is a weak mutagen
compared with MNU, but that it induces A-> T transversions as the most common genetic
change (40% of all mutations) [84].
Mutagenicity increased 2-3 fold in 3MeA-glycosylase
deficient strains, which suggests that the lesion driving the mutations is 3MeA [85].
Interestingly, the methylated adenines in Me-lex treated DNA give rise to mutations in a strictly
sequence-specific manner.
The cytotoxicity of 3MeA is well established in the literature. In vitro studies showing chain
termination one nucleotide 3' to adenines in methylated DNA templates pointed to 3MeA as a
strong block to DNA replication. 3MeA in DNA has also been shown to be toxic in E. coli [82].
Using Me-lex in combination with 3-methyladenine-DNA-glycosylase proficient and deficient
cell lines, Engelward et al. showed that 3MeA can cause p53 induction, S phase arrest, sister
chromatid exchange (SCE), chromosome aberrations, and apoptosis in mammalian cells [86].
As with 7MeG, enhanced repair of 3MeA by DNA glycosylases of the BER pathway can lead to a
flood of AP sites that can also contribute to mutations and lethality [87].
2.9 N7-Methyladenine
N7-Methyladenine (7MeA) is a minor lesion formed at a level 40-fold below that of 7MeG,
which is typically the most abundant lesion in alkylated DNA [88]. Like 7MeG, 7MeA possesses
a cationic imidazole ring, which facilitates depurination and, alternatively, can favor hydrolysis
of the five-membered ring to form the formamidopyrimidine derivative, Fapy-7MeA; this latter
hydrolysis reaction is especially favored for the 7MeA in RNA [89], which has a stabilized
glycosidic bond as compared to DNA. The half life of 7MeA in DNA in vivo is only 2-3 hours,
which is similar to its half life in vitro at pH 7.2, 37 0C [90]. Fapy-7MeA is a mutagenic lesion,
displaying A->G transitions in single-stranded M13mp18 DNA transfected into SOS-induced E.
coli [88;91]. In these studies, dimethylsulfate (DMS) treated DNA was compared before and
after treatment with alkali, which hydrolyzed the imidazole rings of N7-methylated adenines
and guanines, forming the Fapy derivatives. DMS and alkali treated DNA was 60-fold more
mutagenic than DNA treated with DMS alone, and showed mutations primarily at A:T sites.
2.10 N1-Methylguanine
N1-Methylguanine (1MeG) has been found both in vitro [68] and in vivo [92]. With regard to
biological relevance, the AIkB protein can repair 1MeG both in vitro and in vivo [76;93]. The
glycosylase AAG, which repairs 3MeA and a range of other lesions, is also active against 1MeG
in vitro [94] but the in vivo relevance of AAG against this adduct has not been established as
yet. 1MeG is a very strong block to replication, which can be partially overcome when the DNA
lesion is partially repaired by AIkB; lesion bypass of 1MeG in vivo increases 8-fold from 2% in
AIkB cells to 16% in AlkB* cells. Similarly, AlkB causes a reduction in the mutagenicity of 1MeG
44
from a very high frequency of 80% in AIkB cells to 4% in AlkB* cells. Taken together these data
indicate that AIkB is a powerful protection against the mutagenic activity of this dangerous
alkylated base. The mutational fingerprint of 1MeG reveals G -+T (57% of all progeny), G 4A
(17%) and G 4C (6%) mutations.
In many instances, the induction of the SOS bypass
polymerases results in increased bypass of a given lesion at the expense of reduced fidelity at
the site of damage; however, the SOS polymerases are somewhat anti-mutagenic when they
bypass this modified base [76].
2.11 N3-Methylguanine
N3-Methylguanine (3MeG) is thought to block replication in the same way 3MeA does, but it is
formed in DNA at a 15-fold lower level. The half-life of 3MeG in vivo has been shown to be 3 to
4 hours [90]. It has been shown that E.coli alkA mutants are sensitive to alkylating agents even
though they express Tag [95], which repairs 3MeA (a known cytotoxic lesion) as efficiently as
AlkA on double-stranded DNA [81]. This result suggests that 3MeG contributes to the toxic
effects of alkylation seen in these cells.
Using cell extracts from adapted E. co/i, it was shown that the AlkA protein can repair 3MeG
present on methylated DNA in vitro. The same study also shows persistence of this adduct in
unadapted E. coli 30 min after exposure to MNNG [96]. A second in vitro study has shown that
TAG also repairs 3MeG present on a synthetic GC rich double-stranded DNA sequence, albeit
with an efficiency only 1 / 7 0 th that of AlkA [97].
2.12 N7-Methylguanine and its degradation products
The N7 atom of guanine is the most chemically vulnerable site to attack by alkylating
electrophiles as it has the highest negative electrostatic potential of all the other atoms within
the DNA bases [98]. This property also makes it a highly reactive ligand for metal ions such as
platinum [99].
When double-stranded DNA is treated with MMS or MNNG, 82% and 67% of
the methylation occurs on the N7-position of guanine, respectively [14]. Within the cell, N7methylguanine (7MeG) is produced at the rate of 4000 residues/human genome/day by the
non-enzymatic reaction of SAM with DNA [781, and its steady-state level in repair-proficient
cells is estimated to be 3000 bases [100]. 7MeG has been detected in human DNA at the level
of a few adducts per 10 7 bases [101]. 7MeG by itself does not have any major mutagenic or
cytotoxic effects. However, methylation at the N7-position destabilizes the N-glycosidic bond
leading to spontaneous depurination of this lesion [102] and the resulting AP sites are toxic. AP
sites can also be formed during repair of 7MeG by N-alkylpurine DNA glycosylases which are
part of the BER pathway.
Although not examined directly in the context of alkylation, the
mutagenic and toxic properties of AP sites have been thoroughly investigated [87].
In addition to its role as a source of AP-sites, 7MeG can manifest toxicity by converting to its
imidazole ring-opened form. Hydrolysis of the imidazole ring of 7MeG forms 2,6-diamino-4hydroxy-5N-methyl-formamidopyrimidine (Fapy-7MeG).
While this lesion does not cause
misparing with dAMP or dTMP, in vitro experiments using E. coli DNA polymerase I and poly
(dGC) templates [103], or Klenow fragment and M13mp8 template DNA [104] show that Fapy7MeG blocks DNA chain elongation. Fapy-7MeG lesions present on M13mp8 phage template
DNA also leads to a 2-3 fold increase in G-C and G4T transversions when transfected into SOS
induced E. coli [88]. However, DNA polymerase I preferentially incorporates dCMP opposite
Fapy-7MeG and a Fapy-7MeG: C pair is extended most efficiently compared to other
possibilities. This property makes Fapy-7MeG a lesion with weak mutagenic potential [89].
In E. coli, AlkA is known to excise 7MeG from methylated DNA [96]. In humans this reaction is
carried out by AAG/MPG
[105].
There exist specific DNA glycosylases in E. coli
(formamidopyrimidine-DNA-glycosylase (Fpg)) and mammalian cells (human 7,8-dihydro-8oxoguanine DNA glycosylase (hOGG1)) [106] that remove Fapy-7MeG lesions.
E. coli Fpg
repairs 7MeG very efficiently, with a Km in the nanomolar range [89]. It has been shown in a
mammalian cell line by site-directed mutagenesis that overexpression of MPG sensitizes cells to
alkylation damage by converting 7MeG into toxic AP sites, which lead to strand breaks. 7MeG
by itself is not toxic to cells, nor is overexpression of MPG, but in combination they can
overwhelm the cell with AP sites leading to cytotoxicity. Rinne et al. propose that these two
aspects combined with appropriate delivery systems could be exploited for the selective
targeting of tumor cells, thereby reducing the peripheral effects of DNA damage by drugs [107].
2.13 N3-Methylcytosine and N3-ethylcytosine
N3-Methylcytosine (3MeC) is formed by SN2 agents such as MMS and the naturally occurring
methyl halides [16] preferentially in single-stranded DNA. It has been detected both in vitro
[63;65-69;108;109] and in vivo [65;71;72;92;109].
The corresponding ethyl homolog, N3-
ethylcytosine (3EtC), is formed by ethylating agents in single-stranded DNA and also has been
47
detected in vitro [65;66] and in vivo [65;110]. As with the 1-alkyladenines, these lesions likely
exist only or predominantly in single-stranded DNA because this site of modification is normally
protected by base pairing [74]. 3MeC stalls DNA synthesis and is likely to be toxic [16].
In E. coli, the AIkB protein has good activity against 3MeC and 3EtC both in vitro and in vivo
[63;64;76].
The appreciable mutagenesis and toxicity of the 3-alkylcytosines in vivo is
decimated by AIkB, although a portion of the toxicity can also be overcome by induction of the
SOS bypass polymerases.
With regard to mutagenic potential, if a cell has no AIkB and
uninduced SOS bypass polymerases, 3MeC and 3EtC are 30% mutagenic, with the predominant
mutations being C
-
T and C -
A. Basal expression of AlkB of a few molecules per cell
abrogates the mutagenicity of 3MeC and 3EtC, whereas expression of SOS bypass polymerases
in the absence of AkB increases the mutagenicity of both lesions to a striking 70%. Although
investigations involving replication past 1MeA and N1-methylguanine (1MeG), which similarly
have a blocked Watson-Crick hydrogen bonding face, by DNA polymerases in vitro are lacking, it
is known from in vitro studies that 3MeC inhibits replication by DNA polymerase I and does not
cause mutation [108;111;112]. However, some adduct bypass occurs with the incorporation of
dAMP and dTMP opposite 3MeC [108].
Therefore, the rules for misreplication of the 3-
alkylcytosine lesions are the same both in vitro and in vivo, although the replicative system in
cells is capable of a much higher mutation rate than is achieved in vitro [76;108].
2.14 N3-Methylthymine
N3-Methylthymine (3MeT) has been found both in vitro [65;66;68;69;113] and in vivo [113;114]
and is formed through the reaction of DNA with SN2 alkylating agents such as MMS. This
adduct is a very weak substrate for AlkB, and it is a strong block to replication in vivo, which can
be only slightly overcome by SOS bypass polymerase induction [76]. Recently, FTO (fat mass
and obesity associated) protein has been shown as a 2-oxoglutarate-dependent demethylase
for nucleic acid [115;116]. FTO can efficiently repair 3MeT in single-stranded DNA but not in
double-stranded DNA; it also shows strong activity on the demethylation of 3-methyluracil in
single-stranded RNA [115;117]. While there are numerous epidemiological studies associating
the FTO gene with obesity, the biological basis for metabolic effects of this gene are still under
investigation.
From the standpoint of its potential to induce genetic change, 3MeT is approximately 60%
mutagenic in SOS/AlkB cells, providing mostly T
--
A (47%) and T 4
C (9%) mutations.
Studies performed in vitro also show that 3MeT is a strong block to the Klenow fragment of
DNA polymerase I, which slightly increases dTTP incorporation on a poly(dC-d3MeT) template
[118]; interestingly, T is exclusively incorporated opposite the analogous 3-ethyldeoxythymidine
adduct in one study [119], while A is exclusively incorporated in another [120].
2.15 8-Methylguanine
C8-Alkylated DNA bases exist but have not been reported extensively in the literature. Recent
studies have suggested that carbon-centered radicals can be a source of C8-alkylated lesions.
49
8-Methylguanine (8MeG) was shown to be produced in vitro in RNA [121] and DNA [122] by
methyl radicals generated by oxidation of 1,2-dimethylhydrazine and methyihydrazine
respectively. Proof of in vivo DNA alkylation by carbon-centered radicals was given by Netto et
al. who detected 8MeG in DNA isolated from the liver and colon of rats administered 1,2dimethylhydrazine [123]. Other studies have shown that this lesion can also be produced in
vitro and in vivo by genotoxic agents such as tert-butylhydroperoxide, diazoquinones and
arenediazonium ions [124]. These findings are significant as they suggest a possible
contribution of 8MeG in the carcinogenic effects of these agents, especially 1, 2dimethylhydrazine, which induces adenocarcinomas of the colon in rodents.
Site-specific studies using 8MedG-containing-oligonucleotides prepared by phosphoramidite
synthesis have explored the mutagenicity and toxicity of this lesion. It was found that 8MeG on
the template strand blocks in vitro extension of DNA by mammalian polymerase a, but not by
the E. coli Klenow fragment [125]. The products from the primer extension reaction were then
analyzed for mutations. 8MeG was found to direct exo- Klenow fragment-based incorporation
of dCMP most of the time (77%), but also paired occasionally with dGMP (1.1%) and dAMP
(0.41%). Similar numbers were obtained for extension assays with mammalian polymerase M.
Replication with the Klenow fragment also introduced small amounts of one- (0.38%) and two(0.81%) base-pair deletions [125]. These numbers mirror the thermodynamic stability of the
8MeG:dNMP base pair, decreasing in the order dCMP > dGMP > dAMP >> dTMP.
1,2-
Dimethylhydrazine induces both 0 MeG and 8MeG in similar amounts in the DNA of rats [123].
However, the mutation frequencies of 8MeG are two orders of magnitude less than those of
06 MeG
[126]. Therefore, we may conclude that 8MeG is a weakly mutagenic lesion that in
principle can contribute to G -C transversions in cells.
The repair of 8MeG has been studied in vitro by Gasparutto et al. In this study, the authors
incorporated 8MeG site-specifically into oligonucleotides and probed the ability of bacterial,
yeast and mammalian glycosylases to repair this lesion. Of the extensive list of enzymes
evaluated, only AlkA was able to excise 8MeG. Human MPG did not repair 8MeG, nor did any
of the glycosylases involved in repair of oxidative damage (Fpg, Nth of E. coli; Ntgl, Ntg2, Ogg1
of Saccharomyces cerevisiae; and human Ogg1) [124].
8MeG has been shown to stabilize the Z-conformation of DNA in short oligonucleotides even in
low salt concentrations. This property may be relevant in vivo as Z-DNA is thought to have a
role in the regulation of DNA supercoiling [127]. This lesion is also used as a chemical
modification to stabilize quadruplex structures of G-rich sequences of DNA, which are proposed
to have a role in telomeric DNA stability and in repression of transcription at the c-myc
promoter [128]. The wide range of potential biological activities of this lesion makes it a prime
target for future investigations.
2.16 1,N -Ethenoadenine and 1,N -ethanoadenine
The formation of 1,N6 -ethenoadenine (eA) results from the reaction of adenine with products
of unsaturated lipid peroxidation [129-132]. This bifunctional DNA lesion arises endogenously
under normal physiological conditions in both rodents and humans [133;134].
Of great
toxicological concern is the observation that eA is induced by common industrial agent vinyl
chloride and its metabolites, such as chloroacetaldehyde. eA also occurs in chronically inflamed
human and rodent tissues [135]. Oxidative stress associated with inflammation is increasingly
being linked to neurological disease, cancer promotion and accelerated aging [136].
In duplex DNA, eA can be repaired in vitro by glycosylases of the BER pathway [94;137].
Mammalian cells can also repair etheno lesions by this route in vivo [138;139]. Indeed, the BER
enzyme AAG and its homologs are likely to be the primary vehicles of repair of eA in the duplex
genomes of eukaryotes. By contrast, the in vivo repair of etheno adducts in E. coli was not
clearly understood until recently; for example, one early study showed that neither BER nor
NER figures prominently in etheno lesion repair [140].
Early genetic studies on the
mutagenicity of eA in E. coli reinforced this conundrum [141]. The eA adduct was neither toxic
nor mutagenic despite the fact that the base lacks any structural possibility of Watson-Crick
complementarity. The issues raised in these studies were resolved in 2005 when biochemical
studies provided the possibility that the direct-reversal enzyme, AIkB, may play a significant role
in the defense of cells against this type of bifunctional DNA damage. These biochemical studies
showed that AIkB and its human homolog ABH3 can efficiently repair eA in vitro [142;143].
AlkB uses a unique iron-mediated biochemical reaction involving 0-ketoglutarate as a cofactor
to putatively epoxidize the exocyclic double bond of eA. An epoxide may be hydrolyzed to a
glycol with the glycol moiety being liberated as the dialdehyde, glyoxal. The direct reversal
mechanism is also likely to be operative in vivo, as evidenced by genetic studies in which a
single-stranded vector containing a single eA was replicated in AlkB proficient and deficient E.
coli cells. In AlkB-deficient cells, eA is 35% mutagenic, yielding 25% A ->T, 5%A ->G and 5% A
4C mutations.
SOS induction causes an increased incorporation of deoxyadenosine
monophosphate opposite to eA [142].
1,N6-Ethanoadenine (EA) is the chemically reduced form of eA and forms through the reaction
of adenine with the antitumor drug bis-chloroethylnitrosourea (BCNU).
EA can be weakly
repaired by the E. coli enzyme AlkA [1441 and the corresponding human enzyme AAG [94;145],
which suggested that BER is a means of repair of this adduct. Recent work, however, by Frick
et. al. show that the direct-reversal repair enzyme AIkB easily alleviates the toxicity of EA in E.
coli in vivo [146]. In an AIkB-proficient cell, EA is almost nontoxic (i.e., easily bypassed) and not
significantly mutagenic. However, in AlkB-deficient cells, EA is extremely toxic, showing an 86%
reduction in replication. The adduct is weakly mutagenic causing A 4C (2%), A 4G (1%), and A
4T (1%) mutations [146].
2.17 1,N2-Ethenoguanine and 3,N2-ethenoguanine
1,N2-Ethenoguanine (1,2-eG) and its isomer 3,N2-ethenoguanine (2,3-eG) are cyclic DNA
adducts formed, as with eA, by reagents such as chloroacetylaldehyde [147] or 4-hydroperoxy2-nonenal (HPNE) [148]. Significantly, the former has been found in the liver DNA of rodents
exposed to vinyl chloride [147]. 1,2-eG can moderately block DNA polymerase and cause G-T
and G-*C base substitutions, as well as frameshift mutations [149].
mammalian uracil-DNA-glycosylase (MUG) and AAG [94;150].
It can be repaired by
In recent work, 1,2-eG, was
shown to be repaired, albeit weakly, by BER, using a truncated form of the AAG enzyme [94].
The AlkA protein can release 2,3-eG from DNA [151]. The glycosidic bond of 2,3-eG is extremely
labile, a property that has made assessment of biological significance of this modified base a
difficult task [147]. Nevertheless, Loeb and colleagues successfully determined the mutation
frequency of the lesion to be approximately 13% in E. coli, where it primarily induces G 4A
transitions.
2.18 3,N4-Ethenocytosine
3,N 4-Ethenocytosine (eC) is produced from the same precursors and by the same pathways
that generate eA in DNA [129-131;152]. As with eA, the BER pathway (human thymine-DNAglycosylase (hTDG) in human and double-stranded uracil-DNA-glycosylase (dsUDG) in E. coli) is
an established strategy used by nature to suppress the biological effects of this adduct
[138;152]. The cellular defense network against eC additionally involves the AlkB pathway, at
least in E. coli., which should be mechanistically similar to that of eA repair by AlkB [142]. In E.
coli, AlkB has a modest effect on eC toxicity, but reduces the mutation rate of the adduct by
about two-thirds from 82% in AlkB-deficient cells to 37% in AlkB-proficient hosts, implying
incomplete conversion to cytosine prior to polymerase traversal. The mutations of eC in AlkBdeficient and -proficient cells are C->A and C 4T, which are of approximately equal abundance
in each cellular background.
2.19 Perspective
Thirty years ago, when Carcinogenesis was a new Journal, the field of cancer research looked
very different from the way it looks today. The field was richly populated by chemists who
identified carcinogens and studied the molecular transformations whereby those agents
damaged DNA. The work described in this review started shortly after the Journal began, when
the complexities of DNA adduction confounded attempts to relate specific types of DNA
damage with genetic changes that, presumably, attend the conversion of a normal cell into a
fully malignant one. From that time to the present, much has been learned. Many oncogenes
and tumor suppressor genes have been discovered and placed like footsteps on the path
between normalcy and malignancy [1531. More recently, linkages have been made between
the genetic events of oncogene activation and tumor suppressor gene inactivation and parallel
disruptions in biochemical networks. These studies are revealing the secrets of how cancer
cells obtain the energy and the raw materials to finance their growth into a tumor [154;155].
One revelation to come out of the last few decades isthat the number of mutations in cancers
is far in excess of the number one would expect on the basis of normal replication errors, or
perhaps even the enhanced rate of replication errors that occurs when a polymerase tries to
copy past a mutagenic DNA lesion such as those described in this manuscript. While it seems
likely that genetic changes induced by carcinogens are an important step in the early stage of
malignant transformation, it now seems clear that we need to find other chemical or
biochemical events that underpin the "mutator phenotype" of tumors [156]. Answers may
come from studies of virally induced diseases, such as HIV and hepatitis, where recent work has
discovered enzymatic DNA-targeted base deamination systems that cause a high density of
mutations within a genome [157]. Answers might also come from the field of immunology
where enzymes such as AID cause, once again, a high density of mutations in a localized stretch
of DNA [158;159].
One of the most important contributions of work on the chemical biology of mutagenesis has
been the collateral impact of this field on the nearby field of DNA repair. It is now common for
workers in the repair field to use oligonucleotides with single lesions, originally made for
studies of mutagenesis, to characterize the detailed biochemical mechanisms by which repair
enzymes or complexes reverse the damage. Moreover, studies of mutagenesis done using cells
that are defective in a specific repair enzyme [142] or that express specialized polymerases
[160] have provided high quality data that have established the physiological relevance of
specific enzymes as protectors from damage, or as the vehicles by which damage is processed
into events with disastrous consequences for the cell and organism.
Looking ahead, there is much to do. To give one example, the process of inflammation is clearly
associated with cancer development [136].
The range of DNA damages created by
inflammation-generated reactive oxygen and nitrogen species is vast, and the task will be a
large one to determine how each of these lesions contributes to the biological endpoints
downstream of an inflammatory event.
As a second example, workers will soon develop
modified versions of the tools described herein to probe what may become a new field ... RNA
repair. Some mRNA species are so long that it takes a day to transcribe them [161-163]. These
important molecules not only need to have their informational integrity protected, but the
energy used in their synthesis is large and would be wasted if a single lesion, for example an eA
residue, made them un-readable. Finally, while this review focuses on only one class of lesion,
the small alkylated bases, it illustrates how much can be learned about the chemical rules of
mutagenesis. Ten years ago, studies of the mutagenic properties of 5-hydroxycytosine [164],
which induces C to T transitions, were the starting point for a novel application in the
development of antiviral agents [165]. It is expected that additional examples of this nature, in
which basic studies of mutagenesis drive clinical development, will help propel this field into a
robust future.
2.20 Acknowledgments
We thank James C.Delaney and Bogdan Fedeles for editorial and artistic contributions.
2.21 Figures
Figure 2.1: Pathways by which DNA damaging agents induce biologically relevant events.
Agents from the environment, chemically reactive natural species generated within cells, and
the misdirected action of natural intracellular enzymatic systems can result in the formation of
a collection of DNA lesions (symbols attached to the helix). These lesions can be formal
chemical-DNA adducts, such as 06MeG, which is a miscoding lesion during replication. They can
be modifications of the sugar-phosphate backbone, such as MePT, which triggers a change in
gene expression. They can be bases such as uracil, which can appear as the enzymatic
deamination product of cytosine. Or, they can be lethal strand breaks, as would form after
treatment of a cell with ionizing radiation or certain anticancer agents. Finding the
relationships between the structures of each lesion and the biological endpoints of mutation,
lethality and gene expression isthe subject of this review.
DNA Lesions
. Environmental Chemicals
* Endogenously-generated
Chemicals
* Radiation
* DNA-modifying Enzymes
* DNA-damaging Drugs
H3C-O
NH
MeG
\B
o
0
o' MePT
H3C-O-
=O
+-- DNA Repair
0
Uracil
[NH
Replication
B
0
0
~T1
Mutation
Lethality
Altered
Gene
Expression
o YStrand
Break
HO
B
0
HO-A 0
0
0\
Figure 2.2: A. Methods to evaluate the biological relevance of DNA damage. The ability of a DNA lesion
(lollipop structure) to block polymerases in vitro and cause mispairing during DNA synthesis can be evaluated in a
system in which a template containing the lesion is primed with a complementary oligonucleotide that terminates
to the 3' side of the lesion. DNA synthesis may result in incorporation of non-complementary bases or in truncated
products, which can be evaluated on sequencing gels. The same in vitro constructs, in double-stranded or singlestranded form, can also be used as substrates for DNA repair reactions, using purified DNA repair proteins or
cellular extracts. B. To determine the mutagenic properties of the full population of adducts that forms from
treatment of DNA with a mutagen, a plasmid or viral vector istreated with the damaging agent. Replication of the
vector in cells results in repair of some adducts but those that evade repair can possibly be converted into
mutations. Sequencing the genomes of progeny can generate the mutational spectrum, which indicates the types
and frequencies of specific mutations along the DNA sequence being studied. In parallel, one can map the
locations of some of the DNA adducts by using enzymatic or chemical probes. The corresponding damage
spectrum is often compared with the mutational spectrum in order to formulate hypotheses with regard to which
DNA adduct might have caused specific mutations. C. The most sophisticated system for analysis of mutagenesis
involves chemical or enzymatic synthesis of an oligonucleotide that contains a candidate for mutagenesis (often
the candidate is nominated based on the data from experiments shown in part B). The oligonucleotide is inserted
into the genome of a virus or plasmid, which is later replicated within cells, either intra- or extra-chromosomally.
Progeny are analyzed to determine the type, amount and genetic requirements for mutagenesis by the lesion. In
parallel, the reduction in viable progeny is determined as an estimate of the extent to which each lesion inhibits
replication of the genome.
A
Invitro
Replication
,-
Mutation
DNA Synthesis
Inhibition
B
0
Determine
Globally
Damaged
Genome
1 DNA Damage
Spectrum
LAc
Determine
Cell
o
A Mutational
Spectrum
--
r-
7
\yhi
Z
Mutational
E
Damage
, Spectum
CTGCAGGGTACGATTCAGTT
DNA Sequence
Single DNA Lesion
nucleotide
Gapped
Genome
Spectrum
Mutation
-Cell
Site-specifically Modified Genome
Replication
Inhibition
Figure 2.3: Structures of DNA alkylation lesions.
H3C-O
N
H3C
N
</I
O
NH2
N
N
DNA
N
N
-CH 3
NN
DN
N
H3C\
N
NH
O
N
DNA
NH2 H3C
N
N
N
DNA
H3
N
DNA
0 4 MeT
02MeT
NH2
N
0
DNA
DNA
NH2
0
0
N
NH
NH N
DNA
3EtC
3MeC
8MeG
3MeT
N
0
0
N
NN
eA
N
DNA
N
EA
N
DNA
1,2-eG
N
H
N
DNA
DNA
N
N
DNA
NH
3-//N3
NH2
7MeG
3MeG
NH 2
1MeG
7MeA
ON
NI N:5 NH 2
DNA 6H
3
NCH3
N
N
N
I
DNA
N
3MeA
1EtA
1MeA
0
0
N II N
DNA 6H 3
NN
DNA
DNA
MePT
N3C H3 C
DNA
CH3
N
dR
C'
NH2
NH2
dR
0
N
N
&II
O 2MeC
0 6 EtG
0 6MeG
OCH3
H 3 C.
NH2
N
N
DNA
NH2
N
N
H3C'O0
0
N
DNA \==/
2,3-eG
O
DNA
NH2
2.22 Tables
Table 2.1: Mutagenicity, genotoxicity, and repairability of DNA alkylation lesions.
Lesion
Mutagenic
specificity
Genotoxicity
o MeG
G 4A
Toxic in presence of
Ada, Ogt
MGMT
MMR
UvrABC (NER)
NER
MMR
MMR
Not repaired by
Ada or Ogt
MGMT
NER
o EtG
G 4A
Toxic in E.coli
Repaired by
prokaryotic
Repaired by
eukaryotic
enzymes/systems
enzymes/systems
MMR
2
0 MeC
Possibly toxic
0 2MeT
04MeT
MePT
1MeA
Possibly toxic
T -C
T 4A in MMR-
Toxic, but less than
Ada, Ogt
0 MeG and 06EtG in
deficient cells
E.coli
None known
A 4T
None known
Mutagenic and toxic
to E.coli in the
absence of AlkB
Ada
AlkB
Mutagenic and toxic
to E.coliin the
AlkB
ABH2, ABH3
Highly toxic
AlkA
Tag
UvrA
AAG/
APNG/
MPG
G 4T
G 4A
Mutagenic and toxic
to E.coli in the
AlkB
AAG
G 4C
absence of AlkB
1EtA
NER
MGMT (minimal)
AAG
ABH2, ABH3
absence of AlkB
3MeA
A 4T
7MeA
Fapy-7MeA
A
1MeG
4G
3MeG
Possibly toxic
AlkA
Tag
7MeG
3MeC
Fapy-7MeG
G 4C
Toxic via formation
of AP sites and Fapy-
G 4T
7MeG
C4T
Toxic
AlkA
Fpg
hOGG1
AlkB
C 4A
3EtC
4
AAG
ABH2, ABH3
C4*T
C
AAG/
MPG
A
Toxic
AlkB
3MeT
AlkB deficient cell
T 4A
T
G 4C
eA
AlkB deficient cell
A 4T
A 4G
A4C
A 4T (weak)
A 4C (weak)
2,3-eG
eC
FTO
AlkA
Not repaired
by known
enzymes
AAG
Mutagenic and toxic
to E.coliin the
absence of AlkB
AlkA
AlkB
Toxic to E.col in the
absence of AlkB
AlkB
Weak substrate
G 4T
G 4C
G -- A
C 4A
C 4T
AAG
for AlkA
A 4G (weak)
1,2-eG
Weak substrate
for AlkB
4C
8MeG
EA
Strong block to
replication
Mutagenic and
causes frameshift
Mutagenic
Mutagenic and toxic
to E.co/i in the
absence of AlkB
MUG
AAG
AlkA
AlkB
dsUDG
hTDG
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Chapter 3:
DinB relieves the toxicity of N2-dG lesions in vivo.
3.1 Introduction
In Escherichia coli, the dinB gene encodes the Y family DNA polymerase DNA pol IV (or DinB) [1],
which is one of the three translesion synthesis (TLS) polymerases that are part of the SOS
pathway[2]. dinB was first identified as one of the damage inducible genes in E. coli [3] and its
product is known to be involved in untargeted mutagenesis [4-6].
The mammalian DinB
homolog is pol ic.
While it is not essential for life, DinB is unique in that it is the only Y-family DNA polymerase
that is conserved across all domains of life (bacteria, eukaryotes, and archaea) [7], a result of
selective constraints imposed on the encoding gene [8]. It is also present at a high intracellular
concentration, more than that of DNA pol Ill and on par with that of the p-processivity clamp
[9]. In a wild-type E. coli cell, it is estimated that there are 30-50 molecules of pol 11,10-30
molecules of pol 111,250 of pol IV, and 15 of pol V [10;11]. Upon SOS induction, this number
jumps to 2500 for pol IV but only to 200 for pol V [11;12]. Fuchs et al. suggested that DinB may
have a role in bypassing lesions that block replicative polymerases; however, this function is not
unique to DinB and can also be performed by pol 11and pol V [11;13]. DinB is implicated in both
the insertion and extension steps in lesion bypass [14]. It is also known to bind the p-clamp
simultaneously with pol 1I1[15]. The current model is that DinB replaces pol IlIl at stalled
replication forks with high affinity; this swap is made possible by the simultaneous attachment
to the p-clamp. However, at high levels of expression, DinB can interfere with replication fork
progression, leading to cytotoxicity [16]. These properties make DinB a particularly interesting
subject of study.
While it is clear that DinB is an inducible bypass polymerase, the specific substrates for which it
was evolved are, at best, a matter of speculation. In vitro, E. coli pol IV can perform DNA
synthesis across a variety of base modifications, but this translates to a similar result in vivo
only for those induced by benzo[17]pyrene
(BaP),
4-nitroquinoline-1-oxide (4-NQO),
nitrofurazone (NFZ), and reactive oxygen species (ROS). Napolitano et al. showed that DinB is
involved in TLS across a single (+) trans-anti-BaP-N2 dG adduct in vivo [13], the major adduct
formed by the reaction of BaP with DNA [18]. It is also hypothesized that DinB is required for
error-free bypass across this adduct [19] by possibly being the polymerase that inserts the
correct base-pairing nucleotide [20]. AdinB E. coli cells are sensitive to the cytotoxic effects of
4-NQO and NFZ, both of which are known to create lesions on the N2 group of 2'deoxyguanosine (N2 -dG) [21]. DinB may also play a role in the mutagenicity of oxidized DNA
bases. In vitro studies indicate that it is capable of incorporating oxidized DNA bases from the
pool in a fashion that would result in mutation [22]. This activity is also seen in the case of 8oxo-7,8-dihydro-2'-deoxyguanosine 5'-triphosphate (8-oxo-dGTP), where DinB incorporates this
oxidized dNTP from the intracellular nucleotide pool opposite a template deoxyadenine [23].
Furthermore, there is evidence to suggest that DinB may have a role in alleviating the
cytotoxicity of alkylated DNA bases. Bjedov et al. have demonstrated that DinB is essential for
the survival of AalkA Atag cells when exposed to methyl methanesulfonate (MMS), a known
alkylating agent [10]. More recently, Jarosz et al. showed that in vitro, DinB bypassed the N2_
furfuryl-dG lesion, a homolog of the major adduct formed by the reaction of NFZ with DNA in
vivo [21].
Interestingly, DinB was ~15-fold more proficient at inserting a cytosine, and had a
~25-fold higher proficiency in extending beyond a cytosine inserted opposite the lesion over
opposite an unmodified guanine, mostly due to an increased affinity for dCTP [14;21]. DinB has
also
been
shown
to
bypass,
with
efficiency and
accuracy,
N2 -(1-carboxyethyl)-2'-
deoxyguanosine (N2-CEdG) , the major adduct formed by methylglyoxal [24]. This function of
DinB is of particular importance as cells are exposed to alkylating agents through both
endogenous as well as exogenous sources, including chemotherapeutic agents [25].
There are several in vitro studies that explore the function of the mammalian homolog of DinB.
Pol
K
is capable of catalyzing accurate bypass of N2-furfuryl-dG adducts in vitro [21]. It has also
been implicated in bypass across N 2-N -guanine interstrand crosslinks [26]. Human pol K may
have a role in bypass of thymine dimers as evidenced by its ability to extend a primer bearing a
guanine opposite the 3'-dT of a model thymine cyclobutane dimer in vitro [27]. While pol
K performs
proficient insertion of A opposite 8-oxo-dG followed by extension [28], it can
perform error-free bypass of thymine glycol in vitro [29]. Finally, human pol
K
is capable of
incorporating oxidized nucleotides from the pool into template DNA just like its bacterial
homolog. The in vivo properties of pol K are unknown.
Most of the aforementioned studies utilize reagents that generate N2-dG lesions. This work
aims to answer the question whether DinB bypasses N2 -dG lesions in living cells. This thesis
explores four N2-dG lesions as possible substrates for repair by DinB in vivo: 2-methylguanine
(m2G), 2-ethylguanine (e2G), 2-tetrahydrofuran-2-yl-methylguanine (THF) and 2-furan-2-ylmethylguanine (FF or N2-furfuryl-dG). The structures are shown in Figure 3.1. The spectrum of
lesions chosen for this analysis is motivated by several research studies. The first study by
Jarosz et al., discussed above, showed both DinB and pol K being capable of bypass across the
N2-furfuryl-dG lesion in vitro [21]; however it remains unknown whether the same is true in
vivo. N2 -Furfuryl-dG is believed to be the structural analog of the major DNA adduct formed by
NFZ [30], a topical antibacterial agent that was used for treating burns and skin grafts in
patients and animals [31;32]. NFZ reduction metabolites have been shown to be mutagenic
and carcinogenic in rodent models [31;33] and it also causes free radical damage, strand
breaks, and N2-dG adducts in DNA [34-36]. Our study tests whether DinB bypasses the N2furfuryl-dG lesion, and its saturated homolog (THF), in vivo.
The second study by Lu et al. is the first of its kind that measures the endogenous levels of
m2G. This lesion is formed by formaldehyde, a classified human and rodent carcinogen that is
introduced in cells by both endogenous as well as exogenous sources [37]. Lu et al. estimate
that m2G is formed endogenously at the level of ~0.5-0.8 adducts/107 dG. They also estimate
that it is a minor adduct induced by exogenous alkylating agents at a level 100 and 1000 times
lower than that of 06-methylguanine and N7-methylguanine respectively. m2G is a known
modification that occurs on tRNAs across species [38], which may have an effect on the coding
properties and codon-anticodon interactions.
It has also been observed that the tRNA
complement of hepatomas partially lack this modification [39] . However, the biological
significance of m2G in DNA is not yet known. One study explored the miscoding potential of
m2G in an in vitro system with the exonuclease-free Klenow fragment of pol I and found that
the lesion pairs preferentially with cytosine with a 9 % propensity to cause G4A transitions
[40]. Other in vitro studies with human polymerases show low levels of mutagenesis and
toxicity caused by m2G [41;42], except in the case of bacteriophage T7 polymerase (exo-) and
HIV reverse transcriptase [43]. Previous work done in our laboratory has found that m2G is not
mutagenic but is toxic in vivo in all combinations of AIkB and AidB proficiency and deficiency
[44]. However, no studies have explored the effect of DinB on m2G. With the levels now
detected in DNA, it is important to explore the possible repair mechanisms available to process
this lesion.
The third research area that ties into our study is that of e2G, which is formed by the reaction
of deoxyguanine monophosphate with acetaldehyde from both endogenous (metabolic
oxidation of ethanol) as well as environmental (tobacco smoke, vehicle exhaust, food
flavorings) sources. Acetaldehyde is shown to be carcinogenic in rodents and causes a variety
of DNA lesions in human cells [45;46]. e2G is the best known lesion formed by acetaldehyde
[47], and it has been found in the liver of ethanol-treated rodents and in the white blood cells
of human alcoholics [45].
It is believed that acetaldehyde first forms N2 -ethylidene-2'-
deoxyguanosine, which is then reduced to form e2G [48]. It is unclear how this reduction is
achieved in an intracellular context. e2G is also speculated to be formed by the reaction of lipid
peroxidation products with DNA [49]. In vitro studies, one of which shows that e2G can mispair
with a guanine in the presence of exonuclease-free Klenow fragment of pol I [50], suggest that
e2G is mutagenic [41;42]. How exactly this lesion contributes to carcinogenesis is unclear as it
is not known to be mutagenic in vivo. However, e2G is known to block several replicative
polymerases, and has shown to be bypassed by pol K [51], pol 1 [52], and pol
T1
[53], all in in
vitro studies. Previous in vivo work from our laboratory has shown that e2G is not mutagenic
but is toxic in all combinations of presence and absence of both AlkB and AidB [44]. Our study
seeks the answer to the question of whether e2G is a mutagenic and blocking lesion in vivo and
if so, whether it is bypassed by DinB.
By using site-specific mutagenesis, we inserted various N2-dG lesions at a specific location in
single-stranded M13 phage, which were then introduced into E. coli proficient or deficient for
DinB, and the mutation frequencies and bypass efficiencies across the lesions were assayed.
3.2 Materials and methods
The REAP and CRAB assays described here have been modified from the work of Delaney et al.
[54].
Please refer to the original method for more detail and clarification. The assays are
outlined in Figure 3.2.
3.2.1 Cell strains
All the E. coli strains used in this work contain the F' episome, which enables infection by M13
phage. GW5100 was used for large scale preparation of M13 phage DNA, SCS110 (JM110, end
Al) was used for amplification of progeny phage post-electroporation, and NR9050 was the
strain of choice for double agar plating with X-gal for blue-clear detection of plaques.
The E. coli strains used to prepare DinB- cell strains were HK81 (as AB1157, but na/A) and HK82
(as AB1157, but na/A alkB22; AlkB-deficient). The AIkB status of these strains was previously
confirmed by PCR amplification and sequencing of the region encoding the alkB gene (44;55].
P1 vir phage transduction was used to create dinB deficient versions of HK81 and HK82. Briefly,
1 ml of overnight saturated cultures of recipient cells (HK81 and HK82) was centrifuged and the
supernatant discarded. The cells were resuspended in 500 ul LB containing 10 mM MgSO 4 and
5 mM CaC12. Approximately 100 ul of these cells were mixed with 0, 25, 50, or 100 ul of P1
lysate (courtesy Dr. Jamie Foti/Dr. Graham Walker) containing a chloramphenicol resistance
gene (cam) flanked by frt sequences designed for insertion at the dinB site. After 30 min
incubation at 30 0C, 100 ul of 1 M sodium citrate was added to the cells, which stopped the
infection by chelating the Mg/Ca ions. The tubes were further incubated at 30 *Cfor 1 h to aid
the recovery and expression of the chloramphenicol gene. The entire volume of cells was then
plated (individually) on LB + chloramphenicol (10 ug/ml) plates using a glass spreader.
Following an overnight incubation at 37 0C, colonies were obtained, which were then replated
on LB + chloramphenicol plates.
3.2.2 Oligonucleotides
All oligonucleotides and primers were obtained from Integrated DNA Technologies (IDT) unless
specified otherwise. Sixteen-mer oligonucleotides of the sequence 5' GAAGACCTXGGCGTCC 3',
where X is the lesion of interest (m2G [44], e2G [44], THF [21], and FF [21]) were synthesized
and purified as described. Sixteen-mer oligonucleotides with the same sequence but with X =
G,A, T, or C,were used as a control. The 19-mer 'competitor' oligonucleotide of the sequence
5' GAAGACCTGGTAGCGCAGG 3' was used in the CRAB assay. Scaffold oligonucleotides (5' GGT
CTTCCACTGAATCATGGTCATAGC 3' and 5' AAAACGACGGCCAGTGAATTGGACGC 3') were used to
hold the 16-mers in place to the cleaved single-stranded M13 vector during genome
construction, and do not overlap the lesion-bearing region.
Primers used to confirm the DinB status of cell strains were a gift from Dr. Jamie Foti and were
of the sequence 5' GATTATGGTGCTGACCAAAAGTGCG 3' (DinB upstream primer) and 5' CGCTG
GCACTTAAGAGATATCCTGCGGG 3' (DinB downstream primer). The forward primers used for
the mutagenicity (REAP) and toxicity/bypass (CRAB) assays spanned the M13 vector as well as
the 5' end of the inserted oligonucleotide carrying the lesion of interest.
While the REAP
reverse primer also spanned the vector and the 3' end of the same oligonucleotide, thereby
effecting a selective amplification of DNA from the progeny phage resulting from only the
lesion-carrying genomes, the CRAB reverse primer annealed only to the M13 vector
downstream
of the inserted oligonucleotide.
The primers were modified with an
aminoethoxyethyl ether group (Y) at the 5' end to prevent labeling with
31P-y-ATP
in
subsequent reactions. The primers were of the sequence 5' YCAGCTATGACCATGATTCAGTGGA
AGAC 3' (CRAB forward primer; also used as the REAP forward primer), 5' YCAGGGTTTTCCCAGT
CACGACGTTGTAA-3' (CRAB reverse primer), and 5' YTGTAAAACGACGGCCAGTGAATTGGACG 3'
(REAP reverse primer).
3.2.3 Enzymes and chemicals
Pvull, XmnI, EcoRl, Haelll, BbsI, HinFI, T4 DNA Ligase, T4 DNA polymerase, BSA, and the enzyme
reaction buffers were from New England Biolabs. Shrimp alkaline phosphatase (SAP) was from
Roche. P1 nuclease, 5-bromo-4-chloro-3-indolyl- beta-D-galactopyranoside (X-gal), isopropyl $D-1-thiogalactopyranoside (IPTG) were from Sigma Aldrich. T4 Polynucleotide kinase was from
Affymetrix. Sephadex G-50 Fine resin was from Amersham Biosciences . Hydroxylapatite resin,
19:1 acrylamide:bisacrylamide solution, and N,N,N',N'-tetra-methyl-ethylenediamine (TEMED)
were from Bio-Rad. Phenol:chloroform:isoamyl alcohol (25:24:1; pH 8) was from Invitrogen.
3P-y-ATP
was from Perkin Elmer. Non-radioactive ATP was from GE Healthcare Lifesciences.
3.2.4 Double agar overlay plaque method for phage analysis
The double agar overlay method used in this work was adapted from Adams et al. [56]. This
method was used for enumerating initial electroporation events as well as phage titers to
ensure statistical robustness, but not for mutational analyses. Briefly, 10 ml of 2 x YT media
was inoculated with 2 ml of a saturated overnight culture of NR9050 and grown for 1 h at 37 *C
with aeration. Three hundred ul of this culture was mixed with 10 ul IPTG (24 mg/ml), 25 ul of 1
% thiamine, and 40 ul X-Gal (40 mg/ml in DMF), and then added to 2.5 ml of top agar
maintained in a molten state at 52 0C. Appropriate dilutions of supernatants containing phage
particles were immediately mixed with the top agar and poured onto B-broth plates, which
were shaken to evenly spread the agar. After a 10 min incubation at room temperature (to
allow the top agar to solidify), the plates were incubated overnight at 37 *Cto obtain dark blue,
light blue, or clear plaques.
3.2.5 M13 phage DNA
M13mp7(L2) phage single-stranded DNA was isolated as follows.
Various dilutions of a
previous stock of M13 phage supernatant were plated on a lawn of E. coli cells using the double
agar overlay method to obtain phage plaques. A well-isolated plaque was plugged using a
sterile Pasteur pipette and vortexed in 1 ml LB, 200 ul of which was used to make a starter
culture (grown overnight) by mixing with 10 ul of an overnight saturated culture of GW5100
cells in 10 ml LB. One milliliter of this phage starter culture was then used to inoculate GW5100
cells, which had been grown using 500 ul of an overnight saturated culture in 250 ml of fresh 2
x YT medium for 2 h at 37 'C and shaken at 275 rpm. The inoculated culture was grown further
for 8 h at 37 *Cwith aeration, after which the cells were pelleted and discarded. The phage
were precipitated from the supernatant by addition of 4 % PEG 8000 MW and 0.5 M NaCl.
After overnight precipation at 4 *C,the phage were pelleted, resuspended in 5 ml TE pH 8, and
extracted with four washes of 3 ml 25:24:1 phenol:chloroform:isoamyl alcohol (Invitrogen, pH
8). The aqueous phase was passed through a 0.5 g hydroxylapatite column (BioRad), washed
with 5 ml TE, and eluted in 1 ml fractions with 12 ml of 0.16 M phosphate buffer. The DNAcontaining fractions were identified by spotting on an agarose plate containing ethidium
bromide. The phosphate buffer in those fractions was then exchanged for TE by three washes
in Microsep 100K spin dialysis columns (Pall Lifesciences). The DNA obtained was at a yield of>
1 pmol/ml 2 x YT and was stored at -20 0Cuntil further use.
3.2.6 Construction of genomes
The multiple cloning site in single-stranded M13mp7(L2) is designed to form a hairpin structure
that contains a functional EcoRl site. Twenty picomoles of M13 single-stranded DNA were
linearized by incubation with 40 U of EcoRl for 8 h at 23 *C. Scaffolds (25 pmol in 1 ul each)
were annealed to the ends of the linearized genome by incubation at 50 0C for 5 min followed
by cooling to 0 0C over 50 min. In addition, 30 pmol of each 16-mer oligonucleotides insert
were 5' phosphorylated by 15 U of T4 PNK, supplemented with 1x T4 PNK buffer, 1 mM ATP,
and 5 mM DTT and incubated at 37 *Cfor 1 h. The linearized genome was subsequently ligated
with the phosphorylated oligonucleotide for 8 h at 16 0C in a reaction volume of 75 ul
containing 1 mM ATP, 10 mM DTT, 25 ug/ml BSA, and 800 U T4 DNA ligase. To degrade
scaffolds and unligated oligonucleotides, the ligation mixture was treated with 0.25 U/ul T4
DNA polymerase for 4 h at 37 *C. Finally, the reaction volume was brought up to 110 ul with
water and extracted once with 100 ul 25:24:1 phenol:chloroform:isoamyl alcohol. The aqueous
phase was purified by three TE (pH 8) washes in Microsep 1OOK spin dialysis columns (Pall
Lifesciences) to remove any residual phenol and salts.
Recovery yields of 30-45 % were
obtained.
3.2.7 Genome validation and normalization
Prior to proceeding with the bypass and mutagenicity assays, the incorporation of lesioncontaining oligonucleotides was confirmed and the relative concentration of the constructed
genomes was determined and normalized using the following procedure:
A 10-fold molar
excess of scaffolds (previously used in constructing the genomes) were annealed to ~0.35 pmol
of genomes in 5 ul.
The genomes were cleaved with 10 U of HinFI and the resulting 5' end
dephosphorylated with 1 U of Shrimp alkaline phosphatase in a reaction volume of 8 ul by
incubation at 37 *Cfor 1 h followed by a 5 min incubation at 80 0Cand cooling down to 20 *C@
0.2 *C/s. The 5' ends were then labeled with 1.66 pmol of 3 1P-y-ATP (6000 Ci/mmol) in a total
reaction volume of 12 ul containing 1 x buffer 2, 5 mM DTT, 150 pmol cold ATP, 5 U T4 PNK, and
10 U Haelll, incubated at 37 *Cfor 1 h. The reaction was stopped by the addition of 12 ul 2 x
formamide loading buffer, and the products were resolved using 20 % PAGE until the xylene
cyanol dye migrated a distance of 12 cm. The bands corresponding to fully-ligated genomes
were then quantified using phosphoimagery and normalized with respect to one another. The
genomes were then diluted with water such that all the genomes were at the same final
concentration. The band generated from the competitor genome was used as a marker.
Post-normalization, a test electroporation was performed in HK81 competent cells using the
control genome mixed in different ratios with the competitor genome. The results of the test
electroporation were determined by plating the competent cells immediately after
electroporation using the phage-overlay method to yield a dark blue (control): light blue
(competitor) plaque count ratio. The ratio that yielded a 75:25 dark blue:light blue phage count
was selected for the bypass assay of the lesion-containing genomes.
3.2.8 Preparation of electrocompetent cells
Three baffled flasks containing 150 ml LB medium each were inoculated with 1.5 ml of a
saturated overnight culture of the strain to be transformed. The cultures were incubated at 37
*Cand shaken at 275 rpm for ~ 2.5 h until the cultures reached early log phase, as measured by
OD600 of ~0.5. The cell were then pelleted by centrifugation at 9500 rpm (Sorvall GSA rotor),
resuspended in 1 ml cold sterile water, and pooled to a final volume of 175 ml cold sterile
water. This process of washing, pelleting, and resuspending was repeated three times. The
final resuspension was in 4.8 ml 10% glycerol to obtain a final volume of 6 ml electrocompetent
cells, which were then aliquoted and stored at -20 *Cprior to use.
3.2.9 CRAB assay
Genomes containing the lesions were mixed in a 75:25 ratio with the competitor genome in a
total volume of 6 ul and electroporated in triplicate into 100 ul competent cells in a 2 mm-gap
cuvette using 2.5 kV and 125 Q. The cells were immediately transferred to 10 ml LB and an
aliquot of the freshly electroporated cells was immediately plated using the agar overlay
method to ensure that a minimum of 105 independent initial electroporation events occurred in
10 ml of culture. The cells were then grown for 6 h at 37 "Cwith aeration to amplify progeny
phage. The supernatants of the 6 h cultures were retained and plated using the agar overlay
method to confirm 10 4 -fold amplification in the progeny phage titer. Another round of
amplification was performed in order to ensure that the progeny phage being analyzed came
from genomes that entered the E. coli cells, and not the residual genomes that did not get
electroporated into cells but were still present in the milieu, since PCR is subsequently used.
This was done by infecting 10 ul of an overnight culture of SCS110 cells with 100 ul of the 6 h
supernatants in 10 ml LB and incubating for 7 h at 37 0C with aeration, after which the
supernatant was retained. Single-stranded M13 progeny phage DNA was isolated from 0.7 ml
of supernatant using a QlAprep Spin M13 Kit with final DNA suspension in 100 ul elution buffer.
CRAB forward and reverse primers were used to amplify the region of interest from 10 ul per
QlAprep elution sample in a total volume of 25 ul using 1.25U Pfu Turbo DNA polymerase, 25
mM of each dNTP, and 10 x Pfu Turbo buffer. The PCR program started by denaturing at 94 *C
for 5 min, then cycled 30 times at 94 "Cfor 30 s, 67 *Cfor 1 min, and 72 0Cfor 1 min, and finally
extended for 5 min at 72 *C . The volume was then made up to 110 ul using water and
extracted once using 25:24:1 phenol:chloroform:isoamyl alcohol to destroy the DNA
polymerase (including the exonuclease domain). The aqueous phase was passed through a
Sephadex G-50 Fine resin spin column to remove any remaining dNTPs and traces of phenol.
The purified PCR product was then treated with Bbsl (1.5 U for 4 ul of sample in a total volume
of 6 ul) and shrimp alkaline phosphatase (0.3 U) by incubating at 37 *Cfor 4 h, heating to 80 *C
for 5 min, and cooling to 20 *C@ 0.2 *C/s. The 5' ends were then labeled in a total volume of 8
ul with a mixture of non-radioactive ATP (20 pmol), y-32P-ATP (1.66 pmol of 10 uCi/ul at 6000
Ci/mmol), and 5 U of Optikinase/T4 PNK by incubation at 37 *Cfor 15 min, followed by 65 *Cfor
20 min and cooling to 23 *C@ 0.1 *C/s. The labeled product was then trimmed by Haell (10 U
in a final volume of 10 ul) at 37 *Cfor 2 h, followed by addition of 10 ul 2 x formamide loading
dye, which quenched the reaction. The samples were then loaded onto a 20 % denaturing gel
and run for ~3.5 h at 550 V, until the xylene cyanol dye migrated 10.5 cm. The gel was then
exposed to a phosphoimager screen and quantified using a Storm 840 scanner. Band intensities
were quantified using ImageQuant software, and lesion bypass was measured by comparison of
the 18-mer band intensity (lesion signal) to the 21-mer band intensity (competitor signal).
3.2.10 REAP assay
The REAP assay methodology is identical to the CRAB assay except for the PCR primers. The
primers used for the REAP assay span both the vector as well as the insert at each end, thereby
effecting a selective amplification of DNA from only the progeny phage resulting from the
lesion-carrying genomes. Following electrophoresis, the 18-mer bands were excised from the
gel, and crushed and soaked overnight in 200 ul water. After desalting with Sephadex G-50 Fine
resin spin columns, the samples were lyophilized overnight to dryness, resuspended in 5 ul
containing 1 ug P1 Nuclease in 30 mM sodium acetate and 100 mM zinc chloride, and incubated
at 50 *Cfor 1 h. One ul of each sample was then spotted onto PEI-TLC plates and separated
using 200 ml of a saturated solution of (NH4) 2HPO 4 adjusted to pH 5.8.
After 12 h of
development, the TLC plates were air-dried and quantified using phosphoimagery.
3.3 Results
3.3.1 Creation of DinB- cell strains
P1 vir transduction was used to create DinB- cell strains as described in the methods section. To
verify the DinB knockout status of the genetically engineering E. coli, chloramphenicol-resistant
colonies were replated. Colonies that grew after the second plating were picked and tested for
dinB absence by colony PCR followed by sequencing with DinB upstream and downstream
primers.
Restriction digestions with Pvull (single-site cutter for cam) and Xmnl (single-site
cutter for dinB) also confirmed the replacement of dinB by cam in the successful transductants.
Colony PCR was also performed on the DinB+ parent cell strains with the same set of primers
and the products were used as controls for the restriction digestions with Pvull and Xmnl. The
new cell strains were named HK83 (HK81 AdinB::frt cat; dinB deficient) and HK84 (HK82 AdinB::
frt cat; alkB and dinB deficient).
3.3.2 CRAB assay
The competitive replication of adduct bypass (CRAB) assay is a quantitative method that
determines to what extent a given lesion blocks DNA replication if left unrepaired. In essence, a
lesion-bearing genome is mixed with a nonlesion competitor genome in a specific input ratio
and passaged through E. coli cells of a given repair background. The output ratio of progeny
phage indicates the relative growth of the lesion-bearing genome with respect to the nonlesion
competitor genome; a decrease in the lesion: competitor output ratio (and hence the bypass
efficiency) signifies blockage of replication and therefore toxicity. The competitor genome acts
as an internal standard in this competitive assay.
The results of the CRAB assay are summarized in Figure 3.3 and Table 3.1. In DinB- cells, THF
and FF show bypass efficiencies of 28 % each. These results indicate that these lesions are
relatively strong blocks to replication in the absence of DinB. However, they are not a complete
block to replication like m3C is in the absence of AIkB, or as was seen in the case of 1methylguanine and 3-methylthymine in a previous study [57]. The presence of DinB more than
triples the bypass efficiencies of THF and FF to ~100 % (p value = 0.001) and ~90 % (p value =
0.003) respectively, thus relieving the replication block. The effect of DinB on the bypass of
m2G and e2G is negligible and not statistically significant.
3.3.3 REAP assay
The restriction endonuclease and postlabeling (REAP) assay outlined in Figure 3.2 determines
the mutation frequency and mutation composition after the lesion of interest, if not yet
repaired, has been processed by the intracellular replication machinery. None of the N2-dG
lesions are significantly mutagenic in the presence or absence of DinB (Figure 3.4 and Table
3.2). THF and FF show mutation frequencies of ~1 % in both the presence and absence of DinB,
while m2G and e2G also show a small mutation frequency of 3 % and 2 %, respectively, in either
cell background. These numbers do not qualify these lesions as mutagenic, and show no effect
of the presence or absence of DinB on the mutation frequencies of said lesions.
3.4 Discussion
We investigated the role of the error-prone E. coli DinB DNA polymerase in bypassing a
spectrum of N2 -dG lesions. The most important finding in our study is that the N2 -furfuryl-dG
lesion and its saturated homolog are not mutagenic but are toxic to E. coli when introduced in
DinB- cells on a single-stranded vector. This toxicity is alleviated by DinB in vivo. Also, this
effect is seen without a change in the mutational profile of the lesions, most likely due to the
insertion of the correct base (cytosine) opposite the lesions by DinB. This finding supports the
in vitro results obtained by Jarosz et al. [14;21]. This finding also provides support for the role
of DinB in transcription-coupled translesion synthesis across N2 -dG lesions formed by NFZ (see
Chapter 1 for a detailed review). However, there seems to be another mechanism at play that
enables lesion tolerance in the absence of DinB to the extent seen in our assays, since the level
of bypass of THF and FF in the absence of DinB (28 %) is much higher than that seen for other
lesions previously, for example, ~5 % for tetrahydrofuran, an abasic site analog [58].
It is
necessary to point out that while it has been proposed that nucleotide excision repair (and not
TLS) might be the primary repair pathway that deals with NFZ-induced damage [59], it requires
a double-stranded DNA context which is obviated by our experimental system.
We also
analyzed the N2 -dG lesions in AlkB+ and AIkB- cells on the off chance that AIkB could perform a
direct reversal of the alkyl adducts. However, no effect of the enzyme was seen on the
mutagenicity or toxicity of these lesions (data shown in Chapter 5).
Interestingly, m2G and e2G do not block replication in vivo in E. coli. Since no toxicity is seen in
DinB negative cells, the presence of DinB does not show any measurable change in the bypass
efficiency of m2G and e2G. This result isin contrast with what has been observed for e2G in in
vitro assays with other Y-family and replicative polymerases [51-53]. While the mutagenicity
data for m2G and e2G correlate well with that seen in a previous study from our laboratory, the
bypass data are in contrast with the same study (~ 35 - 50 % bypass for both lesions) [44]. The
same DinB+ cell strain and same batch of m2G and e2G oligonucleotides were used both in our
study as well as the previous study. We do not have a good explanation for this discrepancy;
however, the data obtained in the current work are internally consistent. It could be that there
is another enzyme which preferentially and efficiently repairs or bypasses these lesions such
that the supplementary role of DinB in the bypass of m2G and e2G is overshadowed beyond
detection by our assay.
It is possible that DinB is tolerant of N2-dG lesions and hence results in a non-toxic phenotype in
DinB+ cells. N2-dG lesions occupy the minor groove of DNA [60] and can interfere with
polymerase-minor groove interactions [61-63] if the alkyl group swivels into the vicinity of N3
atom of guanine. Several B-family polymerases are known to have a conserved motif that scans
the DNA minor groove for lesions and misincorporations [64], which is lacking in the Y-family
DNA polymerases. It is speculated that this may be the case for pot
K,
as deduced from x-ray
crystal structure studies of the catalytic core of the polymerase with a primer-template DNA
and an incoming nucleotide, which reveal a lack of steric hindrance in the minor groove at the
primer-template junction [65]. It is proposed that DinB can accommodate minor groove lesions
to enable bypass with correct base pairing, even lesions with alkyl groups much bigger than
those of the lesions in the current study (such as BaP) [66].
That the lesions are not mutagenic in any repair background can be explained by the availability
of a hydrogen at the N2 position of guanine and the possibility of free rotation around the
nitrogen-carbon bond. Since the addition of a chemical moiety at the N2 position of guanine
still leaves one hydrogen atom intact on the nitrogen, correct base pairing with cytosine is
possible. In addition, free rotation around the carbon-nitrogen bond would enable the
extraneous alkyl group to swivel away from the base pairing face of guanine, thus alleviating
2
any steric hindrance caused by attachments to the N2 position. Thus, smaller N -dG alkyl groups
should not interfere with Watson-Crick base pairing, while the bulky attachments should be
accommodated by DinB to allow correct base pairing. If steric hindrance cannot be relieved by
this mechanism, a wobble base pair can be formed between an NdG lesion and the incoming
cytosine. This is proposed to be the mechanism by which yeast pol '1 (pol V homolog) and
human pol K bypass such lesions [52]. Furthermore, the formation of Hoogsteen base pairs of
N2 -dG lesions with cytosine where the lesion-bearing guanine is rotated to the syn
conformation has been observed with pol t. Specifically, this has been observed in the case of
m2G [42], and e2G [52] where the ethyl group is positioned into the major groove of DNA to
enable bypass with correct base pairing. However, for an N2 -dG lesion to correctly pair with a
cytosine using either mechanism, the latter has to be in its imine tautomer form (for wobble
base pairing) or its protonated form (for Hoogsteen base pairing). Additionally, while it has
been shown that e2G is mutagenic when encountered by exonuclease-free Klenow fragment of
pol I [50], this may not hold true for pol Ill which is the main replicative enzyme in E. coli that
processes the M13 genome. Our results prove e2G to be non-mutagenic in vivo.
In summary, we have shown that DinB does indeed bypass the N2 -furfuryl-dG lesion in vivo.
Taken together with the in vitro data available for DinB and its homologs, it is highly possible
that these Y-family polymerases bypass the major adduct of NFZ in vivo. Given that NFZ is an
antibiotic agent, DinB may be the shield in the arsenal of E. coli against this type of 'chemical
warfare' from other species. In addition, we have also found that DinB does not have a role in
alleviating any mutagenicity or toxicity of other N2-dG lesions such as e2G, which is in contrast
with what has been observed in previous studies. Lastly, this is the first study to our knowledge
that explores the effect of DinB on the mutagenicity and toxicity of m2G in DNA in vivo. We
conclude that m2G is neither mutagenic nor toxic when introduced into DNA, and that this
result is not mediated by DinB.
3.5 Figures
Figure 3.1: Structures of lesions used in the DinB study. The lesions are referred to by the
following abbreviations in the text: FF: 2-furan-2-yl-methylguanine, THF: 2-tetrahydrofuran-2yl-methylguanine, m2G: 2-methylguanine, e2G: 2-ethylguanine, and G: unmodified guanine.
0
N
N
IN
NH
N
N
H
0
0
DNA
THF
FF
N
KN I
-'NH
'NH
N
DNA
N
H
m2G
N
DI
DNA
N
N4
H
e2G
0
N
N NH
N
N
DNA
NH2
100
Figure 3.2: Overview of the CRAB bypass and REAP mutagenesis assays. Courtesy of Dr. James
Delaney [67].
(+3)
GAAGACCTGGCGTCC
If lesion hinders replication
Lesion
Lesion
Competitor
Genome
Genome
Genome
10%
20%
Competitor
Genome
90%
Input
Cells
GAA( ACTXGGCGTCC
(
80%
Output
REAP
mutagenesis assay
CRAB
bypass efficiency assay
{Lesion
Genome)
GAAGACCTGGTAGCGCAG
GAAGACCTXGGCGTCC
GAAGACCTGGTAGCGCAGG
GAAGACCTXGGCGTCC
GAAGACCTGGTAGCAG
Competitor
Genome)
Genome
Genome
80%
20%
80%
Lesion
AAGACCTG4GTAGCGCAG
20%
Competitor
{
PCR amplify with primers designed to amplify
only region that had contained lesion
PCR amplify with primers designed
to amplify lesion and competitor equally
80% competitor signal (3 bases longer)
100% lesion signal
61 mer
20% lesion signal
For CRAl&REAP analyss:
Cut with Bbsl & dephosphorylate
"P(@)-Label new 5' ends &Haelll trim
Separate products via PAGE
RNt
Bbsl
34mer
30mer
Shown in detail for REAP (right)..
CRAB works similarly CRAB
lesion or non-lesioncontrol signal
m
13mer
[11100ll- NH2
13mer
18mer
Excise oligo whose 2P-labeled
5' base was the lesion site
Digest with Nuclease P1
Resolve 5'dNMPs via TLC
Quantify via Phosphorimagery
-21mer
--
Hal Il
14mer
REAP
011
- 34me rII
+3 competitor signal-*
+
18mer
Quantify frameshifts,
(..
Lesion signal
(non-lesion control
its competitor
x 100%
/
if applicable
0
Mutation
+3 competitor signal /
Bypass =
.)
0
5' dCMP
5' dTMP
5'dGMP
5' dAMP
101
Figure 3.3: Bypass efficiencies of N2-dG lesions in DinB+ and DinB- cells as determined by the
CRAB assay. THF and FF pose a relatively strong block to replication as seen by the low bypass
efficiencies in the absence of DinB. In DinB+ cells, the bypass efficiencies are trebled, thereby
relieving the toxicity. The data are also tabulated in Table 3.1.
'Bypass' of N2-dG lesions in DinB+/DinB- cells
140
*:p-valalue
=0.001
120
*1
120
a DinB=.0
-
DinB+
=0.003
-
100
4
80
60
40
20
0
m2G
e2G
THF
FF
G
102
Figure 3.4: Mutagenicity of the N2-dG lesions as determined by the REAP assay. None of the
lesions show any significant mutations in either DinB+ or DinB- strains. The data are also
tabulated in Table 3.2.
Mutation frequency in DinB+ cells
100%
NG
mA
90%
UT
80%
mC
70%
60%
50%
40%
30%
20%
10%
0%
m--
-
m2G
e2G
THF
FF
TGG
GATC
Mutation frequency in DinB- cells
100%
NG
SA
90%
OT
80%
NC
70%
60%
50%
40%
30%
20%
10%
0%
I
--
m2G
e2G
THF
FF
TGG
GATC
103
3.6 Tables
Table 3.1: Bypass efficiencies of N2 -dG lesions in DinB+ and DinB- cells as determined by the
CRAB assay. The data tabulated below are shown in Figure 3.3.
Lesion/base
Bypass in DinB- cells
Avg.
Std. Dev.
m2G
e2G
THF
FF
G
Bypass in DinB+ cells
Avg.
Std. Dev.
94.64
3.23
122.17
6.73
121.19
27.92
28.06
100.00
3.92
2.39
2.85
0.00
124.20
106.57
88.25
100.00
15.30
13.15
15.46
0.00
Table 3.2: Mutagenicity of the N2-dG lesions as determined by the REAP assay. None of the
lesions show any significant mutations in either DinB+ or DinB- strains. The data tabulated
below are shown in Figure 3.4.
(a)
DinB+ cells
Lesion/base
m2G
e2G
THF
FF
TGG
GATC
%G
96.73
97.20
98.35
98.31
98.28
16.16
Average
%A
%T
2.86
0.11
2.11
0.09
0.53
0.18
0.61
0.15
0.90
0.07
32.75 31.67
%C
0.30
0.60
0.93
0.93
0.75
19.42
Standard
%G %A
0.31 0.26
0.22 0.18
0.36 0.15
0.57 0.19
1.03 0.45
1.87 1.88
deviation
%T %C
0.04 0.11
0.03 0.05
0.07 0.34
0.04 0.38
0.04 0.54
0.99 0.49
%G
96.93
98.73
98.75
99.02
99.45
20.69
Average
%A
%T
2.49
0.23
0.91
0.08
0.39
0.39
0.21
0.57
0.36
0.06
26.83 32.99
%C
0.36
0.28
0.47
0.21
0.13
19.49
Standard
%G %A
0.76 0.75
0.20 0.08
0.37 0.22
0.12 0.05
0.12 0.06
1.15 1.42
deviation
%T %C
0.16 0.15
0.02 0.13
0.05 0.23
0.04 0.11
0.01 0.07
0.79 2.50
(b) DinB- cells
Lesion/base
m2G
e2G
THF
FF
TGG
GATC
104
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mispair formation and mispair extension is dependent on the interaction between the
minor groove of the primer terminus and Arg668 of DNA polymerase I of Escherichia
coli. Biochemistry, 44, 5647-5659.
64. Swan,M.K., Johnson,R.E., Prakash,L., Prakash,S., and Aggarwal,A.K. (2009) Structural
basis of high-fidelity DNA synthesis by yeast DNA polymerase 6. Nat.Struct.Mol.Biol., 16,
979-986.
65. Lone,S., Townson,S.A., Uljon,S.N., Johnson,R.E., Brahma,A., Nair,D.T., Prakash,S.,
Prakash,L., and Aggarwal,A.K. (2007) Human DNA polymerase K encircles DNA:
implications for mismatch extension and lesion bypass. Molecular Cell, 25, 601-614.
66. Chandani,S. and Loechler,E.L. (2009) Y-Family DNA polymerases may use two different
dNTP shapes for insertion: a hypothesis and its implications. JMol Graph.Model., 27,
759-769.
67. Delaney,J.C. and Essigmann,J.M. (2006) Assays for determining lesion bypass efficiency
and mutagenicity of site-specific DNA lesions in vivo. Methods Enzymol., 408, 1-15.
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Chapter 4:
DNA lesion placement relative to the origin of replication and
polymerase obstacles as variables in 0-methylguanine mutagenesis
in vivo
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4.1 Introduction
It is known that the local DNA sequence, usually spanning a few base pairs flanking a chemicalDNA lesion, affects replication error and lesion repair rates. Studies have also found differences
in rates of errors on leading and lagging strands [1].
Functional elements a considerable
distance from the site, such as the origin of replication, may also affect mutagenesis. Pavlov et
al. have shown that the mutation frequency caused
by 8-hydroxyguanine and 6-
hydroxylaminopurine in yeast genomes was influenced in part by the distance from the origin.
The study also indicates that replication origins may establish a strand bias for adduct-induced
mutations [2]. This result suggests that the mutagenic and carcinogenic risk posed by adducts
might be partly determined by the location of susceptible genes relative to origins of
replication. This result may also have an evolutionary significance, as switching off origins of
replication could change the mutability of nearby genes [2].
Several studies comparing sequences of homologous genes of various bacteria have shown that
the distance from the origin of replication influences mutation frequency [1;3-5}.
In a
comparison between homologous genes of E. coli and S. enterica, Sharp et al. have reported a
higher synonymous substitution rate in genes farthest from the origin of replication. However,
this could result from a higher rate of post-replicative recombinational repair closer to the
origin [5]. Furthermore, it has been found that the substitution rates for Mycobacteria show
the opposite trend [3]. A study to infer spontaneous mutation patterns using pseudogenes of
M. leprae by Mitchell et al. reported a negative correlation of G -> T transversions on the
leading strand with distance from the ori [4]. Another study done by Hudson et al. assayed
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reversion rates of lacZ alleles inserted at different positions between the origin and termination
sites in the S. enterica chromosome and found that the highest mutation rates were at an
intermediate locus, concluding that mutation rates do not increase with distance from the
origin [1].
Thus, there are conflicting data and opinions in the literature on the effect of
distance from the origin of replication on mutation frequency.
While some of the aforementioned studies cite an increase in mutation rates and/or decrease
in repair rates with distance from the ori as possible reasons for their observations, neither has
been experimentally probed. This project aims to explore mutation frequency as a function of
distance from the origin of replication, as well as kinetics of repair. The hypothesis of this
project is that the mutation frequency )f a given pro-mutagenic DNA lesion is a factor of its
distance from the origin of replication. This hypothesis is based upon the expectation that a
DNA lesion at a distal site has more time to be repaired than one at a site more proximal to the
origin of replication. We investigated this hypothesis by modulating the time available for
repair of a lesion, O6 methylguanine, by (a) constructing a modified M13 genome to
incorporate a second site of insertion, (b) using the modified M13 to evaluate the effect of
distance of the lesion from the origin of replication on its repair, and (c) introducing replication
stalling elements upstream of the lesion site. The time available for lesion repair can also be
modulated (d) using kinetically slow DNA polymerase IlIl and Ada enzymes, and (e) changing the
intracellular concentration of repair enzymes Ada and Ogt. The focus of this thesis is on
exploring the hypothesis via (a), (b), and (c), and the experimental outline towards this end is
summarized in Figure 4.1. Parts (d)and (e) are discussed briefly in future work.
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4.1.1 M13 - life cycle, genes and possible sites for genetic manipulation
M13 is a member of the filamentous Ff phages (fd, f1) that infect E. coli bearing the F pilus and
has been used widely for cloning and phage display [6-8]. Wild-type (wt) M13 contains a singlestranded circular genome, 6407 nucleotides long, encoding ten genes (Figure 4.2a), the
functions of which are listed in Table 4.1. M13 encodes two origins of replication: one for the
minus (transcribed) strand and one for the plus (template) strand. For the work described
herein, we are concerned with only the plus strand origin since that isthe form of M13 used in
all our experiments.
The mode of replication of M13 makes genetic manipulations and experimental studies possible
in both double-stranded and single-stranded DNA.
The life cycle of the virus begins with
adsorption to the F pilus of the host cell and insertion of single-stranded circular DNA into the
bacterium. The first step in replication of the virus is the conversion of viral single-stranded
DNA (plus strand) to double-stranded replicative form (RF) DNA. This is initiated by E. coli RNA
polymerase in the presence of E. coli single-stranded binding protein (SSB) by production of an
RNA primer at the plus strand ori. Extension of the primer to produce the RF DNA is done by
DNA polymerase l1l, followed by ligation and supercoiling by host enzymes. The second step is
the rolling circle form of replication which is initiated by specific nicking of RF at the plus strand
origin by gp 11endonuclease. The 3'-OH is then elongated by pol Ill displacing the plus strand. A
second gp 11endonuclease cleavage liberates the single-stranded DNA which is subsequently
ligated into a circle, after which RNA priming can occur to repeat the cycle. Approximately 100
copies of the RF DNA are made.
The viral coat proteins are concurrently transcribed and
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translated, until gp V levels reach a certain level [9]. In the third step, gp V single-stranded DNA
binding protein coats this displaced viral strand to block complementary strand synthesis and
enable packaging of the nucleoprotein. gp X is also proposed to play a role in the switch from
synthesis to packaging [6]. M13 causes a chronic infection and is released continuously from
growing E. coli without lysing the cells, with an average of 500 phage particles produced per
infected E. coli cell. Superinfection is prevented by blocking the synthesis of the F pilus [10].
Assays developed in our laboratory for mutagenesis and toxicity studies of lesions employ a
variant of M13, viz., M13mp7(L2), which differs from wt M13 in that it is 7214 nucleotides long
due to insertion of lacZ (containing a multiple cloning site (MCS)) in the intergenic region
between genes IV and II. Figure 4.2a shows the map of M13mp7(L2). Around 89% of the
genome encodes proteins in a non-overlapping manner with very few untranslated nucleotides
in between, and the intergenic spaces usually contain other important genetic elements such as
gene promoters, and transcription termination sites. The two intergenic regions are between
genes IV and II, and between genes Vill and Ill. The former contains the ori and phage
packaging signal while the latter contains a terminator of transcription [7]. Thus, the possible
sites for insertion of oligonucleotides for this study are in the MCS, downstream of (or within)
lacZ, or at the 5' end of (or within) gene Ill. The intergenic region between genes IV and II was
employed initially to insert lacZ without disruption of phage replication and survival elements
[7]; hence, further insertions are possible upstream of lacZ. Insertions at the N-terminal sites of
gene Ill [11] and Vill [8] have been used to create gene fusions for phage display, since these
gene products are coat proteins. However, insertions larger than 6-8 amino acid residues are
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not tolerated for gene VIII [8]. It is possible to insert longer oligonucleotides coding for longer
proteins at the 5' end of gene Ill, as long as they do not disrupt the reading frame.
4.1.2 Lesion of interest: 06-methylguanine
Alkylating agents react with DNA to produce various lesions that can be lethal and/or
mutagenic. Almost all oxygen and nitrogen atoms in DNA can be modified by methylating
agents to form 14 known primary lesions [12]. The sugar-phosphate backbone of DNA can also
be alkylated to form lesions like methylphosphotriesters (MePT) [13]. Some lesions such as 3methyladenine and 1-methyladenine inhibit DNA replication and are therefore cytotoxic, while
lesions like 0 6-methylguanine (06MeG), 04-methylthymine (04MeT), and 3-methylcytosine are
mutagenic as well as lethal [13]. 06MeG, which causes GC->AT transitions [14], is believed to
be the primary mutagenic lesion formed by alkylation damage [12]. It can also lead to sisterchromatid exchanges, chromosome aberrations, double-strand breaks, and toxicity via 'futile'
mismatch repair cycles if left unrepaired [15-18]. 06MeG is formed from both endogenous
[19;20] and exogenous sources [21] and studies have correlated its persistence to organspecific tumorgenicity in rats [22]. We chose O6 MeG as the lesion of interest in this study since
its mutagenic and toxic effects have been widely investigated, and the results from this study
can be compared to those in the literature as well as to studies done previously in our
laboratory [14;23;24].
The repair of O6 MeG in various sequence contexts has also been
explored [24], and this study will compare repair at different distances from the ori in these
sequence contexts.
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4.1.3 Repair of O6 MeG by Ada and Ogt
E. coli has two 06 MeG-DNA-methyltransferases, the constitutive Ogt protein and the inducible
Ada protein, which directly reverse methylation damage by transferring the alkyl group to one
of the internal cysteine residues on the protein.
This transfer irreversibly inactivates the
enzyme making the reaction stoichiometric [13]. The alkyl groups from 0 6-AlkGua and 04AlkThy are transferred to Cys 321 at the C-terminus, while those from MePT are transferred to
Cys 38 at the N-terminus of Ada. Ada is part of the adaptive response, which was discovered
when E. coli treated with a low dose of a methylating agent acquired resistance to the
mutagenicity and toxicity of subsequent higher doses [25].
Methylation of Cys38 of Ada
converts it to a transcriptional activator of the genes forming the adaptive response, ada, alkA,
alkB and aidB.
AlkA serves as a DNA-glycosylase while AIkB is an iron- and a-ketoglutarate-
dependent dioxygenase, both of which repair several alkylated purines and pyrimidines. AidB is
a flavin adenine dinucleotide (FAD)- containing protein speculated to provide resistance to
certain methylating agents but not through direct DNA repair [13;26]. The number of Ada
molecules is estimated to rise from 1-2 molecules in an unadapted state to ~3000 molecules in
a fully adapted cell [27;28]. It has also been shown that levels of Ada increase by 20-fold in
uninduced cells as they enter stationary phase [29]. The second DNA methyltransferase, Ogt,
was discovered by deletion of the ado operon [30;31].
Unlike Ada, Ogt is constitutively
expressed in E. coli, shows a preference for repair of 0 4 MeT and larger alkyl adducts, and does
not repair MePT [31]. It is estimated that there are ~30 molecules of Ogt in wt E. coli [28].
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4.1.4 Repair of O6MeG by the nucleotide excision repair and mismatch repair pathways
In addition to the methyltransferases, the UvrABC nucleotide excision repair (NER) pathway can
also repair 06 MeG. Excision of O6MeG on duplex substrates has been shown to occur in vitro
[32] and in vivo [33]. In the first hour of alkylation, O6MeG was repaired by nucleotide excision
unless the adaptive response had been induced prior to exposure or was constitutively
expressed. Ogt is speculated to provide protection at low levels of exposure, and the adaptive
response against chronic exposure. However, the uvr pathway seems to be the main line of
defense against O6MeG in E. coli for transient exposures to alkylating agents [33].
These
studies have been done in double-stranded DNA and we do not expect the uvr pathway to
contribute significantly to O6MeG repair in our single-stranded system. If repair does occur by
excision, it will be after the RF DNA is made, by which time a mutation would have been
incorporated by translesion polymerases. Nonetheless, there is a possibility that NER may
affect the mutation frequencies obtained from repair of O6MeG. Chambers et al. found a 40
fold decrease in the G->A transition caused by an O6MeG lesion on a <DX174 genome in an NER
deficient (uvrA) cell strain vs. wt [34]. The authors suggested a shielding mechanism by which
uvrA binds to the lesion and protects it from repair by Ada or Ogt, leading to higher mutation
frequencies. Studies done in our lab suggest that nucleotide excision repair does not affect
mutation frequency of O6 MeG in an M13 single-stranded context. The mutation frequency of
0 MeG in uvrBada ogt cells was very similar to that in uvrB adacogt cells [24]. However, a
better comparison would be between uvrB* ada'ogt~ and uvrB ada'ogt cells, as is suggested in
the discussion.
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The mismatch repair (MMR) pathway has also been implicated in 0 MeG repair [35]. 0 MeG
can be processed by post-replicative mismatch repair in E. coli in a double-stranded context,
but in a single-stranded context (gapped plasmid) the mutation frequencies in wt and mutS~
cells were found to be the same [36]. Using an M13 single-stranded system containing an
o
MeG lesion, Rye et al. showed that the Dam and MutH proteins did not impact the repair of
06 MeG by Ada or Ogt, but cells lacking MutS and MutL were less efficient at repairing 06 MeG
by Ada or Ogt [37]. The study also suggested that the MutS and MutL MMR proteins could aid
the repair of 06MeG by identifying and presenting the lesion in a manner that facilitated the
activity of methyltransferases on the lesion. It has also been postulated that MMR may play a
related but independent role in DNA alkylation damage signaling pathways [38].
4.1.5 Replication stalling elements
Factors that interfere with DNA replication can be grouped broadly into three categories:
genetic, intrinsic, and exogenous [39]. Genetic factors are mutations in genes that can affect
the rate of replication and accuracy (such as DNA pol Ill mutants). Intrinsic factors are natural
impediments to replication, such as DNA binding proteins, transcription units, unusual DNA
structures, and replication slow zones [39]. Exogenous factors affect DNA by either depleting
nucleotide pools or damaging the DNA, which may result in replication blocking lesions.
Work done in our laboratory has identified several lesions as strong blocks to replication. These
are listed in Table 4.2. 1-Methylguanine (m1G), 3-methylthymine (m3T), 3,N4-ethenocytosine
(EC), and tetrahydrofuran (THF) abasic site are particularly strong blocks to replication [40;41].
119
None of these lesions are repaired by Ada or Ogt, which is essential for this work as it will avoid
convolution of results. We chose THF and m3T for our experiment, as they are strong blocks to
replication even in wild-type cells.
THF is widely used as a synthetic analog of an abasic (apurinic or apyrimidinic; AP) site. AP
lesions can form in DNA spontaneously or as a result of DNA glycosylase action. These sites can
be dealt with by both translesion DNA synthesis and base excision repair (BER) machinery. A
THF moiety located in the DNA template strand is structurally similar to an AP site and shows
similar behavior as an AP site in in vitro reactions with exonuclease Ill, endonuclease IV, and
several DNA polymerases [42;43]. However, unlike a natural AP site, THF is not susceptible to
p-elimination, making it stable under the alkaline conditions used in solid-state synthesis of
oligonucleotides. The in vivo mutational profile of THF in a TXG nearest neighbor context in
single-stranded M13 is 55% T, 15% C, 12% A, and 13% G, while in a CXG context it is 65% T, 15%
C,4% A, and 16% G [44].
m3T has been found both in vitro [45-49] and in vivo [49;50] and is formed through the reaction
of DNA with SN2 alkylating agents such as MMS. This adduct is a very weak substrate for AIkB
and is a strong block to replication in vivo, which can be only slightly overcome by SOS bypass
polymerase induction [40]. Recently, the FTO enzyme has been shown to repair m3T efficiently
in single-stranded DNA but not in double-stranded DNA [51;52]. m3T is approximately 60%
mutagenic in SOS-/AlkB- cells, leading mostly to T
--
A (47%) transversions and T 4 C (9%)
transitions.
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4.2 Materials and methods
The REAP assay described here and outlined in Figure 4.3 has been modified from the work of
Delaney et al. [53]. Please refer to the original method for more detail and clarification.
4.2.1 Cell strains
All the E. coli strains used in this work contain the F' episome, which enables infection by M13
phage. GW5100 was used for large scale preparation of M13 phage DNA, SCS110 (JM110, end
Al) was used for amplification of progeny phage post-electroporation, and NR9050 was the
strain of choice for double agar overlay plating with X-gal for blue-white selection of plaques.
The E. coli strain used to test for 0 MeG repair was the C216 (ogt::kan) derivative of FC215,
used previously in our laboratory [24]. The repair background of the cells was tested and
confirmed in a previous study [37], and isfunctionally Ada+.
4.2.2 Oligonucleotides
All oligonucleotides, scaffolds, and primers were obtained from Integrated DNA Technologies
(IDT) unless specified otherwise. Sixteen-mer oligonucleotides of the sequence 5' GAAGACCRX
GGCGTCC 3' (R=G or A), where X is either the lesion of interest (06MeG) or a blocking lesion
(THF, m3T) were synthesized and purified as described [41]. Sixteen-mer oligonucleotides with
the same sequence but with X = G, A, T, or C were used as controls. An oligonucleotide of the
sequence 5' TCAACAGTTTCAGCGGAGTGAGAATAG 3' (Mspcomp) was used to create a locally
double-stranded region in M13mp7(L2) to aid cleavage by MspAll.
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The hairpin inserted at the proximal site (4 kb from the ori) was modeled after the existing one
at the distal site (6 kb from ori) in M13mp7(L2). This hairpin was designed to have the following
attributes: 1) A unique EcoRV site to linearize the DNA, 2) a Haelll site to enable analysis by the
REAP assay, 3) a minimal length to avoid loss from the phage genome over multiple rounds of
replication, 4) a sequence that enabled insertion into the M13 genome in only one orientation,
and 5) a sequence that would maintain the correct reading frame both with and without the
presence of the 16mer oligonucleotides. This last property was necessary because the hairpin
was being incorporated in gene Ill of M13 that encodes the minor coat protein, one necessary
for the progeny phage to be able to infect E. coli and hence survive. The hairpin fulfilling all of
these requirements was a 54-mer of the sequence 5'ACATACCGAAGGATATCCGTAGACTAGTCAT
CTACGGATATCCTTCGGGCCTCG
3'. The most probable structure of the hairpin is shown in
Figure 4.4.
The scaffolds used for holding the hairpin in place during ligation for parental vector
construction were 5' ACGGATATCCTTCGGTATGTCGGAGTGAGAATAGAAAGGAACA 3' (Msp LHS
scaffold) and 5' GCTAAACAACTTTCAACAGTTTCAGCGAGGCCCGAAGGATATCCG
3' (Msp RHS
Scaffold). For the main experiment, scaffold oligonucleotides were also used at the proximal
site (5' GGTCTTCATCCTTCGGTATGTCGGAGTG
3' and 5' TTTCAGCGAGGCCCGAAGGATGGACGC
3') and the distal site (5' GGTCTTCCACTGAATCATGGTCATAGC 3' and 5' AAAACGACGGCCAGTGA
ATTGGACGC 3') to hold the 16-mers in place to the cleaved single-stranded M13 vector during
genome construction, and do not overlap the lesion-bearing region.
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Sequencing primers used for confirming the insertion of the hairpin were 5' TTGCTACCCTCGTTC
CGAT 3' (M13EcoRV forward primer) and 5' TACTCAGGAGGTTTAGTACCGC 3' (M13EcoRV
reverse primer). The forward and reverse primers used for the mutagenicity (REAP) assay
spanned the M13 vector as well as the 5' end of the inserted oligonucleotide carrying the lesion
of interest, thereby effecting a selective amplification of DNA from the progeny phage resulting
from only the lesion-carrying genomes. The primers were modified with an aminoethoxyethyl
ether group (Y) at the 5' end to prevent labeling with
32P-y-ATP
in subsequent reactions. The
primers for the REAP assay at the proximal site were of the sequence 5' YTCACTCCGACATACCG
AAGGATGAAGAC 3' (REAP EcoRV forward primer) and 5' YAGTTTCAGCGAGGCCCGAAGGATGGA
CG 3' (REAP EcoRV reverse primer), while those for the distal site were of the sequence 5' YCAG
CTATGACCATGATTCAGTGGAAGAC 3' (REAP EcoRI forward primer) and 5' YTGTAAAACGACGGCC
AGTGAATTGGACG 3' (REAP EcoRl reverse primer).
4.2.3 Enzymes and chemicals
MspAll, EcoRV, EcoRI, Haelll, BbsI, HinFI, T4 DNA Ligase, T4 DNA polymerase, BSA, and the
enzyme reaction buffers were from New England Biolabs. Shrimp alkaline phosphatase (SAP)
was from Roche.
P1 nuclease, 5-bromo-4-chloro-3-indolyl- beta-D-galactopyranoside (X-gal),
isopropyl P-D-1-thiogalactopyranoside (IPTG), chloramphenicol, and kanamycin were from
Sigma Aldrich.
T4 Polynucleotide kinase was from Affymetrix. Sephadex G-50 Fine resin was
from Amersham Biosciences. Hydroxylapatite resin, 19:1 acrylamide:bisacrylamide solution,
and N,N,N',N'-tetra-methyl-ethylenediamine (TEMED) were from Bio-Rad. Phenol: chloroform:
123
isoamyl alcohol (25:24:1; pH 8) was from Invitrogen. 3P-y-ATP was from Perkin Elmer. Nonradioactive ATP was from GE Healthcare Lifesciences.
4.2.4 Double Agar overlay plaque method for phage analysis
The double agar overlay method used in this work was modified from Adams et al. [54]. This
method was used for enumerating initial electroporation events as well as phage titers to
ensure statistical robustness, but not for mutational analyses. Briefly, 10 ml of 2 x YT media
was inoculated with 2 ml of a saturated overnight culture of NR9050 and grown for 1 h at 37 0C
with aeration. Approximately 300 ul of this culture was mixed with 10 ul IPTG (24 mg/ml), 25 ul
of 1 % thiamine, and 40 ul X-Gal (40 mg/ml in DMF), and then added to 2.5 ml of Top agar
maintained in a molten state at 52 *C. Appropriate dilutions of supernatants containing phage
particles were immediately mixed with the top agar and poured onto B-broth plates, which
were shaken to evenly spread the agar. After a 10 min incubation at room temperature (to
allow the top agar to solidify), the plates were incubated overnight at 37 0Cto obtain dark blue,
light blue, or clear plaques.
4.2.5 M13 phage DNA
M13mp7(L2) phage single-stranded DNA was isolated as follows.
Various dilutions of a
previous stock of M13 phage supernatant were plated on a lawn of E. coli cells using the double
agar overlay method to obtain phage plaques. A well-isolated plaque was plugged using a
sterile Pasteur pipette and vortexed in 1 ml LB, 200 ul of which was used to make a starter
culture (grown overnight) by mixing with 10 ul of an overnight saturated culture of GW5100
124
cells in 10 ml LB. One milliliter of this phage starter culture was then used to inoculate GW5100
cells, which had been grown using 500 ul of an overnight saturated culture in 250 ml of fresh 2
x YT medium for 2 h at 37 *Cand shaken at 275 rpm. The inoculated culture was grown further
for 8 h at 37 *Cwith aeration, after which the cells were pelleted and discarded. The phage
were precipitated from the supernatant by addition of 4 % PEG 8000 MW and 0.5 M NaCl.
After overnight precipation at 4 0C,the phage were pelleted, resuspended in 5 ml TE pH 8, and
extracted with four washes of 3 ml 25:24:1 phenol:chloroform:isoamyl alcohol (Invitrogen, pH
8). The aqueous phase was passed through a 0.5 g hydroxylapatite column (BioRad), washed
with 5 ml TE, and eluted in 1 ml fractions with 12 ml of 0.16 M phosphate buffer. The DNAcontaining fractions were identified by spotting on an agarose plate containing ethidium
bromide. The phosphate buffer in those fractions was then exchanged for TE by three washes
in Microsep 100K spin dialysis columns (Pall Lifesciences). The DNA obtained was at a yield of
1 pmol/ml 2 x YT and was stored at -20 0Cuntil further use.
4.2.6 Construction of M13EH
Since M13mp7(L2) contains four MspAll sites in its genome, a complementary scaffold
(Mspcomp) was used to create a locally double-stranded region in order to cleave the DNA at
the target site. Ten pmol of M13 DNA were mixed with 20 pmol of Mspcomp and heated to 59
0C for
5 min followed by cooling to 0 *C @ 0.1 0C/s. The genome was then linearized by
incubating with 30 U of MspAll at 37 *Cfor 2 h. Twenty pmol each of Msp LHS and RHS
scaffolds were mixed with the linearized genome and annealed by heating at 54 0C for 5 min
followed by cooling to 0 0C@ 0.1 *C/s. In parallel, 20 pmol of the hairpin was 5' phosphorylated
125
by 15 U of T4 PNK in a reaction supplemented with 1 x T4 PNK buffer, 1 mM ATP, and 5 mM
DTT and incubated at 37 *C for 1 h. Finally, the phosphorylated hairpin and the linearized
genome were mixed and ligated for 8 h at 16 0Cin the presence of 1 mM ATP, 10 mM DTT, 25
ug/ml BSA, and 800 U T4 DNA Ligase. A cleanup digest was done by adding 100 x Mspcomp and
30 U MspAll and incubating at 37 *Cfor 2 h, so that any circularized genomes that did not
incorporate the hairpin would be linearized and removed in the subsequent purification steps.
The reaction volume was brought up to 120 ul with water and extracted once with 120 ul
25:24:1 phenol:chloroform:isoamyl alcohol. The aqueous phase was purified by three 2 ml
water washes in Centricon YM-100 spin dialysis columns (Millipore) to remove any residual
phenol and salts.
Recovery yields of 70% were obtained. Two controls were included in the
initial linearization step: one without the addition of the Mspcomp scaffold and the other
without the hairpin. The expectation was that these controls would not incorporate the hairpin
and could therefore be used as negative controls to check for insertion of the hairpin in the
modified genome.
Five pmol each of the modified M13 and the two controls were electroporated into 100 ul
electrocompetent C216, and the double-stranded RF DNA was isolated using the QlAprep Spin
miniprep kit from the cells following the 7 h growth in SCS110. The insertion of the hairpin was
checked by digestions with EcoRV and Haelll. Digestions were also performed with MspAll and
EcoRl to check for loss of the MspAll cognate site after insertion and retention of the original
EcoRl functionality in the original MCS, respectively. The presence of the hairpin was also
checked by sequencing with the M13EcoRV forward and reverse primers. As expected, the two
126
control genomes were negative for incorporation of the hairpin. A large-scale prep was
performed to isolate single-stranded DNA of the modified M13 (method described above),
hereon referred to as M13EH. The map of M13EH is shown in Figure 4.2b and its iconic
representation in Figure 4.2c. The complete sequence of M13EH is shown in Figure 4.5.
4.2.7 Construction of genomes
The two MCS in M13EH are designed to form hairpin structures, which contain a functional
EcoRl site at the distal site and an EcoRV site at the proximal site. Sixteen-mers containing
either a lesion or a control nucleotide were inserted at either site in a sequential fashion as
follows: 20 pmol of M13EH single-stranded DNA was linearized by incubation with 40 U of
EcoRV for 1 h at 37 0C. Scaffolds for the proximal site (25 pmol in 1 ul each) were annealed to
the ends of the linearized genome by incubation at 68 0Cfor 5 min followed by cooling to 0 *C
@ 0.1
0C/s.
In parallel, 30 pmol of the appropriate 16 mer oligonucleotides were 5'
phosphorylated by 15 U of T4 PNK, supplemented with 1x T4 PNK buffer, 1 mM ATP, and 5 mM
DTT and incubated at 37 0Cfor 1 h. The linearized genome was subsequently ligated with the
phosphorylated oligonucleotide for 8 h at 16 0C in a reaction containing 1 mM ATP, 10 mM DTT,
25 ug/ml BSA, and 800 U T4 DNA ligase. A cleanup digest was done by adding 40 U EcoRV and
incubating at 37 0C for 1 h, so that any circularized genomes that did not incorporate the 16mers would be linearized and removed in the subsequent purification steps. To degrade the
linearized genomes and excess scaffolds using the exonuclease activity of T4 DNA polymerase,
the enzyme was added to a final concentration of 0.25 U/ul and the mixture was incubated at
37 *Cfor 4 h. Finally, the reaction volume was brought up to 110 ul with water and extracted
127
once with 100 ul 25:24:1 phenol:chloroform:isoamyl alcohol. The aqueous phase was washed
three times with 2 ml water each in Centricon YM-100 spin dialysis columns (Millipore) to
remove any residual phenol and salts. Recovery yields of 50-60% were obtained.
These genomes were then processed in a similar fashion to incorporate 16-mers at the distal
site. Ten pmol of genome were linearized by incubation with 4 U of EcoRI for 8 h at 23 *C.
Scaffolds for the distal site (12.5 pmol each) were annealed to the ends of the linearized
genome by incubation at 50 *Cfor 5 min followed by cooling to 0 *C@ 0.1 *C/s. In parallel, 15
pmol of the appropriate 16-mer oligonucleotides were 5' phosphorylated by 7.5 U of T4 PNK,
supplemented with 1x T4 PNK buffer, 1 mM ATP, and 5 mM DTT and incubated at 37 *Cfor 1 h.
The linearized genome was subsequently ligated with the phosphorylated oligonucleotide for 8
h at 16 *C in a reaction containing 1 mM ATP, 10 mM DTT, 25 ug/ml BSA, and 400 U T4 DNA
Ligase. A cleanup digest was done by adding 4 U EcoRl and incubating at 23 *Cfor 8 h, followed
by a deactivation of the enzyme by heating to 65 0C for 20 min. This step was necessary to
avoid the star activity of EcoRl on single-stranded DNA upon subsequent incubation at 37 0C.
T4 DNA polymerase was then added to a final concentration of 0.25 U/ul and the mixture was
incubated at 37 0C for 4 h to degrade scaffolds. Finally, the reaction volume was brought up to
110 ul with water and extracted once with 100 ul 25:24:1 phenol:chloroform:isoamyl alcohol.
The aqueous phase was washed three times with 2 ml water each in Centricon YM-100 spin
dialysis columns (Millipore) to remove any residual phenol and salts. The genomes were then
incubated in a speed-vacuum for 1.5 h to reduce the volumes of the samples (but not
128
completely dried). An average of 3 pmol of genome was obtained for every 20 pmol of starting
M13EH single-stranded DNA.
4.2.8 Preparation of electrocompetent cells
Ten ml LB containing 10 ug/ml Kanamycin was inoculated with the C216 strain and grown
overnight at 37 *Cwith aeration. Three baffled flasks containing 150 ml LB medium each were
inoculated with 1.5 ml of a saturated overnight culture of the strain to be transformed. The
cultures were incubated at 37 0Cand shaken at 275 rpm for ~ 2.5 h until the cultures reached
early log phase, as measured by OD600 of ~0.5. The cell were then pelleted by centrifugation at
9500 rpm (Sorvall GSA rotor), resuspended in 1 ml cold sterile water, and pooled to a final
volume of 175 ml cold sterile water. This process of washing, pelleting, and resuspending was
repeated three times. The final resuspension was in 4.8 ml 10% glycerol to obtain a final
0
volume of 6 ml electrocompetent cells, which were then aliquoted and stored at -20 C prior to
use.
This process was used to prepare fresh electrocompetent cells for each experiment.
However, within each experiment, the same batch of electrocompetent cells was used for all
genomes in all replicates.
4.2.9 REAP assay
Approximately 0.5 pmol of genomes containing the lesions were electroporated in triplicate
into 100 ul competent cells in a 2 mm-gap cuvette using 2.5 kV and 125 Q. The cells were
immediately transferred to 10 ml LB and an aliquot of the freshly electroporated cells was
immediately plated using the agar overlay method to ensure that a minimum of 10s
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independent initial electroporation events occurred in 10 ml of culture. The cells were then
grown for 6 h at 37 0C with aeration to amplify progeny phage. The supernatants of the 6 h
cultures were retained and plated using the agar overlay method to confirm 104 -fold
amplification in the progeny phage titer. Another round of amplification was performed in
order to ensure that the progeny phage being analyzed came from genomes that entered the E.
coli cells, and not the residual genomes that did not get electroporated into cells but were still
present in the milieu, since PCR is subsequently used. This was done by infecting 10 ul of an
overnight culture of SCS110 cells with 100 ul of the 6 h supernatants in 10 ml LB and incubating
for 7 h at 37 *Cwith aeration, after which the supernatant was retained. Single-stranded M13
progeny phage DNA was isolated from 0.7 ml of supernatant using a QlAprep Spin M13 Kit with
final DNA suspension in 100 ul elution buffer. PCR was performed at the proximal and distal
sites individually using the appropriate REAP forward and reverse primers.
The region of
interest was amplified from 10 ul per QlAprep elution sample in a total volume of 25 ul using
1.25U Pfu Turbo DNA polymerase, 25 mM of each dNTP, and 10 x Pfu Turbo buffer. The PCR
0
program started by denaturing at 94 'C for 5 min, then cycled 30 times at 94 C for 30 s, 67 *C
for 1 min, and 72 0C for 1 min, and finally extended for 5 min at 72 0C . The volume was then
made up to 110 ul using water and extracted once using 25:24:1 phenol: chloroform: isoamyl
alcohol to destroy the DNA polymerase (including the exonuclease domain).
The aqueous
phase was passed through a Sephadex G-50 Fine resin spin column to remove any remaining
dNTPs and traces of phenol.
130
The purified PCR product was then treated with Bbsl (1.5 U for 4 ul of sample in a total volume
0
of 6 ul) and shrimp alkaline phosphatase (0.3 U) by incubating at 37 *Cfor 4 h, heating to 80 C
for 5 min, and cooling to 20 0C@ 0.2 *C/s. The 5' ends were then labeled in a total volume of 8
ul with a mixture of non-radioactive ATP (20 pmol),
P-y-ATP (1.66 pmol of 10 uCi/ul at 6000
32
Ci/mmol), and 5 U of Optikinase/T4 PNK by incubation at 37 *Cfor 15 min, followed by 65 *Cfor
20 min and cooling to 23 "C@ 0.1 *C/s. The labeled product was then trimmed by Haelll (10 U
in a final volume of 10 ul) at 37 0Cfor 2 h, followed by addition of 10 ul 2 x formamide loading
dye, which quenched the reaction. The samples were then loaded onto a 20 % denaturing gel
and run for ~3.5 h at 550 V, until the xylene cyanol dye migrated 10.5 cm.
Following
electrophoresis, the 18-mer bands were excised from the gel, and crushed and soaked
overnight in 200 ul water. After desalting with Sephadex G-50 Fine resin spin columns, the
samples were lyophilized overnight to dryness, resuspended in 5 ul containing 1 ug P1 Nuclease
0
in 30 mM sodium acetate and 100 mM zinc chloride, and incubated at 50 Cfor 1 h. One ul of
each sample was then spotted onto PEI-TLC plates and separated using 200 ml of a saturated
solution of (NH4) 2HPO 4 adjusted to pH 5.8. After 12 h of development, the TLC plates were airdried and quantified using phosphoimagery.
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4.3 Results
4.3.1 GATC controls
The REAP mutation-analysis assay was initially designed around the distal insertion site in M13.
Therefore, a control experiment (Figure 4.1a) was done to test that the REAP assay works as
expected at both the proximal and distal sites in the modified version of M13, i.e., M13EH. An
approximately equimolar mixture of genomes containing a 'G', 'A', 'T', or 'C' insertion at the
distal site (but a pure 'G' at the proximal site) were mixed, electroporated into E. coli cells, and
processed via the REAP assay. On interrogating the distal insertion site, an approximately 25%
output of each of the four bases was obtained, while ~100% G was detected at the proximal
site. Similar results were obtained for the equimolar mixture of genomes with the four bases
inserted (individually) at the proximal site but with a pure 'G' at the distal site. These results
are shown in Figure 4.6. Several conclusions can be drawn from this experiment: (a) The REAP
assay is effective in determining the base composition at both the proximal and distal sites, (b)
a 3 - 4 % basal level of 'background' mutation frequency (mostly A) is detected at the newly
incorporated proximal site in M13EH, and (c) there is little, if any, recombination taking place
between the proximal and distal sites. The background mutation frequency at the proximal site
is detected in all experiments at the same level and may be investigated further. One caveat to
keep in mind isthat no lesions were used in this test run.
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4.3.2 Test of recombination between the proximal and distal sites
The aim of this experiment (Figure 4.1b) was two-fold. While the control experiment described
above confirmed a working system, it remained to be seen whether this system would work
well when lesions, instead of unmodified nucleotides, were inserted into the genomes. The
second aim was to test whether recombination between the two sites occurred in the presence
of a lesion, which if present, would convolute the results. To this end, two genomes containing
a pure 'T' at the proximal site and either O6MeG or a pure 'G' at the distal site were
electroporated individually into cells and taken through the REAP assay. The primary mutation
caused by 06MeG is a G->A transition. Hence, the expected base composition at the distal site
would contain either pure G, or a mixture of G and A. Since we were also testing for possible
recombination between the two sites, we chose to insert a thymine base at the proximal site
which would not result in a G or Awhen processed by the REAP assay.
The results, depicted in Figure 4.7, show that (a) The REAP assay works as expected when a
lesion is introduced at the distal site as evidenced by the typical 40% 'A' mutation profile
obtained for O6MeG [24], and (b) no recombination takes place between the proximal and
distal sites even in the presence of a lesion, as evidenced by no thymine 'contamination' in the
mutational profile of the distal site. The last observation is particularly important as the cell
strains used in this work are not recA-, the two sites of insertion are similar in structure, and the
lesions are always inserted in the same 16-mer oligonucleotide sequence context at either site.
Hence, there is a possibility of the mutations resulting from the lesion inserted at the proximal
site being 'rescued' by homologous recombination with the pristine distal site (and vice versa).
133
However, our results from this experiment and the previous one show that said recombination
does not take place. Note that the basal low level of 'A' at the proximal site is also seen in this
set of experiments.
In hindsight, a third genome carrying 06MeG at the proximal site and a pure 'T' at the distal site
should have been included in this experiment for completion. However, future experiments
included genomes with lesions at either site and the results were in line with our findings
above. These experiments are discussed below.
4.3.3 Effect of distance from the origin of replication on the mutation frequency of O6 MeG
Our initial hypothesis was that the farther a lesion is placed from the origin of replication the
more time it would have for repair, which should translate into a lower mutation frequency.
To test this hypothesis, we engineered M13 to include a site of insertion proximal to the ori for
comparison with the distal site. We then created several genomes to include all combinations
of O6MeG and a pure 'G' at either site.
Additionally, these lesions were inserted in two
different nearest-neighbor sequence contexts, GXG and AXG, as per a previous study which
showed relatively strong and weak repair, respectively, of O6 MeG by Ada [24]. The various
genomes were individually electroporated into Ada+ Ogt cells and taken through the REAP
assay. Figures 4.8, 4.9, and 4.10, and Tables 4.3 and 4.4 show the results of this experiment.
Since the primary mutation caused by O6MeG is a G-A mutation with negligible amounts of T
and C,the mutation frequency of this lesion can be wholly represented by the percentage of A
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obtained in the REAP assay.
This property of O6 MeG eliminates the need to show the
percentages of all bases in the mutation frequency graphs, thus simplifying the depiction of
data. This concept is represented in Figure 4.8 and is used in Figures 4.9 and 4.10.
Figure 4.9a depicts the mutation frequency as a function of distance from the ori. In a GXG
context, a lesion placed at the distal site shows a ~5% higher mutation frequency than the same
lesion placed at the proximal site.
This finding is in contrast with our initial hypothesis
according to which a distal site would have more time for repair and hence would show a lower
mutation frequency. This result is significant (p value = 0.004) in at least one case, where both
the lesions are present on the same genome. We do notice that the basal mutation frequency
at the proximal site shows up once again. If we were to 'normalize' the mutation frequency at
the proximal site by deducting this basal level of mutation frequency, we would get a difference
of ~8% in the mutation frequency between the two sites. Also, this result is significant both
when the two lesions at the two sites are present on different genomes (p value = 0.05) as well
as when they are present on the same genome (p value = 0.001). This data is not shown in the
graph. One thing to note is that the statistical significance of results in the GXG context is
dependent on the fact that an extremely small standard deviation was obtained for one of the
data sets, namely, the mutation frequency at the distal site in a two-lesion genome. It is
possible that the small error obtained for this particular result is a statistical outlier; further
statistical analyses will be needed to determine whether this is the case. However, while this
observation may call into question the statistical significance of the results, it does not negate
the trends seen in our results described in this section and the next.
135
Figure 4.9b shows the same data in an AXG context. The first observation is that the mutation
frequencies across the board are higher than those obtained in the GXG context. This has been
observed in a previous nearest-neighbor sequence context study from our laboratory [24] and
is therefore as expected. Interestingly, the basal mutation frequency at the proximal site is
higher (~5.5 %) compared to that observed in the GXG context (~2.5%).
The same trend of
mutation frequency with distance from the ori is observed in the AXG sequence context.
However, the difference in the mutation frequencies at the two sites is only ~1-3%, and is not
statistically significant.
If we 'correct' the proximal site mutation frequencies for the basal
level, the difference jumps to 6-8%, which is statistically significant when the two lesions at the
two sites are on the same genome (p value = 0.008, not shown in the graph).
4.3.4 Effect of genome lesion load on the mutation frequency of 0 MeG
In analyzing the data from the above experiments, an interesting effect was observed. A
genome carrying two lesions showed a lower mutation frequency at either site, compared to
the single-lesion-carrying genomes. These results are shown in Figure 4.10a (GXG context) and
Figure 4.10b (AXG context).
At either site, the mutation frequency in atwo-lesion genome is lower by 5-6% in GXG and 4-5%
in AXG sequence context as compared to genomes that carry a single lesion. However, this
finding is statistically significant only in the GXG context (p value = 0.02).
In this case,
normalizing the proximal site data by subtracting the basal level of 'mutation frequency' would
136
not change the relative difference in mutation frequencies between the two-lesion and onelesion genomes at the proximal sites.
4.3.5 Effect of a blocking lesion at the proximal site on the mutation frequency of 0 MeG at
the distal site
Another method to test our hypothesis would be to stall the replication machinery by placing a
blocking lesion in its path, thus giving the lesion at the distal site more time for repair. Also, the
ideal lesion would be one which was not acted on by either Ada or Ogt so that it would not
interfere in the kinetics of O6 MeG repair. The blocking lesion should also not be acted upon by
any other cellular repair elements. Several lesions fit the bill and are listed in Table 4.2. We
chose two of the strongest, simplest in structure, replication-blocking lesions, m3T and THF, as
determined by the CRAB assay in previous studies (7%and 3% bypass efficiencies respectively).
These lesions were inserted at the proximal position in genomes that had 0 MeG inserted at
the distal position. The genomes were electroporated individually into cells and the mutation
frequency of O6 MeG was measured via the REAP assay. Since m3T is a thymine lesion, and THF
usually pairs with an adenine (hence behaving like a thymine), we used a genome containing a
non-blocking pure 'T' at the proximal position (but O6MeG at the distal position) as a control.
For comparison, we also included a control genome containing 06MeG at both sites.
The results obtained were in sharp contrast to those that would be expected if our intial
hypothesis were correct. We observed that placing a strong block to replication at the proximal
site increased the mutation frequency of O6 MeG at the distal site. These findings are shown in
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Figure 4.11a. The presence of m3T increased the mutation frequency of O6 MeG from 50% to
71% (p value = 0.001) while THF increased the mutation frequency of O6 MeG from 50% to 67%
(p value = 6 x 10-6). One interesting thing to note is that the genomes carrying two lesions (one
o
MeG and one blocking lesion) did not show a lower 06 MeG mutation frequency as was
observed in our previous experiment (Section 4.3.4).
That finding seems to be limited to
genomes carrying two 0 MeG lesions. We also performed the REAP assay at the proximal site
to confirm the identity of the lesion inserted there. The results were as expected and this data
is shown in Figure 4.11b.
4.4 Discussion and future work
Our initial hypothesis was that the farther a lesion is placed from the origin of replication, the
more time it would have for repair before the replication machinery encountered it, resulting in
a lower mutation frequency. This possible result was also the expectation if we were to slow
down the replication polymerase by placing a replication-blocking obstacle upstream of the
lesion. Our results disprove this hypothesis and show some interesting trends. We found that
the lesion mutation frequency increases, rather than decreases, with distance from the ori. The
increase was found to be approximately 5 - 8 % between two sites placed 2 kb apart. It was
also noticed that a genome carrying a higher O6MeG lesion load had a lower mutation
frequency at either interrogation site compared to genomes containing a single lesion at either
site.
Finally, when we placed a replication-blocking lesion upstream, in the path of the
138
polymerase, we obtained strikingly high mutation frequencies for 06MeG. Our results call for a
new hypothesis, of which several are discussed below.
The result that mutation frequency increases with distance from the ori is in line with studies
done on a genomic scale, which found a 2x lower mutation frequency closer to the origin [4;5].
However, post-replication repair, especially by homologous recombination, was thought to play
a role in keeping the mutation frequency near the ori low. We eliminated that possibility in our
system in two ways. Firstly, we used a single-stranded M13 genome to house the lesions and
homologous recombination is known to occur in double-stranded DNA. The mutations get 'set'
in the genome during the formation of RF, if they are not repaired by direct demethylation.
Secondly, we tested our system for any occurrence of homologous recombination between the
two sites and found none.
While it is not known exactly how Ada searches for and locates a repairable lesion, there are
three proposed mechanisms [55]. The first envisions a model in which Ada actively scans the
DNA, rotating out each base to check for damage. The second mechanism is one whereby Ada
does not scan the DNA but rather directly detects a lesion hidden in the intrahelical structure of
DNA by the distortion it causes by its presence. The third mechanism takes this further by
proposing that the lesion rotates out on its own and is then detected by Ada. It is a reasonable
expectation that an ATPase domain would be required for Ada to migrate linearly along the
genome. However, structural studies have not identified such a domain in Ada. What is known
139
is that Ada has a helix-turn-helix DNA binding domain as well as an 'arginine finger' that helps it
rotate out the damaged base for repair in an extrahelical fashion [56].
One theory that we believe could best explain our observations is the preferential binding of
Ada at or near the ori every time it starts to scan the DNA for potential alkylation damage.
Once bound, it would move along the DNA until it came across an 0 6 -alkylated base, a blocking
lesion, or a stalled replication polymerase. In the first case, Ada would repair the lesion in a
suicidal fashion, which would result in the recruitment of a new molecule of Ada at the ori to
resume the scan of the remaining length of the genome. In the second and third scenarios Ada
would 'fall off' the DNA and resume its scan at the ori in an iterative fashion until a scan of the
entire stretch of DNA is accomplished. This hypothesis depicts a scenario that would explain
why we see a lower mutation frequency at the proximal site and a higher one at the distal site.
It would also explain the higher mutation frequency obtained at the distal site when a
replication-blocking lesion is placed at the proximal site. There are several ways by which we
could test this theory: (1) modifying the M13 genome to reverse the orientation of the ori, (2)
repeating the experiments in an Ada-Ogt+ cell strain, and (3) using our system to analyze the
mutation frequency of other nonalkylated lesions.
While (1) would test this theory in a
definitive manner, (2) and (3) might show that other repair proteins repair other lesions by an
approach similar to the one proposed above. We should keep in mind that the exact timelines
of repair and replication starts in the cell are not known. When does replication start once the
phage DNA enters the cell? How long does it take to traverse the distance between the two
sites? How long does it take for the initial RNA primer to be formed? These are pertinent
140
questions, the answers to all of which are not yet known. DNA polymerase Ill can replicate the
genome at the rate of 1000 nt/s and can therefore traverse the entire length of the M13
genome in less than 10s. However, the initiation of replication requires the assembly of several
proteins and the formation of an 18-20nt long RNA primer by E. coli RNA polymerase [57;58],
which has a replication rate (12nt/s [59] ) considerably slower than that of DNA polymerase Ill.
It is quite possible that the repair proteins function on a time scale that is much faster than the
time required for initiation and completion of replication.
In our scenario, the higher lesion load possibly stimulates a stronger repair response in the
cells. The cells used in our experiments contain only a few molecules of Ada and none of Ogt
(since
Ogt- cells were used).
The adaptive response requires the
presence of
methylphosphotriesters for induction. Since we are not exposing the cells to any alkylating
agents and are instead introducing a single lesion in a site-specific manner, the adaptive
response is not expected to be induced. A plausible hypothesis is that the role of lesion
'beacon' is played by alkyltransferase-like (ATL) proteins. These proteins have a high binding
affinity for 06MeG but no repair capabilities and are known to exist in various species [60]. It
has been shown that while they bind O MeG tightly, repair by alkyltransferases is possible
eventually via a competitive displacement of ATL from the substrate [61]. They can thus serve
as a 'lightning rod' to attract repair proteins. In fact, recent studies have shown that the E. coli
ATL protein may have a role in protecting 06-alkylG lesions (but not O6 MeG) from MMR
processing [62] and redirecting the repair to NER [63;64] by pairing with UvrA, much like Mfd.
However, we do not expect NER to play a role in our single-stranded system.
141
Another favored possibility is that the mutation frequencies we observe are the result of some
long-range sequence context effects. While the lesions at both the proximal and distal sites are
housed in the same 16-mer oligonucleotides, there may be sequence effects at play beyond
that distance. For example, in a study by Parris et al. the introduction of a mutation at one site
in the supF tRNA led to creation of a hotspot 8 bases upstream, while a mutation at another site
led to the suppression of a hotspot 48 bases upstream [65]. This effect has been shown to exist
over longer distances, and in the downstream direction as well, as was shown by Levy et al.
[66]. Both studies, taken together, showed by the process of elimination that this 'distance'
effect was not a consequence of differential formation of lesions, differential repair of lesions,
or the positioning of the lesions in the leading or lagging strand. Thus it was concluded that the
long-range effect seen was somehow a function of the sequence context of the lesions. While
this effect remains unexplained, and no such studies exist specifically for O6 MeG, we cannot
negate the possibility of a similar phenomenon occurring in our system. If this happens to be
the case, our study would be the first to report a long-range sequence effect on the order of 2
kb, which is the distance between the proximal and distal sites. One method to test whether
such an effect is at play in our system would be to change the relative positions of the proximal
and distal sites with respect to the ori. This can be done by (1) reversing the orientation of the
ori, or (2) moving the ori such that the distal site is encountered first in the direction of
replication.
Yet another theory which we considered was that the lesions were titrating the miniscule
amount of Ada present in the cells. The replication machinery would encounter the proximal
142
lesion first and possibly attract Ada to that site. However, with the number of Ada molecules
per cell in single digits, it is possible that not enough would remain to repair the distal lesion.
We refer to this as the 'Cowcatcher' model, a term originally proposed by Lawrence Grossman
(Lawrence Grossman to John Essigmann, personal communication). This theory would explain
the lower mutation frequency seen at the proximal site and the higher mutation frequency at
the distal site only in genomes carrying two 06MeG lesions, and that too if Ada were scanning
the genomes in a processive rather than distributive manner. This theory would also fail to
explain the overall lower mutation frequency seen in genomes carrying a higher 06 MeG lesion
load, compared to single-lesion-carrying genomes.
A further possibility is that the proximal site is under a tighter repair pressure. As it is a part of
an important M13 gene required for infection of the host, transcription-coupled repair (TCR)
could lead to a lower mutation frequency. The distal site, on the other hand, is located in a
non-essential (for the virus) lacZ gene in the MCS, which is an artificial insertion in the M13
genome. TCR is known to occur when RNA polymerase is stalled at a lesion. The transcription
repair coupling factor Mfd binds to and displaces the RNA polymerase to enable the
recruitment of UvrA at the site. The subsequent steps involve lesion verification by UvrB, DNA
strand dual incision by UvrC, excision by UvrD, repair by DNA polymerase I, and ligation (see
[67] for a review). As mentioned earlier, the expectation is that the lesion is either repaired or
results in a mutation the very first time it is processed by a polymerase, namely, during the
formation of the double-stranded RF phage. In the latter scenario the lesion would exist in a
double-stranded mispaired state in the phage genome. However, even if transcription of viral
143
genes is initiated by E. coli RNA polymerase at this stage (which is believed to be the case), we
do not expect TCR to play a major role in lesion repair. The reason is three-fold. Firstly, and of
paramount importance, it is the plus-strand of single-stranded M13 that carries the lesion.
When it is converted to the RF form, the minus-strand serves as the template DNA for
transcription. This strand will have either the correct base or a mispaired base opposite the
lesion, but no transcription-stalling moiety. The second reason, while less important, but
nevertheless relevant to the thought process, is that E. coli RNA polymerase is known to be
blocked by only bulky lesions, and seems to bypass smaller lesions such as 8-oxo-G and abasic
sites with ease [68]. In fact, it has been shown that E. coli RNA polymerase can bypass 06 MeG
in vitro in an error-prone fashion [69]. Lastly, a previous study from our laboratory found that
the mutation frequencies of 0 MeG at the distal site in ada-ogt-uvrB+ cells and ada-ogt-uvrBcells were very similar [24]. While it has been speculated that TCR on the non-transcribed
strand is a possibility, it hasn't been explicitly tested [67]. We could test whether TCR (by
involvement of UvrABC) occurs across 06 MeG by using ada+ogt-uvrB+ cells, and comparing it
with the mutation frequency obtained in Ada proficient but uvrB- cells, both in single-stranded
as well as double-stranded vector systems. If it turns out that TCR was still occurring in our
system, it would be a variable that would be hard to get around, as M13 has very few intergenic
sites available for introducing new genetic elements, which would not be under the influence of
TCR, without disrupting phage viability.
MMR could also be playing a role in repair of O6 MeG at either site. It takes two rounds of
6
replication for O6MeG to result in a G->A transition. In the first round, O MeG mispairs with a
144
thymine, which in turn pairs with an adenine in the second round of replication. MMR can kick
in after the first round when the O6 MeG:T mispair is detected. However, it is believed that the
repair machinery would identify the 06MeG-carrying strand as the template, excise the
thymine, and reinsert another thymine opposite the lesion, thereby initiating a series of 'futile'
cycles of repair [70]. If somehow MMR correctly identifies and excises 06MeG from the RF
form, a mutation will have been 'set' into the pairing strand leading to a mutational outcome
which would not reflect the effect of MMR or of Ada.
What if there exists more than one origin of replication in M13? If this were true, the ori would
have to be closer to what we consider the distal site in our system in order to be consistent
with the initial hypothesis. Literature suggests that M13 contains just one origin of replication
[71]. There is one study in which the ori sequence was deleted, and while the formation of
plaques was severely affected, plaques still occurred [72], hinting at the possibility of a second
ori. However, in wild-type M13, it is expected that the known ori is dominant for replication
starts. It would be relatively easy to test for the presence of a second ori by creating a version
of M13EH with the known ori modified for loss of function.
The prospect of processive translesion synthesis across O6 MeG was also analyzed.
It is
imaginable that one of the TLS polymerases that helps bypass the blocking lesions carries on
replicating the template instead of 'falling off, thereby performing error-prone replication at
the distal O6MeG.
It has been shown that if single-stranded M13 is for some reason not
converted to the double-stranded RF form (say, due to the deletion of the ori), it can induce the
145
SOS response in E. coli [73]. We can imagine that a replication-blocking lesion that impedes the
formation of RF might do the same.
If this hypothesis were true, which TLS polymerase
performs this 'bypass' of the blocking lesions and subsequently of 06MeG? There are three TLS
polymerases in E. coli, all of which bypass THF in vitro. Of those, pol IV almost exclusively
results in a -2 deletion when it encounters a THF base, something not seen in our experiments.
In vivo, Pol 11and pol V have a preference of incorporating adenine opposite THF [44], which is
congruent with our data. In addition, pol V seems to be the most important polymerase for
THF bypass when the SOS system is not induced [44]. While no studies exist to pinpoint the TLS
polymerase for m3T, the fact that the SOS polymerases play a small but significant role in the
bypass of m3T has been seen in previous work by Delaney and Essigmann [40]. However,
replicative pol III could be also performing error-prone bypass across this lesion in E. coli. It has
been shown in vitro that pol III can bypass the O6 MeG lesion placed in single-stranded M13 with
about the same efficiency as the non-lesion control [74]. The same study showed that pot I and
pol 11did not mediate any TLS across 06 MeG. Other studies with related TLS polymerases show
that Pol P is unlikely to play an important role in 0 MeG bypass [75], while Dpo4, a Y-family
polymerase present in Sulfolobus solfataricus, can perform TLS across this lesion [76], which
hints at the possibility of the related E.coli enzyme pol IV doing the same. Yeast and human pol
il are known to have this capability as well [77]. All these studies taken together imply that
0 MeG is 'bypassed' most likely by either pol V or by the replicative pol Ill itself. Even if pol V
replaces pol IlIl at the proximal site and decides to hang on, it seems to have too low a
processivity compared to pol IllI in order to be able to replicate the 2kb between the proximal
and distal sites in a timely manner. There are ~10 molecules of pol Ill per E. coli cell, with a
146
processivity of 105 and a replication rate of ~700 nucleotides / second [6]. In contrast, pol V has
a processivity of ~18 nucleotides in conjunction with the p-clamp [78]. We could also speculate
that the TLS polymerase recruited at the proximal site hangs on to the p-clamp as the
replicative polymerase resumes replication and reaches the distal site (without physically
traversing along the DNA itself and thus obviating the need for processivity), where it leads to a
higher mutation frequency.
If this were true, it could result in non-random clusters of
mutations, or 'mutation showers', in other regions of the M13 genome as well [79;80] . It is
also possible that all TLS polymerases of E. coli have a propensity to insert a 'T' opposite an
o
MeG lesion, regardless of which one is recruited to 'bypass' the lesions. A simple way to
check whether this hypothesis of processive TLS is true would be to repeat the experiment with
the positions of the blocking lesions and 06MeG exchanged. We could also check for the
presence of 'mutation showers' by sequencing the progeny phage population.
Further experiments can be done to increase the time available for repair of an 06MeG lesion
using options (d) and (e) mention in the introduction. These options were: (d) to use kinetically
slow DNA polymerase Ill mutants and Ada mutants, and (e) to modulate the level of expression
of Ada and Ogt proteins. The a-subunit of the enzyme responsible for the polymerization
activity of pol Ill is encoded by dnaE. Several temperature-sensitive dnaE mutants of pol IlIl
have been reported, some of which have slower growth and slower rate of DNA synthesis [8184]. Similarly, kinetic mutants of Ada that repair 06MeG at slower rates as compared to wt
have been reported in the literature. 'Ada3' and 'Ada5' transferases have been shown to repair
0 -MeG in DNA 20 and 3000 times slower than wt Ada. These proteins also display deficient
147
induction of the ada and alkA genes but exhibit normal DNA phosphotriester repair [85;86].
Lastly, the level of gene expression of Ada and Ogt can be controlled by replacing the native
promoters. While Ada can be induced by the adaptive response on exposure to methylating
agents [25;27], the number of Ada molecules is estimated to rise from 1-2 molecules in an
unadapted state to ~3000 molecules in a fully adapted cell. Control of Ogt expression will
require genetic modification as it is constitutively expressed in the cell. A fine control of genetic
expression of Ada and Ogt can be achieved using the promoter system developed by the
Keasling group, which was shown to direct a homogenous induction for the entire population of
cells based on the concentration of the inducer [87;88]. These experimental systems will
require extensive manipulation of the E. coli genomic DNA before they can be used to further
decipher the effect of distance from the ori on the mutation frequency of a lesion.
This exploration of the effect of distance from the origin of replication on the lesion mutation
frequency has resulted in some interesting and exciting data that point to molecular
mechanisms not yet fully understood. We have performed some thought analyses outlining
what might possibly be going on in the intracellular milieu. Further investigation (by way of
some of the aforementioned experiments) may clarify the chronological and hierarchical roles
of the various players in the repair of and replication across DNA alkylation damage.
148
4.5 Figures
Figure 4.1: Experimental Outline.
(a)GATC controls for distal site(top row) and proximal site (bottom row)
1. Make an
Equimolar
Mixture
0
Ada
Q&
REAP
k\~J 2. Electroporate
Into E.coli
K\J
4
kG C
ti
I
1. Make an
Equimolar
Mixture
Mutation
Frequency
Ada
99t
Assay
2. Electroporate
Into E.coli
(b) Controls to check for recombination between the distal and proximal sites
D
N j
j
Electroporate
Indcividuall
REAP Assay
into E.coli
Lesion
Mutation
Frequency
(c) Main experiment genomes in GXG (top row) and AXG (bottom row) sequence contexts
Electroporate
Individually
into E.coli
Ada
Qo
REAP Assay
Lesion
Mutation
Frequency
REAP Assay
Lesion
Mutation
Frequency
DOElectroporate
Individually
into E.coli
+1
-
(d) 'Resistor' experiment genomes with blocking lesions placed at the proximal site
"" \
'""'"\
*'"'
GXG sequence context
Electroporate
Individually
into E.coli
AXG sequence context
Aa
:+
t
REAP Assay
Lesion
Mutation
Frequency
O'MeG
149
Figure 4.2: (a) Map of M13mp7(L2) showing the location of all genes and the unique EcoRI site in the
lacZ gene used for site-specific mutagenesis. The EcoRI site is at a distance of ~6 kb from the origin of
replication (ori). (b) The proposed M13EH structure includes an additional site of insertion with a
unique EcoRV site at ~4 kb from the ori. (c)The M13EH genome, with lesions (shown as black 'lollipops')
inserted at both the sites of insertion, depicted as an icon. The EcoRI site is referred to as the distal site
while the EcoRV site is referred to as the proximal site, reflecting the distances from the ori.
(a)
R4
206
1304
"30
EcoRI
2853
28!-6
4242
3191
3106
-2kb
EcoRV
-4kb
(c)
Distal
site
4I
Proximal
site
150
Figure 4.3: REAP mutagenesis assay. Adapted from [53].
GAAGACCTGGCGTCC-
Lesion
Genome
Cells
,I-GAAGACCTAGGCGTCC-
,
GAAGACCTGGGCGTCG.
Unrepaired
Repaired
Genome
Genome
20%
80%
)
PCR amplify with primers
designed to amplify only
region of interest
13AACCTAGGCGTC
Unrepaired
\
Genome
Repaired
(GAAGCCTGGGCTC
Genome
80%
20%
61 mer
oL
Itj
Cut with Bbsl & dephosphorylate
"P()-LabeI new 5' ends & HaellI trim
Separate products via PAGE
NH,
HN-
BbsI t*
34mer
H2N
a
30mer
HaeI|i
14mer
I
18mer
13mer
X
NH,
13mer
REAP
34mer
4'
Excise oligo whose 32P-labeled
5' base was the lesion site
Digest with Nuclease P1
Resolve 5' dNMPs via TLC
Quantify via PhosphorImagery
18mer
Quantify frameshifts,
) if applicable
5'dCMP
...........
Mutation =
S
5' dTMP
S
5' dGMP
5' dAMP
151
Figure 4.4: Structure of hairpin inserted in the modified M13 vector. The 'OligoAnalyzer 3.1'
software by Integrated DNA Technologies was used to obtain this hairpin structure, which is the
hairpin that is most likely to be formed according to energy minimization calculations. The
unique EcoRV site is shown as well.
T
G
C
II
C
30
G C
I
C
G
IC
I
I
GC
I
I
EcoRV
11
4
site
GIO
IC
40
I
I'
C 9G
G COG
9C
CO
\
I
I
I
I
ROT
I' I
G C
TT
I
I
c OG
I
I'
5
G
G
A IC
T
C
I
AI
C
T
%
3.
152
,
Figure 4.5: Complete sequence of M13EH. The
gene, M sites,
and EcoRV sites sequences are highlighted below.
gene 11,
,
5,
GTTCCAAACTGGAACAACACTCAACCCTATCTCGGGCTATTCTTTTGATTTATAAGGGATTTTGCCGATTTCGGAAC
CACCATCAAACAGGATTTTCGCCTGCTGGGGCAAACCAGCGTGGACCGCTTGCTGCAACTCTCTCAGGGCCAGGCGG
TGAAGGGCAATCAGCTGTTGCCCGTCTCGCTGGTGAAAAGAAAAACCACCCTGGCGCCCAATACGCAAACCGCCTCT
CCCCGCGCGTTGGCCGATTCATTAATGCAGCTGGCACGACAGGTTTCCCGACTGGAAAGCGGGCAGTGAGCGCAACG
CAATTAATGTGAGTTAGCTCACTCATTAGGCACCCCAGGCTTTACACTTTATGCTTCCGGCTCGTATGTTGTGTGGA
ATTGTGAGCGGATAACAATTT-CACACAGGAAACAGCT
MMtkAAATGA
GCTGATTTAACAAAAATTTAACGCGAATTTTAAC
GCTTTTCTGATTATCAACCGGGGTACATATGATTGACATGCTAGTTTTACGATTACCGTTCATC
GATTCTCTTGTTTGCTCCAGACTCTCAGGCAATGACCTGATAGCCTTTGTAGATCTCTCAAAAATAGCTACCCTCTC
CGGCATTAATTTATCAGCTAGAACGGTTGAATATCATATTGATGGTGATTTGACTGTCTCCGGCCTTTCTCACCCTT
TTGAATCTTTACCTACACATTACTCAGGCATTGCATTTAAAATATATGAGGGTTCTAAAAATTTTTATCCTTGCGTT
GAAATAAAGGCTTCTCCCGCAAAAGTATTACAGGGTCATAATGTTTTTGGTACAACCGATTTAGCTTTATGCTCTGA
GGCTTTATTGCTTAATTTTGCTAATTCTTTGCCTTGCCTGTATGATTTATTGGATGTTAATGCTACTACTATTAGTA
GAATTGATGCCACCTTTTCAGCTCGCGCCCCAAATGAAAATATAGCTAAACAGGTTATTGACOATTTGCGAAATGTA
TCTAATGGTCAAACTAAATCTACTCGTTCGCAGAATTGGGAATCAACTGTTACATGGAATGAAACTTCCAGACACCG
TACTTTAGTTGCATATTTAAAACATGTTGAGCTACAGCACCAGATTCAGCAATTAAGCTCTAAGCOATCCGCAAAAA
TGACCTCTTATCAAAAGGAGCAATTAAAGGTAOTCTCTAATCCTGACCTGTTGGAGTTTGCTTCCGGTCTGGTTCGC
TTTGAAGCTCGAATTAAAACGCGATATTTGAAGTCTTTCGGGCTTCCTCTTAATCTTTTTGATGCAATCCGCTTTGC
TTCTGACTATAATAGTCAGGGTAAAGACCTGIXTTTTTGATTTATGGTCATTCTCGTTTTOTGAACTGTTTAAAGCAT
TTGAGGGGGATTCAATGAATATTTATGACGATTCCGCAGTATTGGACGCTATCCAGTCTAAACATTTTACTATTACC
CCCTCTGGCAAAACTTCTTTTGCAAAAGCCTCTCGCTATTTTGGTTTTTATCGTCGTCTGGTAAACGAGGGTTATGA
TAGTGTTGCTCTTACTATGCCTCGTAATTCCTTTTGGCGTTATGTATCTGOATTAGTTGAATGTGGTATTCCTAAAT
CTCAACTGATGAATCTTTCTACCTGTAATAATGTTGTTCCGTTAGTTCGTTTTATTAACGTAGATTTTTCTTCCCAA
CGTCCTGACTGGTATAATGAGCCAGTTCTTAAAATCGCATAAGGTAATTCACAATGATTAAAGTTGAAATTAAACCA
TCTCAAGCCCAATTTACTACTCGTTCTGGTGTTTCTCGTCAGGGCAAGCCTTATTCACTGAATGAGCAGCTTTGTTA
CGTTGATTTGGGTAATGAATATCCGGTTCTTGTCAAGATTACTCTTGATGAAGGTCAGCCAGCCTATGCGCCTGGTC
TGTACACCGTTCATCTGTCCTCTTTCAAAGTTGGTCAGTTCGGTTCCCTTATGATTGACCGTCTGCGCOTCGTTCCG
GCTAAGTAACATGGAGCAGGTCGCGGATTTCGACACAATTTATCAGGCGATGATACAAATCTCCGTTGTACTTTGTT
TCGCGCTTGGTATAATCGCTGGGGGTOAAAGATGAGTGTTTTAGTGTATTCTTTCGCCTCTTTCGTTTTAGGTTGGT
GCCTTCGTAGTGGCATTACGTATTTTACCCGTTTAATGGAAACTTCCTCATGAAAAAGTCTTTAGTCCTCAAAGCCT
CTGTAGCCGTTGCTACCCTCGTTCCGATGCTGTCTTTCGCTGCTGAGGGTGACGATCCCGCAAAAGCGGCCTTTAAC
TCCCTGCAAGCCTCAGCGACCGAATATATCGGTTATGCGTGGGCGATGGTTGTTGTCATTGTCGGCGCAACTATCGG
TATCAAGCTGTTTAAGAAATTCACCTCGAAAGCAAGCTGATAAACCGATACAATTAAAGGCTCCTTTTGGAGCCTTT
TTTTTTGGAGATTTTCAACGTGAAAAAATTATTATTCGCAATTCCTTTAGTTGTTCCTTTCTATTCTCACTCCG
GATATC
ATAT
CTGAAACTGTTGAAAGTTGTTTAGCA
AAACCCCATACAGAAAATTCATTTACTAACGTCTGGAAAGACGACAAAACTTTAGATCGTTACGCTAACTATGAGGG
153
TTGTCTGTGGAATGCTACAGGCGTTGTAGTTTGTACTGGTGACGAAACTCAGTGTTACGGTACATGGGTTCCTATTG
GGCTTGCTATCCCTGAAAATGAGGGTGGTGGCTCTGAGGGTGGCGGTTCTGAGGGTGGCGGTTCTGAGGGTGGCGGT
ACTAAACCTCCTGAGTACGGTGATACACCTATTCCGGGCTATACTTATATCAACCCTCTCGACGGCACTTATCCGCC
TGGTACTGAGCAAAACCCCGCTAATCCTAATCCTTCTCTTGAGGAGTCTCAGCCTCTTAATACTTTCATGTTTCAGA
ATAATAGGTTCCGAAATAGGCAGGGGGCATTAACTGTTTATACGGGCACTGTTACTCAAGGCACTGACCCCGTTAAA
ACTTATTACCAGTACACTCCTGTATCATCAAAAGCCATGTATGACGCTTACTGGAACGGTAAATTCAGAGACTGCGC
TTTCCATTCTGGCTTTAATGAAGATCCATTCGTTTGTGAATATCAAGGCCAATCGTCTGACCTGCCTCAACCTCCTG
TCAATGCTGGCGGCGGCTCTGGTGGTGGTTCTGGTGGCGGCTCTGAGGGTGGTGGCTCTGAGGGTGGCGGTTCTGAG
GGTGGCGGCTCTGAGGGAGGCGGTTCCGGTGGTGGCTCTGGTTCCGGTGATTTTGATTATGAAAAGATGGCAAACGC
TAATAAGGGGGCTATGACCGAAAATGCCGATGAAAACGCGCTACAGTCTGACGCTAAAGGCAAACTTGATTCTGTCG
CTACTGATTACGGTGCTGCTATCGATGGTTTCATTGGTGACGTTTCCGGCCTTGCTAATGGTAATGGTGCTACTGGT
GATTTTGCTGGCTCTAATTCCCAAATGGCTCAAGTCGGTGACGGTGATAATTCACCTTTAATGAATAATTTCCGTCA
ATATTTACCTTCCCTCCCTCAATCGGTTGAATGTCGCCCTTTTGTCTTTAGCGCTGGTAAACCATATGAATTTTCTA
TTGATTGTGACAAAATAAACTTATTCCGTGGTGTCTTTGCGTTTCTTTTATATGTTGCCACCTTTATGTATGTATTT
TCTACGTTTGCTAACATACTGCGTAATAAGGAGTCTTAATCATGCCAGTTCTTTTGGGTATTCCGTTATTATTGCGT
TTCCTCGGTTTCCTTCTGGTAACTTTGTTCGGCTATCTGCTTACTTTTCTTAAAAAGGGCTTCGGTAAGATAGCTAT
TGCTATTTCATTGTTTCTTGCTCTTATTATTGGGCTTAACTCAATTCTTGTGGGTTATCTCTCTGATATTAGCGCTC
AATTACCCTCTGACTTTGTTCAGGGTGTTCAGTTAATTCTCCCGTCTAATGCGCTTCCCTGTTTTTATGTTATTCTC
TCTGTAAAGGCTGCTATTTTCATTTTTGACGTTAAACAAAAAATCGTTTCTTATTTGGATTGGGATAAATAATATGG
CTGTTTATTTTGTAACTGGCAAATTAGGCTCTGGAAAGACGCTCGTTAGCGTTGGTAAGATTCAGGATAAAATTGTA
GCTGGGTGCAAAATAGCAACTAATCTTGATTTAAGGCTTCAAAACCTCCCGCAAGTCGGGAGGTTCGCTAAAACGCC
TCGCGTTCTTAGAATACCGGATAAGCCTTCTATATCTGATTTGCTTGCTATTGGGCGCGGTAATGATTCCTACGATG
AAAATAAAAACGGCTTGCTTGTTCTCGATGAGTGCGGTACTTGGTTTAATACCCGTTCTTGGAATGATAAGGAAAGA
CAGCCGATTATTGATTGGTTTCTACATGCTCGTAAATTAGGATGGGATATTATTTTTCTTGTTCAGGACTTATCTAT
TGTTGATAAACAGGCGCGTTCTGCATTAGCTGAACATGTTGTTTATTGTCGTCGTCTGGACAGAATTACTTTACCTT
TTGTCGGTACTTTATATTCTCTTATTACTGGCTCGAAAATGCCTCTGCCTAAATTACATGTTGGCGTTGTTAAATAT
GGCGATTCTCAATTAAGCCCTACTGTTGAGCGTTGGCTTTATACTGGTAAGAATTTGTATAACGCATATGATACTAA
ACAGGCTTTTTCTAGTAATTATGATTCCGGTGTTTATTCTTATTTAACGCCTTATTTATCACACGGTCGGTATTTCA
AACCATTAAATTTAGGTCAGAAGATGAAATTAACTAAAATATATTTGAAAAAGTTTTCTCGCGTTCTTTGTCTTGCG
ATTGGATTTGCATCAGCATTTACATATAGTTATATAACCCAACCTAAGCCGGAGGTTAAAAAGGTAGTCTCTCAGAC
CTATGATTTTGATAAATTCACTATTGACTCTTCTCAGCGTCTTAATCTAAGCTATCGCTATGTTTTCAAGGATTCTA
AGGGAAAATTAATTAATAGCGACGATTTACAGAAGCAAGGTTATTCACTCACATATATTGATTTATGTACTGTTTCC
ATTAAAAAAGGTAATTCAAATGAAATTGTTAAATGTAATTAATTTTGTTTTCTTGATGTTTGTTTCATCATCTTCTT
TTGCTCAGGTAATTGAAATGAATAATTCGCCTCTGCGCGATTTTGTAACTTGGTATTCAAAGCAATCAGGCGAATCC
GTTATTGTTTCTCCCGATGTAAAAGGTACTGTTACTGTATATTCATCTGACGTTAAACCTGAAAATCTACGCAATTT
CTTTATTTCTGTTTTACGTGCTAATAATTTTGATATGGTTGGTTCAATTCCTTCCATAATTCAGAAGTATAATCCAA
ACAATCAGGATTATATTGATGAATTGCCATCATCTGATAATCAGGAATATGATGATAATTCCGCTCCTTCTGGTGGT
TTCTTTGTTCCGCAAAATGATAATGTTACTCAAACTTTTAAAATTAATAACGTTCGGGCAAAGGATTTAATACGAGT
TGTCGAATTGTTTGTAAAGTCTAATACTTCTAAATCCTCAAATGTATTATCTATTGACGGCTCTAATCTATTAGTTG
TTAGTGCACCTAAAGATATTTTAGATAACCTTCCTCAATTCCTTTCTACTGTTGATTTGCCAACTGACCAGATATTG
ATTGAGGGTTTGATATTTGAGGTTCAGCAAGGTGATGCTTTAGATTTTTCATTTGCTGCTGGCTCTCAGCGTGGCAC
TGTTGCAGGCGGTGTTAATACTGACCGCCTCACCTCTGTTTTATCTTCTGCTGGTGGTTCGTTCGGTATTTTTAATG
GCGATGTTTTAGGGCTATCAGTTCGCGCATTAAAGACTAATAGCCATTCAAAAATATTGTCTGTGCCACGTATTCTT
ACGCTTTCAGGTCAGAAGGGTTCTATCTCTGTTGGCCAGA-ATGTCCCTTTTATTACTGGTCGTGTGACTGGTGAATC
TGCCAATGTAAATAATCCATTTCAGACGATTGAGCGTCAAAATGTAGGTATTTCCATGAGCGTTTTTCCTGTTGCAA
TGGCTGGCGGTAATATTGTTCTGGATATTACCAGCAAGGCCGATAGTTTGAGTTCTTCTACTCAGGCAAGTGATGTT
ATTACTAATCAAAGAAGTATTGCTACAACGGTTAATTTGCGTGATGGACAGACTCTTTTACTCGGTGGCCTCACTGA
TTATAAAAACACTTCTCAAGATTCTGGCGTACCGTTCCTGTCTAAAATCCCTTTAATCGGCCTCCTGTTTAGCTCCC
GCTCTGATTCCAACGAGGAAAGCACGTTATACGTGCTCGTCAAAGCAACCATAGTA 3'
154
Figure 4.6: GATC controls. An approximately equimolar mixture of genomes containing a 'G',
'A', 'T', or a 'C' insertion at the distal site (but a pure 'G' at the proximal site) were mixed and
processed via the REAP assay. On interrogating the distal insertion site, an approximately 25%
output of each of the four bases was obtained. Similar results were obtained for the equimolar
mixture of genomes with the four bases inserted (individually) at the proximal site but with a
pure 'G'at the distal site.
GATC controls
100%
.. "
MA
90%
ST
RC
80%
70%
60%
50%
40%
30%
20%
10%
0%
C
155
Figure 4.7: Test of possible recombination between the distal and proximal sites. A 'T' base was
inserted at the proximal site, in combination with either an O6MeG lesion or a pure 'G' base
inserted at the distal site. The mutation frequencies obtained through the REAP assay confirm
that even though the E. coli strains are recA* and hence capable of performing recombination
across similar sequences, this does not occur between the proximal and distal sites.
Test of recombination
100%
90%
80%
MG
MA
ST
Mc
70%
60%-50%
40%
30%
20%
10%
0%
: O6MeG
156
Figure 4.8: Graphical representation of mutation frequency data. (a) Graph depicting actual
base composition numbers obtained at the site of interrogation after the REAP assay. Since the
lesion of interest in this study, O6MeG, is known to mutate solely to adenine (if not repaired),
the mutation frequency can be represented completely by the percentage conversion to
adenine. This observation is borne out by our results as well, and the new depiction of
mutation frequency for the same set of data as graph (a)is shown in graph (b) (and repeated in
Figure 4.8a). This format will be used for the remainder of this study.
(a)Actual data
100%
,
.
90%
80%
70%
60%
50%
40%
30%
20%
10%
M
0%
-
I..
K
(b) Graphical representation
50%
45%
40%
35%
30%
25%
20%
15%
10%
5%
-M
I
I
157
Figure 4.9: Mutation frequency of O6 MeG as a function of distance from the ori in an Ada+ Ogtrepair background in (a)GXG (n=3) and (b)AXG (n=2) nearest neighbor contexts.
(a) Mutation frequency as a function of distance from the ori in a GXG sequence context
50%
45%
40%**:
p-value
=0.004
C
.2 35%
ba
2
30%
25%
0
.IA 20%
4a
-15%
*:p-value
10%
0.002
5%
0%
(b) Mutation frequency as a function of distance from the ori in an AXG sequence context
50%
45%
40%
C
.2 35%
2 30%
0
25%
0
w
:t 20%
M
15%
-au
=0.002
10%
7T
158
Figure 4.10: Effect of lesion load on mutation frequency in an Ada+ Ogt- repair background in
(a) GXG (n=3) and (b) AXG (n=2) nearest neighbor contexts.
(a) Effect of lesion load in a GXG sequence context
50%
*:p-value =
0.02
45%
40%
35%
30%
25%
20%
15%
10%
5%
0%
(
fi
q
(b) Effect of lesion load in an AXG sequence context
50%
45%
40%
35%
30%
25%
20%
15%
10%
5%
0%
oOj
""""T
f : OMeG
159
Figure 4.11: (a) Mutation frequency of O6MeG (m6G) placed at the distal site when a known
replication block (m3T, THF) is placed at the proximal site. (b) Mutation frequency of blocking
lesions or 'T' control at proximal site.
(a) Mutation frequency at distal site
**: p-value = 6 x 10-6
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
- T
4,
(b) Mutation frequency at proximal site
100%
90%
80%
70%
60%
50%
T
T
40%
30%
20%
10%
0%
-4,4
O'MeG
160
4.6 Tables
Table 4.1: M13 genes and their functions. Reproduced from [6].
Gene
I
Il
III
IV
V
VI
VII
VIII
IX
X
Function
Copies
per virion
Morphogenesis
Synthesis of phage single-stranded and double-stranded DNA
Minor coat protein needed for adsorption to host
Morphogenesis
Single-stranded DNA binding protein; binds single-stranded DNA for
packaging
Minor coat protein (adsorption complex)
Minor coat protein (morphogenic end)
Major coat protein
Major coat protein (morphogenic end)
Synthesis of double-stranded DNA; switch from synthesis to
packaging
0
0
5
0
0
5
5
2000
5
0
161
Table 4.2: Possible candidates for replication-blocking lesions to be placed at the proximal site.
xG: benzo-expanded guanine, xC: benzo-expanded cytosine, m3T: 3-methylthymine, m1G: 14
methylguanine, THF: tetrahydrofuran, gC: 3,N -ethenocytosine.
Lesion
Bypass Efficiency
Mutagenicity
xG
10% in SOS- cells
50% in SOS+ cells
35% in SOS- cells
55% in SOS+ cells
7% in AlkB+ SOS- cells
5% in AlkB- SOS- cells
95% 'A' in SOS- cells
100% 'A' in SOS+ cells
90% 'C' in SOS- cells
85% 'C' in SOS+ cells
40% in AlkB+ SOS- cells
60% in AlkB- SOS- cells
22% in AlkB- SOS+ cells
47% in AlkB- SOS+ cells
cells
cells
cells
cells
cells
cells
cells
cells
5% in AlkB+ SOS- cells
80% in AlkB- SOS- cells
68% in AlkB- SOS+ cells
5% in AlkB+ SOS- cells
78% in AlkB- SOS- cells
70% in AlkB- SOS+ cells
55-60% 'T' in w.t. cells
xC
m3T
m1G
m1G
THF
17% in AlkB+ SOS3% in AlkB- SOS9% in AlkB- SOS+
20% in AlkB+ SOS3% in AlkB- SOS7% in AlkB- SOS+
2% in AlkB+ SOS5% in AlkB- SOS-
Reference
[89]
[89]
[40]
[40]
[41]
[41;44]
7% in AlkB- SOS+ cells
FC
25% in AlkB+ SOS- cells
18% in AlkB- SOS- cells
40% in AlkB+ SOS- cells
82% in AlkB- SOS- cells
30% in AlkB- SOS+ cells
85% in AlkB- SOS+ cells I
[41]
162
Table 4.3: Mutation frequency data in a GXG nearest neighbor sequence context. This data is
also shown in Figures 4.9a and 4.10a. The data in each row corresponds to the lesion/base
enclosed within a red box in that row.
? : OMeG
163
Table 4.4: Mutation frequency data in an AXG nearest neighbor sequence context. This data is
also shown in Figures 4.9b and 4.10b. The data in each row corresponds to the lesion/base
enclosed within a blue box in that row.
Y
Standard deviation
Average
Lesion/base
%G
%A
%T
O'MeG
%C %G %A
%T
%C
99.09 0.42 0.11 0.40 0.23 0.13 0.02 0.07
92.15 5.47 0.24 2.16 0.32 0.32 0.08 0.08
51.55 47.98 0.05 0.43 0.33 0.28 0.01 0.06
92.67
5.25
0.23
1.86
0.17
0.06
0.06
0.16
99.16
0.38
0.09
0.38
0.04 0.13
0.01
0.07
53.16 44.91 0.29 1.65 1.37 1.10 0.01 0.28
0.11
0.36
2.72
2.86
0.01
0.11
56.90 41.00 0.05
2.07
0.73
1.29 0.37
0.18
57.26
42.28
164
4.7 References
1. Hudson,R.E., Bergthorsson,U., Roth,J.R., and Ochman,H. (2002) Effect of chromosome
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Chapter 5:
Investigating the role of AlkB in the repair of N2-dG lesions and
demethylation of m6A in vivo
173
5.1 Introduction
The role of AIkB has been widely studied in the last thirty years. It is known to be an irondependent dioxygenase, which repairs a plethora of toxic lesions as part of the adaptive
response in E. coli, especially those present in a single-stranded context. Several homologs of
AIkB exist in prokaryotic and eukaryotic species, nine of which are known to exist in humans
alone. The conservation of this enzyme (and its function) across species makes it an important
weapon in the cellular arsenal against DNA damage.
AIkB was discovered in 1983 [1] and established as a part of the adaptive response soon after
[2;3]. Introducing the alkB gene into human cells conferred resistance to SN 2 alkylating agents
(but not SNI), and it was found that AlkB was involved in the repair of damaged DNA rather
than prevention of its formation [4]. A seminal discovery was made when it was found that
alkB mutants were especially defective in repairing methylation damage in a single-stranded
context, suggesting a role for AIkB in single-stranded DNA repair [5]. Although it was shown
that AIkB preferentially bound alkylated single-stranded DNA over the double-stranded version
[6], it was subsequently established that AIkB could repair 1-methyladenine and 3methylcytosine in both single-stranded and double-stranded DNA via a mechanism typical of
the iron- and ax-ketoglutarate-dependent dioxygenases [7;8]. AIkB can also reverse alkylation
damage in RNA [9;10].
methyladenine,
The spectrum of lesions repaired by AIkB now includes 1-
3-methylcytosine,
3-ethylcytosine
[11],
1-methylguanine
[11;12],
3-
methylthymine [11;12], 1-ethyladenine [13], and 1,N -ethanoadenine [14] in DNA. The enzyme
also repairs unsaturated exocyclic lesions such as 1,N -ethenoadenine and 3,N - ethenocytosine
174
by a unique mechanism that proceeds via an epoxide intermediate to generate a repaired base
with the release of glyoxal [15].
Although the alkB gene is not conserved across all species [16], AIkB homologs have been found
in viruses, bacteria, C.elegans, D.melanogaster, mice, rats, and humans [17]. In addition, there
is evidence in yeast to suggest that there might be other unrelated proteins that have a
functional homology with AIkB in species lacking an obvious AIkB homolog. Out of nine human
homologs, hABH2, hABH3, and the recently discovered FTO (fat mass and obesity associated
gene)
[18] show the mechanism characteristic of c -ketoglutarate- and iron-dependent
dioxygenases [13]. hABH2 repairs methylation damage by rotating out the adduct from the
DNA helix using an aromatic 'finger' as seen in the case of DNA glycosylases. However, AlkB
lacks this 'finger' residue and instead squeezes out the damaged base, leading the flanking
bases to stack and therefore cause a helix distortion. The difference in the base extrusion
mechanism between hABH2 and AlkB is speculated to explain the affinity of the former for
double-stranded templates and the latter for single-stranded ones [19]. hABH3, on the other
hand, shares the preference of AIkB for single-stranded templates (both DNA and RNA) [20],
and localizes in the nucleus and cytoplasm, whereas hABH2 is localized to the nucleus [21]. In
terms of substrate specificity, hABH2 and hABH3 are similar to AlkB and have similar substrate
recognition sequences.
Few of the other homologs have been studied in great detail. The FTO gene is involved in
regulating energy balance and has been associated with obesity [22;23]. Its product is a nuclear
175
protein that has been shown to repair 3-methylthymine,
1-methyladenine, and 3-
methylcytosine in single-stranded DNA [18], and 3-methyluracil in single-stranded RNA [241.
hABH1, bearing the strongest homology to AIkB, was shown to partially protect E. coli alkBcells from methyl methanesulfonate damage [251. More recently, it has been shown to be a
mitochondrial enzyme that repairs 3-methylcytosine in single-stranded DNA and RNA in vitro
[26].
hABH8 is different from the other homologs dicussed above in that it contains an
additional tRNA methyltransferase domain and an RNA binding motif, which enables it to
hydroxylate 5-methoxycarbonyl-methyluridine residues present at the wobble position in
tRNAs [27;28] . The function of the remaining homologs is unknown.
In this study, we seek to expand the in vivo substrate specificity of AlkB to a number of lesions,
namely 2-methylguanine (m2G), 2-ethylguanine (e2G), 2-tetrahydrofuran-2-yl-methylguanine
(THF), 2-furan-2-yl-methylguanine (FF or N 2-furfuryl-dG), and 6-methyladenine (m6A).
The
structures of these lesions are shown in Figure 5.1. The motivation for this work came from in
vitro studies done in our laboratory by Dr. Deyu Li, which hinted at the possibility of partial
repair of the N2 -alkylguanines and partial demethylation of m6A (data not shown).
The
experiments performed on the N2-dG lesions also tie in with those done with DinB presented in
Chapter 3 using the same set of lesions. While a previous study from our laboratory has
explored the mutagenicity and toxicity of m2G and e2G in AlkB+/- cells [29], we included these
lesions in the current study to tease out the influence of alkyl group size on N2-dG adduct
repairability by AIkB. 3-methylcytosine (m3C) is a known substrate for AlkB [11] and serves as
a positive control in our experiments.
176
6-Methyladenine is one of the three most common post-replicative base methylations (the
other two being 5-methylcytosine and N4-methylcytosine), and is found in both prokaryotic and
eukaryotic genomes [30]. It functions as an epigenetic signal in bacteria, modulating several
DNA-protein interactions, thereby serving a variety of regulatory functions including bacterial
defense against viruses, regulation of the start of replication in E. coli, separation and
reorganization of chromosomes after replication, strand discrimination for mismatch repair,
regulation of bacterial conjugation, and the packaging of phage DNA into capsids (see [30] for a
detailed review). The function of m6A in eukaryotes is not yet known. Recently, it was shown
that m6A present in RNA could be repaired by the human FTO enzyme [31]. While several DNA
methyltransferases, existing either as part of a restriction-modification system or as standalone
enzymes, form m6A in vivo, no demethylase has been found in bacteria that can remove this
modification. Our work tests AIkB as a possible candidate for that purpose.
5.2 Materials and Methods
The REAP and CRAB assays described here have been modified from the work of Delaney et ol.
[32]. Please refer to the original method for more detail and clarification.
5.2.1 Cell strains
All the E. coli strains used in this work contain the F' episome that enables infection by M13
phage. GW5100 was used for large scale preparation of M13 phage DNA, SCS110 (JM 110, end
Al) was used for amplification of progeny phage post-electroporation, and NR9050 was the
177
strain of choice for double agar plating with X-gal for blue-white selection of plaques. The E.
coli strains used to test for AlkB function were HK81 (as AB1157, but na/A) and HK82 (as
AB1157, but nalA alkB22; AlkB-deficient). The AIkB status of these strains was previously
confirmed by PCR amplification and sequencing of the region encoding the alkB gene [14;29].
5.2.2 Oligonucleotides
All oligonucleotides and primers were obtained from Integrated DNA Technologies (IDT) unless
specified otherwise. Sixteen-mer oligonucleotides of the sequence 5' GAAGACCTXGGCGTCC 3',
where X is the lesion of interest (m2G [29], e2G [29], THF [33], FF [33], m3C [11], m6A) were
synthesized and purified as described. Sixteen-mer oligonucleotides with the same sequence
but with X = G, A, T, or C, were used as a control. The 19-mer 'competitor' oligonucleotide of
the sequence 5' GAAGACCTGGTAGCGCAGG 3' was used in the CRAB assay.
Scaffold
oligonucleotides (5' GGTCTTCCACTGAATCATGGTCATAGC 3' and 5' AAAACGACGGCCAGTGAATT
GGACGC 3') were used to hold the 16-mers in place to the cleaved single-stranded M13 vector
during genome construction, and do not overlap the lesion-bearing region.
The forward primers used for the mutagenicity (REAP) and toxicity/bypass (CRAB) assays
spanned the M13 vector as well as the 5' end of the inserted oligonucleotide carrying the lesion
of interest. While the REAP reverse primer also spanned the vector and the 3' end of the same
oligonucleotide, thereby effecting a selective amplification of DNA from the progeny phage
resulting from only the lesion-carrying genomes, the CRAB reverse primer annealed only to the
M13 vector downstream of the inserted oligonucleotide. The primers were modified with an
178
in
aminoethoxyethyl ether group (Y) at the 5' end to prevent labeling with "P-y-ATP
subsequent reactions. The primers were of the sequence 5' YCAGCTATGACCATGATTCAGTGGA
AGAC 3' (CRAB forward primer; also used as the REAP forward primer), 5' YCAGGGTTTTCCCAGT
CACGACGTTGTAA 3' (CRAB reverse primer), and 5' YTGTAAAACGACGGCCAGTGAATTGGACG 3'
(REAP reverse primer).
5.2.3 Enzymes and chemicals
Pvull, Xmnl, EcoRl, Haelll, Bbsl, HinFl, T4 DNA Ligase, T4 DNA polymerase, BSA, and the enzyme
reaction buffers were from New England Biolabs. Shrimp alkaline phosphatase (SAP) was from
Roche. P1 nuclease, 5-bromo-4-chloro-3-indolyl- beta-D-galactopyranoside (X-gal), isopropyl @D-1-thiogalactopyranoside (IPTG) were from Sigma Aldrich. T4 Polynucleotide kinase was from
Affymetrix. Sephadex G-50 Fine resin was from Amersham Biosciences . Hydroxylapatite resin,
19:1 acrylamide:bisacrylamide solution, and N,N,N',N'-tetra-methyl- ethylenediamine (TEMED)
were from Bio-Rad. Phenol: chloroform: isoamyl alcohol (25:24:1; pH 8) was from Invitrogen.
3P-y-ATP
was from Perkin Elmer. Non-radioactive ATP was from GE Healthcare Lifesciences.
5.2.4 Double agar overlay plaque method for phage analysis
The double agar overlay method used in this work was adapted from Adams et al. [34]. This
method was used for enumerating initial electroporation events as well as phage titers to
ensure statistical robustness, but not for mutational analyses. Briefly, 10 ml of 2 x YT media
0
was inoculated with 2 ml of a saturated overnight culture of NR9050 and grown for 1 h at 37 C
with aeration. Three hundred ul of this culture was mixed with 10 ul IPTG (24 mg/ml), 25 ul of 1
179
% thiamine, and 40 ul X-Gal (40 mg/mI in DMF), and then added to 2.5 ml of top agar
maintained in a molten state at 52 0C. Appropriate dilutions of supernatants containing phage
particles were immediately mixed with the top agar and poured onto B-broth plates, which
were shaken to evenly spread the agar. After a 10 min incubation at room temperature (to
allow the top agar to solidify), the plates were incubated overnight at 37 'C to obtain dark blue,
light blue, or clear plaques.
5.2.5 M13 phage DNA
M13mp7(L2) phage single-stranded DNA was isolated as follows.
Various dilutions of a
previous stock of M13 phage supernatant were plated on a lawn of E. coli cells using the double
agar overlay method to obtain phage plaques. A well-isolated plaque was plugged using a
sterile Pasteur pipette and vortexed in 1 ml LB, 200 ul of which was used to make a starter
culture (grown overnight) by mixing with 10 ul of an overnight saturated culture of GW5100
cells in 10 ml LB. One milliliter of this phage starter culture was then used to inoculate GW5100
cells, which had been grown using 500 ul of an overnight saturated culture in 250 ml of fresh 2
x YT medium for 2 h at 37 0Cand shaken at 275 rpm. The inoculated culture was grown further
for 8 h at 37 *Cwith aeration, after which the cells were pelleted and discarded. The phage
were precipitated from the supernatant by addition of 4 % PEG 8000 MW and 0.5 M NaCl.
After overnight precipation at 4 0C,the phage were pelleted, resuspended in 5 ml TE pH 8, and
extracted with four washes of 3 ml 25:24:1 phenol:chloroform:isoamyl alcohol (Invitrogen, pH
8). The aqueous phase was passed through a 0.5 g hydroxylapatite column (BioRad), washed
with 5 ml TE, and eluted in 1 ml fractions with 12 ml of 0.16 M phosphate buffer. The DNA-
180
containing fractions were identified by spotting on an agarose plate containing ethidium
bromide. The phosphate buffer in those fractions was then exchanged for TE by three washes
in Microsep 100K spin dialysis columns (Pall Lifesciences). The DNA obtained was at a yield of >
1 pmol/ml 2 x YT and was stored at -20 0Cuntil further use.
5.2.6 Construction of genomes
The multiple cloning site in single-stranded M13mp7(L2) is designed to form a hairpin structure
that contains a functional EcoRl site. Twenty picomoles of M13 single-stranded DNA were
linearized by incubation with 40 U of EcoRl for 8 h at 23 "C. Scaffolds (25 pmol in 1 ul each)
were annealed to the ends of the linearized genome by incubation at 50 *Cfor 5 min followed
by cooling to 0 *C over 50 min. In addition, 30 pmol of each 16-mer oligonucleotides insert
were 5' phosphorylated by 15 U of T4 PNK, supplemented with 1x T4 PNK buffer, 1 mM ATP,
and 5 mM DTT and incubated at 37 *Cfor 1 h. The linearized genome was subsequently ligated
with the phosphorylated oligonucleotide for 8 h at 16 *C in a reaction volume of 75 ul
containing 1 mM ATP, 10 mM DTT, 25 ug/ml BSA, and 800 U T4 DNA ligase. To degrade
scaffolds and unligated oligonucleotides, the ligation mixture was treated with 0.25 U/ul T4
DNA polymerase for 4 h at 37 0C. Finally, the reaction volume was brought up to 110 ul with
water and extracted once with 100 ul 25:24:1 phenol:chloroform:isoamyl alcohol. The aqueous
phase was purified by three TE (pH 8) washes in Microsep 100K spin dialysis columns (Pall
Lifesciences) to remove any residual phenol and salts.
Recovery yields of 30-45 % were
obtained.
181
5.2.7 Genome validation and normalization
Prior to proceeding with the bypass and mutagenicity assays, the incorporation of lesioncontaining oligonucleotides was confirmed and the relative concentration of the constructed
genomes was determined and normalized using the following procedure:
A 10-fold molar
excess of scaffolds (previously used in constructing the genomes) were annealed to ~0.35 pmol
of genomes in 5 ul.
The genomes were cleaved with 10 U of HinFl and the resulting 5' end
dephosphorylated with 1 U of Shrimp alkaline phosphatase in a reaction volume of 8 ul by
incubation at 37 *Cfor 1 h followed by a 5 min incubation at 80 *Cand cooling down to 20 *C@
0.2 *C/s. The 5' ends were then labeled with 1.66 pmol of 1 2P-y-ATP (6000 Ci/mmol) in a total
reaction volume of 12 ul containing 1 x buffer 2, 5 mM DTT, 150 pmol cold ATP, 5 U T4 PNK, and
10 U Haelll, incubated at 37 *Cfor 1 h. The reaction was stopped by the addition of 12 ul 2 x
formamide loading buffer, and the products were resolved using 20 % PAGE until the xylene
cyanol dye migrated a distance of 12 cm. The bands corresponding to fully-ligated genomes
were then quantified using phosphoimagery and normalized with respect to one another. The
genomes were then diluted with water such that all the genomes were at the same final
concentration. The band generated from the competitor genome was used as a marker.
Post-normalization, a test electroporation was performed in HK81 competent cells using the
control genome mixed in different ratios with the competitor genome. The results of the test
electroporation were determined by plating the competent cells immediately after
electroporation using the phage-overlay method to yield a dark blue (control): light blue
182
(competitor) plaque count ratio. The ratio that yielded a 75:25 dark blue:light blue phage count
was selected for the bypass assay of the lesion-containing genomes.
5.2.8 Preparation and SOS induction of electrocompetent cells
Three baffled flasks containing 150 ml LB medium each were inoculated with 1.5 ml of a
saturated overnight culture of the strain to be transformed. The cultures were incubated at 37
0C and
shaken at 275 rpm for ~ 2.5 h until the cultures reached early log phase, as measured by
OD600 of ~0.5. The cell were then pelleted by centrifugation at 9500 rpm (Sorvall GSA rotor),
resuspended in 1 ml cold sterile water, and pooled to a final volume of 175 ml cold sterile
water. This process of washing, pelleting, and resuspending was repeated three times. The
final resuspension was in 4.8 ml 10% glycerol to obtain a final volume of 6 ml electrocompetent
cells, which were then aliquoted and stored at -20 *Cprior to use.
To induce the SOS response in cells, the three log phase cultures (with OD600 of ~0.5) were
pelleted, resuspended in 25 ml ice-cold MgSO 4 , and transferred to individual 150 x 15 ml
petridishes. SOS induction was achieved by irradiating the cells in each dish (without lids) with
45 J/m2 of 254 nm light. Immediate transfer to 125 ml of 2 x YT was followed by growth for 40
min at 37 *C. The cells were then centrifuged, resuspended in cold sterile water and pooled.
The washing, pelleting, and resuspending steps described above were followed and the cells
were finally stored at -20 0C.
183
5.2.9 CRAB assay
Genomes containing the lesions were mixed in a 75:25 ratio with the competitor genome in a
total volume of 6 ul and electroporated in triplicate into 100 ul competent cells in a 2 mm-gap
cuvette using 2.5 kV and 125 0. The cells were immediately transferred to 10 ml LB and an
aliquot of the freshly electroporated cells was immediately plated using the agar overlay
method to ensure that a minimum of 105 independent initial electroporation events occurred in
10 ml of culture. The cells were then grown for 6 h at 37 0C with aeration to amplify progeny
phage. The supernatants of the 6 h cultures were retained and plated using the agar overlay
method to confirm 10 4 -fold amplification in the progeny phage titer. Another round of
amplification was performed in order to ensure that the progeny phage being analyzed came
from genomes that entered the E. coli cells, and not the residual genomes that did not get
electroporated into cells but were still present in the milieu, since PCR is subsequently used.
This was done by infecting 10 ul of an overnight culture of SCS110 cells with 100 ul of the 6 h
supernatants in 10 ml LB and incubating for 7 h at 37 0C with aeration, after which the
supernatant was retained. Single-stranded M13 progeny phage DNA was isolated from 0.7 ml
of supernatant using a QlAprep Spin M13 Kit with final DNA suspension in 100 ul elution buffer.
CRAB forward and reverse primers were used to amplify the region of interest from 10 ul per
QlAprep elution sample in a total volume of 25 ul using 1.25U Pfu Turbo DNA polymerase, 25
mM of each dNTP, and 10 x Pfu Turbo buffer. The PCR program started by denaturing at 94 0C
for 5 min, then cycled 30 times at 94 *Cfor 30 s, 67 0C for 1 min, and 72 *Cfor 1 min, and finally
extended for 5 min at 72 0C . The volume was then made up to 110 ul using water and
extracted once using 25:24:1 phenol: chloroform: isoamyl alcohol to destroy the DNA
184
polymerase (including the exonuclease domain). The aqueous phase was passed through a
Sephadex G-50 Fine resin spin column to remove any remaining dNTPs and traces of phenol.
The purified PCR product was then treated with Bbsl (1.5 U for 4 ul of sample in a total volume
of 6 ul) and shrimp alkaline phosphatase (0.3 U) by incubating at 37 0Cfor 4 h, heating to 80 0C
for 5 min, and cooling to 20 *C@ 0.2 *C/s. The 5' ends were then labeled in a total volume of 8
ul with a mixture of non-radioactive ATP (20 pmol),
32
P-y-ATP (1.66 pmol of 10 uCi/ul at 6000
Ci/mmol), and 5 U of Optikinase/T4 PNK by incubation at 37 0Cfor 15 min, followed by 65 0Cfor
20 min and cooling to 23 *C@ 0.1 0C/s. The labeled product was then trimmed by Haelli (10 U
in a final volume of 10 ul) at 37 *Cfor 2 h, followed by addition of 10 ul 2 x formamide loading
dye, which quenched the reaction. The samples were then loaded onto a 20 % denaturing gel
and run for ~3.5 h at 550 V, until the xylene cyanol dye migrated 10.5 cm. The gel was then
exposed to a phosphoimager screen and quantified using a Storm 840 scanner. Band intensities
were quantified using ImageQuant software, and lesion bypass was measured by comparison of
the 18-mer band intensity (lesion signal) to the 21-mer band intensity (competitor signal).
5.2.10 REAP assay
The REAP assay methodology is identical to the CRAB assay except for the PCR primers. The
primers used for the REAP assay span both the vector as well as the insert at each end, thereby
effecting a selective amplification of DNA from only the progeny phage resulting from the
lesion-carrying genomes. Following electrophoresis, the 18-mer bands were excised from the
gel, and crushed and soaked overnight in 200 ul water. After desalting with Sephadex G-50 Fine
185
resin spin columns, the samples were lyophilized overnight to dryness, resuspended in 5 ul
containing 1 ug P1 Nuclease in 30 mM sodium acetate and 100 mM zinc chloride, and incubated
at 50 *Cfor 1 h. One ul of each sample was then spotted onto PEI-TLC plates and separated
using 200 ml of a saturated solution of (NH4) 2HPO 4 adjusted to pH 5.8.
After 12 h of
development, the TLC plates were air-dried and quantified using phosphoimagery.
5.3 Results
5.3.1 CRAB assay
The competitive replication of adduct bypass (CRAB) assay is a quantitative method that
determines to what extent a given lesion blocks DNA replication if left unrepaired. In essence, a
lesion-bearing genome is mixed with a nonlesion competitor genome in a specific input ratio
and passaged through E. coli cells of a given repair background. The output ratio of progeny
phage indicates the relative growth of the lesion-bearing genome with respect to the nonlesion
competitor genome; a decrease in the lesion: competitor output ratio (and hence the bypass
efficiency) signifies blockage of replication and therefore toxicity. The competitor genome acts
as an internal standard in this competitive assay. The CRAB assay was performed on all the N2_
dG lesions (m2G, e2G, THF, FF) but not on m6A. An unmodified 'G' genome and an m3C
genome were used as controls.
186
None of the N2-dG lesions showed any signs of being toxic to the cells (Figure 5.2 and Table
5.1). The bypass efficiencies in the absence of AIkB were close to 100 %. Introducing the
genomes in AlkB+ cells did not make a statistically significant change in the bypass of any of the
N2-dG lesions. A dramatic change is seen in the toxicity of m3C, going from extremely toxic in
AIkB- cells to completely 'bypassed' (probably through repair prior to the DNA polymerase
encountering the lesion site) in the presence of AIkB. The results for m3C echo those obtained
in a previous study [11], while the results for m2G and e2G are in contrast with another study
done in our laboratory [29].
Since all the N 2 dG lesions show almost complete bypass
efficiencies in AIkB+ and AIkB- cells, it would be non-informative to perform the assay using
SOS-induced cells to see if the SOS polymerases increase bypassability in vivo. Hence, the
analysis of progeny in SOS-induced cells using CRAB (bypass) primers was not performed.
5.3.2 REAP assay
The restriction endonuclease and postlabeling (REAP) assay determines the mutation frequency
and mutation composition after the lesion of interest, if not yet repaired, has been processed
by the intracellular replication machinery. The REAP assay was performed on all the N2 dG
lesions, m6A, m3C, unmodified G, unmodified A, and an approximately equimolar mixture of
genomes carrying unmodified 'G', 'A', 'T', and 'C' bases at the lesion site (denoted as GATC).
The only lesion to show a mutation specificity in the absence of AIkB was m3C, once again
confirming previously seen data [11]. None of the other lesions show any substantial mutations
in either the presence or absence of AIkB (Figure 5.3 and Table 5.3), which has been observed
187
previously for m2G and e2G [29]. We do see a 3 - 4 % mutation frequency for both m2G and
e2G in all three cell strains, but this number does not vary significantly on changing the AlkB or
SOS status of the cells. No change was seen when the SOS repair system was induced, except
for m3C which showed an increase in the mutation frequency from 78 % in AIkB-SOS- cells to 86
% in AlkB-SOS+ cells, mainly due to an increase in 'T' at the lesion site (after m3C pairing with
'A' by the SOS-induced DNA polymerases).
5.4 Discussion
In the present study, we tested a number of lesions as possible substrates for AlkB. While some
repair of these lesions was seen in vitro, the results from this study show that AIkB does not
have any discernable effect on either the toxicity or mutagenicity of these lesions. The data
would seem to indicate that the lesions under consideration are not mutagenic or toxic at all, or
that some other enzyme is responsible for the repair or bypass of said lesions. Other work
done by the author with the same set of N2-dG lesions in the presence or absence of DinB has
shown that at least two of those lesions (THF and FF) are toxic in the absence of DinB. While
the presence of DinB alleviates the toxicity, AIkB has no effect on the bypass efficiencies of the
lesions in DinB- cells (Figure 5.4 and Table 5.2). The bypass efficiencies for m2G and e2G in this
study (~100 %) are in contrast with those obtained in a previous study (~35-50 %) [29]. The
same cell strains and same batch of m2G and e2G oligonucleotides were used both in our study
as well as the previous study. We do not have a good explanation for this discrepancy;
however, the data obtained in the current work are internally consistent. Additionally, the
188
conclusion of the current work that AIkB does not have an effect on m2G and e2G is consistent
with this previous study. It is possible that the lesions (other than THF and FF) in fact pose no
toxic or mutagenic threat to the replicative machinery. While the alkyl groups in this set of N2_
dG lesions do lie along the base-pairing face, it is likely that they swivel out of the way due to
free rotation around the endocyclic carbon-N2-nitrogen bond.
Tolerance of minor groove
lesions (such as the set under study here) by replicative polymerases has also been speculated
[35].
It is unclear whether m6A should be toxic at all since it is found in the DNA as a marker for
normal cellular processes. Also, the methyl group is not big enough to be a steric hindrance to
replicative polymerases. Therefore, the fact that we do not see any mutagenicity with m6A
either in the presence or absence of AlkB does not imply that AIkB cannot demethylate this
adduct. It is possible that the methyl group does not block Watson-Crick pairing with thymine,
leading to negligible mutation frequency in AIkB- cells.
Further experiments employing a
different assay, possibly one involving mass spectrometry to detect the percentage of m6A
converted to A, would be required to ascertain whether AlkB can demethylate m6A in vivo.
In summary, the lesions shown in Figure 5.1 are not substrates for AIkB in vivo at the level of
one molecule of lesion-containing genome per E. coli cell containing its normal level of AIkB.
The lesions show neither mutagenicity nor toxicity as determined by the REAP and CRAB assays.
This study demonstrates that results obtained in an in vitro scenario do not always translate in
189
vivo. Further tests need to be designed and performed to confirm whether AlkB has any in vivo
enzymatic activity on m6A.
190
5.5 Figures
Figure 5.1: Structure of lesions in the AlkB study. The lesions are referred to by the following
abbreviations in the text: FF: 2-furan-2-yl-methylguanine, THF: 2-tetrahydrofuran-2-ylmethylguanine, m2G: 2-methyl guanine, e2G: 2-ethyl guanine, m6A: 6-methyladenine, and
m3C: 3-methylcytosine.
0
N
K<ZJ
N
0
DNA
FF
N
N:
N N
DNA
N
0
H
THF
-"NH
N
N
DNA
m2G
H3C,'NH
N
N
H
N
NN
I N
H
DNA
e2G
NH 2
N
</I
NN
DNA
m6A
N O
DNA
m3C
191
Figure 5.2: Bypass efficiencies in AlkB+ and AlkB- cells (both non-SOS-induced) as determined
by the CRAB assay. These data are also tabulated in Table 5.1.
'Bypass' of N2-dG lesions inAlkB+/AlkB- cells
140
mAIkBmAkB+
120
100
0
60
40
20
0
m2G
e2G
THF
FF
m3C
G
192
Figure 5.3: Mutagenicity results in (a) AlkB+/SOS-, (b)AlkB-/SOS-, and (c) AlkB-/SOS+ cells as determined by the
REAP assay. None of the lesions are mutagenic in any of the three repair backgrounds. The status of AlkB and SOS
were validated by using m3C, which serves as a positive control showing elimination of mutational signature in
AlkB+ cells and a statistical difference in mutation pattern in AlkB-/SOS- vs. AlkB-/SOS+ cells. These data are also
tabulated in Table 5.3.
(a) Mutation frequency in AlkB+ SOS-E G 0 A 2 T N C
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
GATC
m2G
e2G
THF
FF
m6A
G
m3C
(b)Mutation frequency in AlkB- SOS-m G MA
A
T AC
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
GATC
1
m2G
e2G
THF
FF
m6A
G
m3C
A
(c)Mutation frequency in AlkB- SOS+m G mA a T mC
100%
90%
80%
70%
60%
50%
40%
-
-FF----
30%
20%
10%
LI L
0%
GATC
m2G
e2G
TH F
FF
m6A
m3C
G
193
Figure 5.4: Bypass efficiencies of N2-dG lesions in AlkB+ and AlkB- (both DinB-) cells as
determined by the CRAB assay. See Chapter 3 for a detailed discussion. These data are also
tabulated in Table 5.2.
'Bypass' of N2-dG lesions in AlkB+/AlkB- cells
160
*AlkB* AkB+
140
120
$ 100
Ln
> 80
60
40
20
m2G
e2G
THF
FF
m3C
G
194
5.6 Tables
Table 5.1: Bypass efficiencies of N2-dG lesions in AlkB+ and AlkB- cells (both DinB+) as
determined by the CRAB assay. The data tabulated below is also shown in Figure 5.2.
Lesion/base
% Bypass in AlkB- cells
Std. Dev.
Avg.
% Bypass in AlkB+ cells
Std. Dev.
Avg.
m2G
e2G
THF
85.91
11.23
122.17
6.73
99.66
89.37
12.16
4.25
124.20
106.57
15.30
13.15
FF
76.20
3.31
88.25
15.46
1.24
100.00
0.13
0.00
72.14
100.00
13.90
0.00
m3C
G
Table 5.2: Bypass efficiencies of N2 -dG lesions in AlkB+ and AlkB- cells (both DinB-) as
determined by the CRAB assay. The data tabulated below is also shown in Figure 5.4.
Lesion/base
m2G
e2G
% Bypass in AlkB- cells
Std. Dev.
Avg.
9.65
92.16
10.39
106.14
% Bypass in AlkB+ cells
Std. Dev.
Avg.
3.23
94.64
3.92
121.19
THF
39.58
6.31
27.92
2.39
FF
m3C
G
36.12
7.48
100.00
2.44
0.63
0.00
28.06
136.62
100.00
2.85
11.37
0.00
195
Table 5.3: Mutagenicity results in (a) AlkB+/SOS-, (b) AlkB-/SOS-, and (c) AlkB-/SOS+ cells as determined
by the REAP assay. None of the lesions are mutagenic in any of the three repair backgrounds. The
status of AlkB and SOS were validated by using m3C, which serves as a positive control showing
elimination of mutational signature in AlkB+ cells and a statistical difference in mutation pattern in AlkB/SOS- vs. AlkB-/SOS+ cells. The data tabulated below is also shown in Figure 5.3.
(a)
AlkB+/SOS- cells
Lesion/base
GATC
m2G
e2G
THF
FF
m6A
m3C
G
A
%G
21.74
Average
%T
%A
25.95 28.97
%C
23.35
Standard deviation
%C
%G %A %T
0.74 0.58 0.50 0.53
95.70
3.31
0.19
0.80
1.14
1.08
0.04
0.20
96.46
97.70
98.51
0.58
0.53
2.35
0.51
0.32
98.51
0.30
0.18
0.25
0.22
0.14
0.16
1.01
1.54
0.95
0.77
99.01
0.37
1.33
0.16
0.30
0.24
0.28 0.03
0.20 0.10
0.08 0.05
0.23 0.03
0.03 0.03
0.12
1.05
0.08
0.06
0.26
97.67
0.74
0.13
1.46
0.32
0.15
0.05
0.32
0.50 98.59
0.08
0.83
0.12
0.36
0.06 0.25
(b) AlkB-/SOS- cells
Lesion/base
%G
Standard deviation
Average
%A
%T
%C
%G
%A
%T
%C
1.26
0.17
0.31
0.21
0.12
0.18
1.45
0.09
0.23
0.15
0.06
0.13
0.87
0.02
0.02
0.03
0.02
0.01
2.03
0.24
0.06
0.05
0.04
0.04
0.21
0.10
0.01
0.89
0.03
0.11
1.41
0.01
0.01
0.95
0.06
0.10
GATC
m2G
e2G
THF
FF
m6A
22.30
96.57
96.89
99.12
99.04
0.57
23.46
2.69
2.32
0.21
0.21
98.76
28.57
0.13
0.16
0.17
0.15
0.12
25.67
0.62
0.64
0.50
0.59
0.55
m3C
G
A
5.91
99.15
0.42
34.49
0.32
99.00
37.24
0.07
0.06
22.36
0.47
0.51
(c) AlkB-/SOS+ cells
Lesion/base
%G
18.14
GATC
96.12
m2G
96.55
e2G
98.42
THF
98.49
FF
0.51
m6A
4.49
m3C
98.39
G
0.45
A
Average
%C
%T
%A
27.31 32.28 22.27
0.61
0.22
3.04
0.54
0.25
2.65
0.58
0.28
0.72
0.58
0.27
0.67
0.64
1.14
97.71
32.47 50.18 12.86
0.41
0.15
1.05
0.27
0.08
99.19
Standard
%G %A
2.19 2.46
1.41 1.25
1.19 1.06
0.33 0.11
0.37 0.37
0.15 0.39
1.49 1.90
0.93 0.79
0.15 0.18
deviation
%T %C
2.38 2.31
0.07 0.36
0.03 0.18
0.05 0.28
0.06 0.16
0.23 0.08
2.48 0.49
0.08 0.13
0.04 0.07
196
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200
Appendix:
Publication: Delaney,J.C., Gao,J., Liu,H., Shrivastav,N., Essigmann,J.M.,
and Kool,E.T. (2009) Efficient replication bypass of size-expanded DNA
base pairs in bacterial cells. Angewandte Chemie International Edition,
48, 4524-4527.
Reprinted here with the permission of the publisher.
201
DOI: 10.1 002/anie.200805683
Unnatural
DNA
Bases
Efficient Replication Bypass of Size-Expanded DNA Base Pairs in
Bacterial Cells**
James C. Delaney, Jianmin Gao, Haibo Liu, Nidhi Shrivastav, John M. Essigmann,* and
Eric T Kool*
One of the long-standing aims of biomimetic chemistry has
been to develop molecules that function as much as possible
as the natural ones do.I1' Among biopolymers, nucleic acids
have long served as a test bed for bioinspired design. The
earliest focus of altered structures for DNA was the redesign
of the phosphodiester backbone,[2 ] and numerous studies
found backbone variants that assembled well into helices.
Moreover, in recent years, some altered sugars have also been
shown to be substrates for certain polymerase enzymes.[31
Such work suggests the possibility of future biological
activities associated with altered DNA backbones.
More recently, a number of laboratories have focused on
design of replacements for the DNA bases themselves, which
encode the chemical information of the cell.14 1 This is a
challenging goal, because biological enzymes have evolved
the ability to manipulate these bases and base pairs with
extraordinarily high selectivity. The polymerase replication of
designed DNA pairs has been studied in a number of
laboratories. Significant successes have been reported using
varied strategies, including base pairs with altered hydrogenbonding patterns,l4al pairs that lack hydrogen bonds altogether, [*-- p and pairs that have larger-than-natural dimen14
sions.q ,51 However, to date such designed base pairs have
generally not been tested in living cells. One recent exception
is a single isosteric replacement for a natural base that was
6
replicated efficiently.
Herein we describe the first tests of a nonnatural DNA
base-pair geometry in a living cell. We have inserted single
size-expanded DNA (xDNA) bases into phage genomes and
measured their replication in Escherichia coli cells. Surprisingly, although xDNA base pairs are considerably larger than
their natural counterparts, we find that they are bypassed by
the cellular replication machinery remarkably well. Indeed,
two of the designed pairs possess biological function that is
nearly indistinguishable from that of natural base pairs.
[*] Dr. J.C. Delaney, N. Shrivastav, Prof. Dr. J. M. Essigmann
Departments of Chemistry and Biological Engineering
Massachusetts Institute of Technology
Cambridge, MA 02139 (USA)
E-mail: jessig@mit.edu
Dr. J.Gao, Dr. H. Liu, Prof. Dr. E. T. Kool
Department of Chemistry, Stanford University
Stanford, CA 94305 (USA)
E-mail: kool@stanford.edu
[*
M
We thank the U.S. National Institutes of Health (CA80024 to J.M.E.
and GM63587 to E.T.K.) for support. J.G. and H.L. acknowledge
Stanford Graduate Fellowships.
Supporting information for this article is available on the WWW
under http://dx.doi.org/l0.1002/anie.200805683.
The four size-expanded DNA base pairs are shown in
Figure 1. The hydrogen-bonded pairs involve benzo homologues opposite complementary natural partners, yielding
base pairs 2.4 A larger than Watson-Crick pairs.16 J The
a)
H
N
H3
H-N 1
/N
N-H
4
INN
N
N
dR
dR
dRN
N
interScience-
©
2009
-.
dR
H-N
H
T-xA
H\'N-H
C-xG
CH3
0
NN
-N\
dR
-xN
0
H
A -xT
G -xC
5'- GAAGACCT X GGCGTCC - 3'
Figure 1. a) Structures of the four xDNA bases evaluated in cellular
replication, shown with their paired structures. b) DNA sequence
context for xDNA bases (at position X) inserted into single-stranded
M1 3 bacteriophage.
concept of a benzo-expanded base was first developed by
Leonard and co-workers, who developed lin-benzoadenine
and -guanine and studied them as ribonucleotide analogues
three decades agol We developed analogous benzopyrimidine C-nucleosides and studied the ability of these four
deoxynucleoside compounds (xA, xC, xG, xT) to form helices
composed of expanded-size pairs. Work to date has shown
that fully substituted duplexes of xDNA pairs are highly
stable and form right-handed helices with a backbone
conformation resembling B-DNA.Isa -f However, single
xDNA pairs substituted within natural DNA are destabilizing, presumably because of the mismatch in size between the
large pair and the naturally sized backbone surrounding it.?1d1
This mismatch could well present a challenge for a natural
DNA polymerase, since such enzymes (especially replicative
681
enzymes) can be highly sensitive to the sizes of base pairs.
However, living cells possess several different specialized
classes of polymerases, some of which function to bypass
damaged bases that are often larger than those of normal
DNA.19 Before completing extensive studies of in vitro
replication of xDNA bases by natural and modified polymerases, we decided to test whether the cellular machinery
already exists that might process these expanded bases.
WILEY
4524
0
H-NO
Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Angew. Chem.
Int.
Ed. 2009, 48, 4524 -4527
We measured the information-encoding capability of the
single xDNA bases by incorporating them separately into
oligonucleotides using standard phosphoramidite solid-phase
synthesis. 5 "a, The intact incorporation of expanded bases into
the DNA strands was confirmed by MALDI-TOF mass
spectrometry in all cases (see the Supporting Information).
The oligonucleotides were then ligated into an
M13 mp7(L2) single-stranded viral genome.["'I These modified genomes were then passaged through E. coli to quantify
the biological responses. The responses to be derived are
replication bypass efficiency (as scored by the amounts of
daughter phage produced with respect to an unmodified
competitor genome) and replication fidelity (measured by
isolation and composition analysis of the daughter sequence,
see below). A feature of this system is that there is no
complementary strand opposite the unnatural bases in this
single-stranded bacteriophage. This situation ensures that
outcomes from replication and mutagenesis are derived solely
from the initial replicative bypass of the modified bases,
rather than from preferential replication of an opposing
unmodified strand.
The ability of each of the four xDNA bases to support
DNA replication in vivo was addressed using a competitive
replication assay in which genomes bearing a nonstandard
base are mixed with an unmodified internal standard prior to
electroporation into cell." The concentration of each xDNA
base construct was determined in triplicate, with subsequent
normalization and transfection using a 2:1 ratio of xDNA/
competitor. The results showed that two of the four sizeexpanded bases were bypassed highly efficiently (Figure 2).
In E. coli that were not induced for a damage (SOS) response
to translesion DNA replication, the xA and xT bases were
bypassed with efficiencies that were, respectively, 80% and
73% of the natural guanine control. The xC base was
moderately well bypassed, with an efficiency of 29%, while
the signal dropped to 11 % for the xG base. When UV light
was used to induce the SOS response polymerases, bypass was
improved by a significant amount in all cases, with xA and xT
reaching the level of the unmodified guanine control. The xC
base increased to 53%, while the base that was the strongest
block to replication, xG, had the greatest response to the UVinduced SOS DNA polymerases, increasing to 45 % relative
to the phage genome containing a natural guanine base at the
test site. This result suggests that flexible damage-response
enzymes may assist in the synthesis or extension of a large
xDNA pair.
Having established that single xDNA base pairs can be
processed by the E. coli replication machinery, we proceeded
to evaluate which natural bases replaced the xDNA analogues
in the daughter phage that was recovered, using a published
restriction endonuclease/postlabeling assay." This assay
serves as a quantitative measure of the ability of the bacterial
polymerases to accurately read the chemical information
encoded in the large-size bases. In the first round of DNA
synthesis to produce the (-) strand of the phage DNA, a
natural base would be incorporated opposite the xDNA base,
thus making a size-expanded pair. In the next round, the allnatural DNA (-) strand would be replicated normally,
producing new, all-naturally substituted (+)-strand daughter
phage genomes that carry the sequence information encoded
by the xDNA base at the original test site.
We found that two of the four xDNA bases, xA and xC,
encoded their analogous replacements correctly (Figure 3).
The fidelity of replacement of xA by A in the daughter phage
- SOS
- SOS
E-) +sOs
120
100
80
-
G A T C
G A T C
G A T C
G A T C
G A T C
xT
xC
Template
a
60
Base:
40
Figure3. The replicative encoding capability of single xDNA bases in
E. coli. Shown are the relative amounts of each natural base at the test
site (X) that replaced the xDNA base shown after phage replication.
cc
G
xG
xA
20
0-
Template
Base:
NH
0
NH2
N~NH
R
G
dR
dR
xG
XA
xT
CR
X
Figure 2. The efficiency of replication bypass of single xDNA bases
in vivo. Bases were ligated into single-stranded M13 bacteriophage
and replicated in E. coli. Bypass efficiency was measured by quantifying
relative output signals from test and internal standard genomes and
normalizing to those from G at the test-site control. Results are shown
without (-SOS) and with (+ SOS) induced damage response.
Angew. Chem.
Int.
Ed. 2009, 48, 4524 -4527
is particularly striking, with 99% of the daughter phage
containing adenine at the test site in the genome. This finding
establishes that the xDNA-replicating polymerase correctly
incorporates T opposite xA despite the large size of this pair.
The xC analogue was also read correctly the majority of the
time, with replacement by C in 88% of the cases and by A
(implicating T-xC mispairing) in 10%. The other two xDNA
bases were read incorrectly, with T incorporation opposite
both xT and xG being dominant over the "correct" xDNA
@ 2009 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.angewandte.org
4525
pairing. Despite the misreading of xG, its coding ambiguity
was low, since it coded as A 95 % of the time. We also carried
out the same experiments with phage that had been passaged
through E. coli in which the SOS response had been induced
by UV light. The results (see the Supporting Information)
showed that this procedure did not significantly change the
bases encoded by the large-sized bases.
Although the xA and xC expanded bases are read
correctly by the bacterial replication machinery, xT and xG
are not, at least in this context. We speculate that the
mispairings that are observed with these latter two bases arise
from an alternative pairing geometry and from an alternative
protonation state (Figure S5 in the Supporting Information).
It is possible to pair T opposite xT with a geometry analogous
to the T-G wobble, which may be closer to a Watson-Crick
geometry. This might explain the observation of "correct" AxT pairing only 26% of the time as compared with 73% T-xT
mispairing. As for the xG base, in some contexts it can be
deprotonated at neutral and basic pH values.5 f' If such
deprotonation occurred during replication, xG would present
a structure that is more complementary to T than C. We note
that although two xDNA bases are incorrectly processed in
this context, a substantial information-encoding capability
remains. The correct coding of two xDNA pairs involves four
different bases, which is, in principle, the same amount of
information content as the natural genetic system.
The finding of efficient replication for two of the largesized pairs is surprising, given that replicative polymerases
8
can be highly sensitive to nucleobase size.l'
To examine this
aspect in more detail, we carried out preliminary in vitro
experiments with DNA polymerase I (Pol I; Klenow fragment (Kf)), the most extensively studied of the E. coli
polymerases. We used 28 mer DNA templates containing a
single xDNA base immediately downstream of a primer. To
evaluate enzymatic efficiency and selectivity, we performed
steady-state kinetics measurements on the addition of single
natural nucleotides opposite each of these large bases. The
results are given in Table 1.
The kinetics data confirm that at least one natural enzyme
can correctly read sequence information stored in sizeexpanded bases. Not surprisingly, the Kf enzyme is inefficient
in constructing these large base pairs, with Vm/Km values
approximately 100-1000-fold below those for a natural base
Table i: Steady-state kinetics data for insertion of nucleoside triphosphates opposite single xDNA bases by the Klenow fragment of DNA Pol I
(exonuclease-free (exo-)). [a
Base Nucleoside Vmx
triphos(% min- 1)[b]
phate
Km[ILM]
(VmaxKm)
Relative
efficiency
Efficiency
xA
xA
xA
xA
dATP
dCTP
dGTP
dTTP
0.0041 (0.0012) 23 (10)
0.0035 (0.0005) 9.9 (2.3)
0.0046 (0.0019) 43 (20)
0.15 (0.01)
18 (3)
1.9 x 102
3.9x 102
2
1.1 x 10
8.0x 10'
2.4 x 10-2
4.9x10-2
1.4 x 10-2
1
xC
xC
xC
xC
dATP
dCTP
dGTP
dTTP
0.40 (0.04)
0.065 (0.004)
3.5 (0.2)
0.19 (0.01)
8.7 x 104
2.0 x 104
1.3 x 10'
1.2 x 104
6.7 x10-1
1.5 x 10-1
1
9.2 x 10-2
xG
xG
xG
xG
dATP
dCTP
dGTP
dTTP
0.0034 (0.0004) 13 (10)
8 (10)
0.26 (0.06)
0.0046 (0.0011) 20(9)
0.0035 (0.0022) 15 (17)
3.8x10 2
7.5 x 104
2.7x10 2
1.3 x 102
5.1 x10-3
1
3.6x10-3
1.7x10-3
xT
xT
xT
dATP
dCTP
dGTP
dTTP
9.0 (0.3)
0.19 (0.03)
0.064 (0.008)
0.59 (0.03)
50(6)
160 (30)
880 (10)
17(2)
1.8x 105
1.2 x 10 3
7.7 x 10'
3.6x10 4
1
6.7 x 104.3 x10-4
2.0x 10-
T
dATP
3.4 (1.4)
1.1 x 107
-
xT
36 (4)
4.6 (1.5)
4.1 (1.5)
28 (3)
17 (5)
[a] Conditions: 200 nm 23 mer/28 mer primer-template duplex and
varied polymerase concentrations in a buffer containing 50 mm Tris-HCI
(pH 7.5), 10 mm MgCl 2 , 50 pgmL 1 BSA, and 1 mm dithiothreitol,
incubated at 37*C in a reaction volume of 10 mL. Standard deviations
(n = 3-5) are given in parentheses. [b] Normalized for the lowest enzyme
concentration used.
size-expanded bases, and 2) that the full replication machinery of E. coli is able to recognize the sequence encoded by
two of the xDNA bases correctly and efficiently. This
intriguing outcome suggests that it may be possible in the
future to incorporate multiple xA or xC bases into phage
genomes or to incorporate xDNA pairs into plasmids that
encode protein expression. In addition, it would be of interest
to explore whether other organisms might also possess the
ability to read xDNA pairs. The findings may ultimately lead
to new strategies for modifying biological systems in useful
ways.
pair. Interestingly, this polymerase selectively chose the
correct pairing partner with each of the four xDNA bases.
Moreover, preliminary experiments on extension of an xDNA
pair by this enzyme also show selective bypass of a correct
pair over mismatched ones, again with very low efficiency (see
the Supporting Information). It appears that enzyme(s) other
than Pol I are responsible for the efficient replication of xG
and xT in vivo, as these bases exhibited different cellular
coding efficiencies (Figure 3). It will be of interest in the
future to explore the other classes of polymerases, including
types that are functionally more flexible,112 to evaluate which
are able to efficiently replicate such large pairs. It will also be
important to study the replication in new sequence contexts
and to evaluate exonuclease proofreading of such pairs.
Taken together, the results show 1) that a DNA polymerase is able read the chemical information stored in the
4526
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4
2009
Experimental Section
Synthesis: The deoxynucleoside phosphoramidite derivatives of xA,
xT, xC, and xG were prepared as described previously."'I They were
incorporated into oligodeoxynucleotides using the published methods, purified by HPLC, and characterized by MALDI-TOF mass
spectrometry (see the Supporting Information).
Enzyme kinetics: 28-nt oligonucleotides containing single xA, xC,
xG, xT residues were prepared along with a 23-nt complementary
primer, which was labeled at its 5' end with 'P. Polymerase reaction
conditions were as follows: DNA concentration 5 pM in a 37*C buffer
containing 100 mm Tris-HCI (pH 7.5; Tris =tris(hydroxymethyl)aminomethane), 20 mm MgC 2 , 2 mm dithiothreitol, and 0.1 mgmL-acetylated bovine serum albumin (BSA). Enzyme and nucleotide
concentrations were varied. See the Supporting Information for
details.
Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Angew. Chemn.
Int. Ed. 2009, 48, 4524-4527
Cellular assays: The competitive replication of adduct bypass
(CRAB) assay[lobI was used to determine the replication blocking (if
any) by the size-expanded bases in M13 phage. Figure SI in the
Supporting Information shows an outline of the assay. Quantification
of the modified and control phage sequences was performed on the
daughter phage population as described. The restriction endonuclease and postlabeling determination of mutation frequency
(REAP) assayllobI was used to quantify the type and amount of
mutagenesis at the modified base site by obtaining the base
composition at that position after cellular replication (Figure S1 in
the Supporting Information). After PCR amplification, products were
cleaved, labeled, and enzymatically digested, then analyzed by TLC
and quantified by phosphorimagery. Experiments were performed in
triplicate.
Received: November 20, 2008
Published online: May 14, 2009
Keywords: DNA replication - nucleobases - polymerasesteric hindrance
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4527
Angewandte
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Zeitschri
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Geselschap
Deutscher
Chemike
e
Supporting Information
@Wiley-VCH 2009
69451 Weinheim, Germany
Supporting data
Efficient Replication Bypass of Size-expanded DNA Base Pairs in Bacterial Cells
James C. Delaneyj Jianmin Gao, Haibo Liu, Nidhi Shrivastavl, John M. Essigmann*j and Eric T.
Kool*
Department of Chemistry, Stanford University, Stanford, CA 94305; *Departmentsof Chemistry
and BiologicalEngineering, MassachusettsInstitute of Technology, Cambridge, MA 02139.
Contents
Oligodeoxynucleotide synthesis methods. (p. S2)
MALDI-TOF data for characterization of xDNA-containing oligonucleotides (p. S2)
Enzyme kinetics methods. (p. S3)
Methods for cellular assays (p. S3)
Figure Si. Schematic representation of CRAB bypass efficiency assay and REAP mutagenesis assay (p.
S4)
Figure S2. Gels for lesion bypass determination (p. S5)
Figure S3. Example of TLC chromatogram used for mutation determination (p. S6)
Figure S4. Mutational analysis for replication of xDNA-containing phage after induction of SOS
response (p. S7)
Figure S5. Hypothesized structures that may explain mispairing of T with xT and xG in the cellular
experiments. (p. S8)
Table S1. Steady-state kinetics for polymerase extension beyond pairs and mispairs of an xA base. (p.
S8)
Oligonucleotide Synthesis. Oligodeoxynucleotides were synthesized on an Applied Biosystems 394
DNA/RNA synthesizer on a 1 [tmole scale and possessed a 3'-phosphate group. Coupling employed
standard p-cyanoethyl phosphoramidite chemistry, but with extended coupling time (600 s) for
nonnatural nucleotides. All oligomers were deprotected in concentrated ammonium hydroxide (55 'C,
16 h), purified by preparative 20% denaturing polyacrylamide gel electrophoresis, and isolated by
excision and extraction from the gel, followed by dialysis against water. The recovered material was
quantified by absorbance at 260 nm with molar extinction coefficients determined by the nearest
neighbor method. Molar extinction coefficients for unnatural oligomers were estimated by adding the
measured value of the molar extinction coefficient of the unnatural nucleoside (at 260nm) to the
calculated value for the natural DNA fragments. Previous studies have shown that xDNA bases have
very low hypochromicity in xDNA oligomers. Molar extinction coefficients for xDNA nucleosides
used were as follows: dxA, E260=19,800 M-c;m- dxG, E260=8,100 M-cm'; dxT, c260=1,200 M'-cm';
dxC, F260=5,800 M--cm'.
MALDI-TOF mass spectrometry was performed on the oligonucleotides that were to be ligated into
the M13 viral genome, using a PerSeptive Biosystems Voyager-DE STR BioSpectrometry Workstation
run in the negative ion linear mode, and conditions as described.' Masses for single, negatively charged
16mer DNA molecules of sequence 5' GAAGACCTXGGCGTCC 3' are as follows:
X = G 4,906.22 observed (4,906.18 calculated)
X = xG 4,956.19 observed (4,956.24 calculated)
X = xA 4,940.27 observed (4,940.24 calculated)
X = xT 4,931.19 observed (4,931.23 calculated)
X = xC 4,930.24 observed (4,930.24 calculated)
(1) Delaney, J. C., and Essigmann, J. M. (2004) Mutagenesis, genotoxicity, and repair of 1-methyladenine, 3alkylcytosines, 1-methylguanine, and 3-methylthymine in alkB Escherichiaco/i. Proc. Natl. Acad. Sci. U. S. A. 101,
14051-14056.
Enzyme kinetics methods.
The 5'-terminus of the primer was labeled using [y- 32 P]ATP and T4 polynucleotide kinase. The labeled
primer as annealed to the template in a buffer of 100 mM Tris*HCl (pH 7.5), 20 mM MgCl 2 , 2 mM
DTT, and 0.1 mg/mL acetylated BSA. Polymerase reactions were started by mixing equal volumes of
solution A containing the DNA-enzyme complex and solution B containing dNTP substrates. Solution
A was made by adding Klenow fragment (exo-) (Amersham) diluted in annealing buffer to the annealed
duplex DNA and incubating for 2 min at 37 'C and terminated by adding 1.5 volumes of stop buffer
(95% formamide, 20 mM EDTA, 0.05% xylene cyanol and bromophenol blue).
2
Steady state kinetics for standing start single nucleotide insertions was carried out as described.
The final DNA concentration was 5 [tM. Final concentrations of triphosphates were 15, 25, 40, 60, 90,
120 tM. Amount of polymerase used (0.05-0.1 u/pL) and reaction time (2-60 min) were adjusted to
give <20% conversion. Extents of reaction were determined by running quenched reaction samples on
20% denaturing polyacrylamide gel. Relative velocities were calculated as extent of reaction divided by
reaction time and normalized to 0.005u/RL enzyme concentration.
Methods for cellular assays.
The Competitive Replication of Adduct Bypass (CRAB) assay determines the replication blocking
power of lesions, whereby a lesion-bearing genome is mixed with a nonlesion internal standard genome
prior to transfection (Figure S1, left). Lesions that block replication cause an increased percentage of
competitor signal in the output phage population, whereas the output from non-blocking lesions should
maintain the original lesion:competitor genome input ratio. Quantification of the lesion and competitor
2
output is performed on the output population as described, using a specific set of PCR primers that
amplifies both lesion and competitor signals equally. PCR products from the competitor signal are 3
bases longer than from the lesion signal, and can therefore be discriminated on a denaturing gel, and
quantified by PhosphorImagery.
The Restriction Endonuclease And Postlabeling determination of mutation frequency (REAP) assay
quantifies the type and amount of mutagenesis at the lesion site by obtaining the base composition at the
lesion site after cellular replication (Figure S1, right). The methodology is similar to the CRAB assay;
however, the primer set used will amplify only signal stemming from the lesion. PCR product from this
100% lesion signal is digested with BbsI, cleaving a fixed number of bases outside of its recognition
sequence to expose the lesion site as a 5'-overhang that is ultimately radiolabeled, trimmed with HaeIII,
and the products are separated on a denaturing gel. After the band containing the lesion site is excised,
eluted, desalted, and dried, the residue is digested to 5'-dNMPs with nuclease P1, thus releasing the
bases at the former lesion site as radioactive species, that are resolved by TLC and quantified by
PhosphorImagery.
If lesion hinders replication
(+3)
C
GAAGA
AG,( /
GAAN.ACCTCGC
GGCGTC
67%
25%
33%
Input
GAAACCGiCGAAGCCTGGTAGCCAG(
GAAGACCTGGTAGCGCAG
Lesion
Genome
Competitor
Genome
25%
75%
75%
Output
REAP
mutagenesis assay
CRAB
bypass efficiency assay
CC
AC"T!
XGGCGT
Competitor
Genome
Lesion
Genome
Cells
Competitor
Genome
Lesion
Genome
GAAiGAC
TGGTACGCG
GAGACCTGGCGTCC
\
Lesion
Genome
Com petitor
Genome
75%
25%
PCR amplify with primers designed to amplify
only region that had contained lesion
PCR amplify with primers designed
to amplify lesion and competitor equally
75% competitor signal (3 bases longer)
100% lesion signal
61mer
25% lesion signal
H:N
For CRAB &REAP analysis:
Cut with Bbsl &dephosphorylate
"P(l)-Label new 5' ends &Haelll trim
Separate products via PAGE
Bbsl t
34mer
HN
Shown in detail for REAP (right)..
CRAB works similarly CRAB
+3 competitor signallesion or non-lesion-*
HaeIII
14mer
18mer
13mer
13mer
NH
REAP
Excise oligo whose P-labeled
5' base was the lesion site
Digest with Nuclease P1
Resolve 5'dNMPs via TLC
21mer
.
-
30mer
34mer
1
NH2
at
-4
Quantify via Phosphorlmagery
.1mer
control signal
(
Lesion signal
+3 competitor signal
Bypass=
Quantify frameshifts'
if applicable
\J0)
Control signal
/
x
F
Mutation = 1 e
10(
its competitor
5'dCMP
5'dTMP
5'dGMP
5'dAMP
M
Origin
Figure S1. Schematic representation of CRAB bypass efficiency assay (left) and REAP mutagenesis
assay (right), adapted from prior work. The concentration of each genome construct (xDNA or G at
lesion site reference control) was determined in triplicate and normalized. Each 2:1 formulation of
lesion genome:competitor genome made for the xG, xA, xT, xC and G genomes was electroporated in
the order shown, and this transfection process was repeated twice. Greater than 4 x 106 initial events
were obtained per electroporation, with greater than 4 x 105 events stemming from the least bypassed
lesion, providing for statistical robustness.
(2) Kim, T. W.; Delaney, J. C.; Essigmann, J. M.; Kool, E. T. Proc. Nati. Acad. Sci. USA 2005, 102, 1580315808.
(3) Delaney, J. C., and Essigmann, J. M. (2006) Assays for determining lesion bypass efficiency and mutagenicity
of site-specific DNA lesions in vivo. Methods Enzymol. 408, 1-15.
(4) Delaney, J. C., and Essigmann, J. M. (2008) Biological properties of single chemical-DNA adducts: A twenty
year perspective. Chem. Res. Toxicol. 21, 232-252.
Gels used for lesion bypass determination
(CRAB assay) in uninduced (-SOS) cells
wells-)
+3competitor-*
AW *W %
lesion site -0
401W4
W
0
Template
Base:
0
H3C
NH2
N
N
dR
NH2
I
NH
N
N
N
dR
G
NH2
0
NH2
xG
NH2
dR
xA
dR
H
xT
dR
H
xC
Figure S2. Gels used for lesion bypass determination (CRAB assay) in uninduced (-SOS) cells,
showing the output after growth in liquid culture from each normalized genome construct mixed with
competitor genome (2:1 lesion:competitor ratio) electroporated in triplicate. The xG lesion was the least
bypassed, providing a lesion site signal of 10%, and a +3 competitor signal of 90%. The lower limit
estimate of 4 x 105 for the number of initial events arising from xG is calculated by multiplying the
lesion site signal by the number of total initial events (obtained by immediate plating of a portion of the
transformed cells). The actual number of xG initial events may be even higher, since the
lesion:competitor output from immediate plating, which can underestimate the genotoxicity of DNA
lesions, is greater than that seen after growth in liquid culture.'
TLC used for mutation determination
(REAP assay) in uninduced (-SOS) cells
5' dCMP
S
5'dTMP
5'dGMP
0
Template
Base:
0
0
0
.4-a
*
U)
.4-
5' dAMP
4- Origin
G xG xA xT xC
Figure S3. TLC used for mutation determination (REAP assay) in uninduced (-SOS) cells (one of three
replicates shown).
100-
+SOS
90
G A T
C
G A T C
Template
Base:
G A T C
0
0
N
NZNH
2
N
N-
dR
G
xG
C
NH2
N
xA
NH2
ON
N
H
N
dR
G A T C
0
H3C
NH
dR
G A T
NH2
dR
xT
H3 C
N
0
dR
H
xC
Figure S4. The fidelity of replication of single xDNA bases in Escherichiacoli. Shown are the relative
amounts of each natural base that replaced the xDNA base shown after phage replication occurred in E.
coli cells that had the SOS response induced with UV light. The largest change with respect to
uninduced cells was for xT, in which the value for T increased from 26% to 36%, with a decrease in A
from 73% to 63% (compare to Fig. 3 in main text).
CH3
N-H-
0
0
~13L
N
0
0
dR
H-N
dR/
0
NH
dR
N
N-H
N
N
H-N
H
0
T - xG mismatch
T - xT mismatch
Figure S5. Hypothesized structures that may explain mispairing of T with xT and xG in the cellular
experiments.
Table S1. Steady-state kinetics data for extension of xDNA pairs and mispairs by the
Klenow fragment of DNA Pol I (exo-). Data are for addition of dCTP opposite a natural
template G at the primer-template terminus beyond the xDNA pairs shown.a
primer
base
(%min)b
m (t )
efficiency
(Vmax / Km)
relative
efficiency
xA
A
0.0011 (0.0001)
141 (6)
7.7 x 100
1.4 x 10-1
xA
C
ND
ND
< 100C
<1.8 x 10-2
xA
G
0.00092 (0.00003) 131 (13)
7.2 x 100
1.3 x 10-1
xA
T
0.011 (0.002)
248 (87)
5.5 x 101
1
template
base
Vmax
K
M
aConditions: 200 nM 23mer/28mer primer-template duplex and varied polymerase concentrations in a
buffer containing 50 mM Tris-HCI (pH 7.5), 10 mM MgCl 2 , 50 ug/mL BSA and 1 mM dithiothreitol,
incubated at 37C in a reaction volume of 10 pL Standard deviations are given in parentheses.
bNormalized for the lowest enzyme concentration used.
cValue is an upper limit; extremely poor efficeincy (virtually no dCTP insertion) was observed.