Biochemical Characterizationof the E. coli
Very Short Patch Repair Pathway and its Coordination with
Methyltransferase Repair of 0 6 -Methylguanine
by
Peter Thomas Rye
B.S., Biochemistry (1999)
Syracuse University in New York
Submitted to the Department of Chemistry in Partial
Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biological Chemistry
at the
Massachusetts Institute of Technology
June 2006
© 2006 Massachusetts Institute of Technology. All Rights Reserved.
The author hereby grants to MIT permission to reproduce
and to distribute publicly paper and electronic
copies of this thesis document in whole or in part.
Signature of Author:
Department of Chemistry
April 7, 2006
Certified by:
Jo~haM. Essigmann
William and Betsy Leitch Professor of Chemistry and Toxicology
Thesis Supervisor
Accepted by:
Robert W. Field
ti. jm;
o .Li.UrI
.TalaLmo,.AnThwr,,,3.
Chairman, Department Committee on Graduate Students
,r
MASSACHUSETTS INSTITUTE
OF TECHNOLOGY
AUG 0 1 2006
ARCHIVES
LIBRARIES
This doctoral thesis has been examined by a committee
ofte Department of Chemistry as follows:
Gerald
N.Wogan.,
-
Professor of Chemisfry ancdBiologicalEngineering
Chairman
John M. Essigmann
William and Betsy LeitchffRessor ohemistry
Thesis Supervisor
and Toxicology
Barbara Imperiali
Class of 1922 Professor of Chemistry and Professor of Biology
2
Biochemical Characterization of the E. coli
Very Short Patch Repair Pathway and its Coordination with
Methyltransferase Repair of 06-Methylguanine
by
Peter Thomas Rye
Submitted to the Department of Chemistry on April 28, 2006
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy in Biological Chemistry
The E. coli Very Short Patch Repair (VSPR) system corrects T:G mismatches that
arise through Dcm-mediated methylation and subsequent deamination of the underlined
cytosine residue in the palindromic sequence 5'-CCWGG-3' (W is an adenine or
thymine). Vsr initiates VSPR by producing a single stranded nick on the 5' side of the
mismatched T. The MutS and MutL mismatch recognition proteins stimulate this
activity, as cells lacking either of these proteins display diminished VSPR. Genetic
studies also indicate that Pol I is responsible for removing and replacing a short tract of
nucleotides downstream of the incision site and that DNA Ligase seals the nick to
complete the repair event. However, until now, biochemical investigation of the repair
steps downstream of Vsr incision have been lacking.
Herein, we describe two novel in vitro assays used to probe the biochemical
events of VSPR. The first was used to verify the reconstitution of VSPR using purified
E. coli Vsr, Pol I, and DNA Ligase enzymes, while the second was used to measure the
distribution of VSPR patch sizes in whole cell extracts. By monitoring the loss of
radiosignal from a series of substrates that contained the label at prescribed distances
downstream of the T:G mismatch, we were able to determine that VSPR patches are
distributed around 2 to 4 deoxynucleotides in length. Interestingly, under certain reaction
conditions, the addition of DNA Ligase improved the efficiency of repair initiation by
Vsr, suggesting that VSPR may be optimal in the context of a multi-protein complex.
Lastly, we investigated the effect of VSPR proteins on methyltransferase
(MTase) repair of O6-methylguanine (6mG). MTase repair of O6mG opposite T results
in a G:T mismatch that must be further processed to yield the native G:C base pairing.
The G:T mismatch is therefore an intersection of the two pathways and led us to
hypothesize that MTase and VSPR proteins might interact. Indeed, cells lacking the
functions of MutS, MutL, or Vsr proteins displayed decreased MTase repair in vivo,
revealing a previously unknown interaction. The cooperation between proteins of these
two repair systems may shed light on the biological significance of the VSPR system.
Thesis Supervisor: John M. Essigmann
Title: William and Betsy Leitch Professor of Chemistry and Toxicology
3
ACKNOWLEDGMENTS
I would like to thank my advisor, Dr. John Essigmann, for his guidance and
support throughout the years. John was consistently receptive to my ideas and very
helpful in thinking them through. His energy in his work provided a great model to
follow and inspired me to achieve the standards of excellence he set for himself. He is a
brilliant scientist and has an amazing talent to communicate ideas. I thank him for his
time and persistence in teaching me those skills, and providing, all the while, an
enjoyable and productive environment in which to work. John is an extremely kind and
selfless individual, and he has taught me more than he will ever be know.
I would also like to thank my thesis committee members, Drs. Gerald Wogan and
Barbara Imperiali, for taking the time and energy to assess my work and provide valuable
feedback. I am especially indebted to Dr. Wogan for continually inspiring me to seek a
greater understanding of my work, the implications it may have, and how to take it to the
next level.
Everyone in the Essigmann Laboratory has positively impacted me in some way.
I need to thank Sarah, Kyle, Jim, Lauren, and Sreeja for being such great friends and
reading my many thesis drafts and suggesting how to make them better. Your many
comments were appreciated and helped enormously in defining the quality of my work.
Many thanks go to Kim for her attention and kindness in dealing with administrative
matters. I am convinced no other could take her place. I would also like to thank many
individuals (past and present) for their friendship and scientific discussions. Aida, Alfio,
Annie, Bob, Bogdan, Charles, Denise, Eunsuk, Francis, Jeannette, Jen, Jim, John,
Kaushik, Maryann, Michelle, Neena, Nicole, Paul, Pei-sze, Shawn, Uday, Will, Yuri, and
Zoran were all excellent people with whom to work. I hope I stay in touch with you all
and I look forward to seeing where your careers lead you. I consider all of you my
friends, and I hope that I had impacted your life in a positive way like you have impacted
mine.
Special thanks go to the numerous undergraduates I have had the privilege to
mentor. Lucien, Meghan, Ai-ris, Jenny, Dana, and Jelly were all amazing students to
teach and also to learn from. The saying that "there's no greater way to learn than by
teaching" holds very true. You all kept me on my toes by demanding an unparalleled
understanding of our work. I owe a great amount of what I understand to our discussions.
I also owe a great amount of what we accomplished in the lab to your efforts. I truly
appreciate the devotion you all displayed in your work and the passion you had towards
accumulating meaningful data. I hope that you feel great value, as I do, in the UROP
experience we shared.
Many thanks go to fellow chemistry graduate students. I remember the great
friendships formed in my first semester at MIT. I thank Aaron, Abi, Fran, Jen, Lyn,
Mary, Matt, Shawn, Woody, and Zarixia for helping to make MIT the right choice. Our
first year was one of the best. I look forward to seeing where you all end up. To Hector,
4
thanks for everything. You are an amazing person and friend. I hope we never lose
touch.
I would like to thank the "Merrimack Crew" for keeping me grounded, cheering
me on, and always being around. I will forever be grateful for the friendship of Andy,
Cory, Derek, Erin, Greg, Holly, Jamie, Jen, Jennie, Jesse, Jessica, Mandy, Mark, Matt,
Meredith, Mike, Mike, Salstrom, Sarah, and Steve. I feel blessed every day that I have
such wonderful people in my life who I can count on for anything. Thanks to you, and
your significant others, for everything in the past, present, and (most exciting) the future.
One word, LIFE. I love you all.
I am also thankful for Syracuse folks including Aleta, Ben, Bry, Cindy, Edi,
Green, Hosmer, Jill, Jordan, Kyoko, Larissa, Maria, Meth, Mooney, Morgan, Moran,
Rachael, Rob, Steve, and Zander. You all had a positive impact on my life and helped
me to remember the things that are truly important. May we continue to keep the good
times rolling.
I would also like to acknowledge friends that I have made along the way. I feel
fortunate to have had the support from people such as Cara, Carolyn, Kristen, Molly,
Paul, Sylvia, and TK. You all have made the last years of my life more enjoyable.
Special thanks goes to Sarah for her irreplaceable friendship. She has always helped me
to enjoy life and appreciate its many graces. I will forever be touched by her unyielding
optimism and the friendship we share.
Lastly, I would like to extend the largest thank you possible to my wonderful
family. I have received so much love and support from my two sisters, Elise and Heide,
their husbands, Paul and Randy, and their families. I am deeply touched by how much
they selflessly helped me through the tough times and celebrated with me through the
good ones. It was also amazing to have received copious amounts of letters, drawings,
pictures, phone calls, and hugs from my wonderful nieces and nephews Ben, Billy,
Claire, Hope, Jane, Katie, Kay, Morgan, and Sara. You all are amazing people and I
thank you for the love and support that you gave me.
To my parents, the two most generous, kind, wise, fun, and loving people in my
life, thank you for everything. You guys are always such an amazing source of direction,
strength, and inspiration. I cannot thank you enough for the endless support you give me.
Your attitude and advice keep me going through the rocky times and your excitement
during the highs is incomparable. I thank God that I have such wonderful parents to
push, pull, and stand by when needed. I also thank you for your talents in knowing just
how to help me see things through. I know, from experience, that I can always look to
your teachings for clarity and your example for guidance. By far and away, you
contribute to my life's work the most. Therefore, I dedicate this thesis to you.
Remember, this is only the beginning.
PETERT. RYE
5
TABLE OF CONTENTS
COMMITTEE PAGE
2
ABSTRACT
3
ACKNOWLEDGMENTS
.
4
TABLE OF CONTENTS
.
6
ABBREVIATIONS
9
LIST OFFIGURES
.
.
.
.
.
.
.
.
.
12
LIST OFTABLES
.
.
.
.
.
.
.
.
.
15
LIST OF SCHEMES
.
.
.
.
.
.
.
.
.
16
MismatchRepair in Escherichiacoli.
.
.
.
.
17
1.1.
Mutation Avoidance
.
.
.
.
18
1.2.
The Very Short Patch Repair System.
.
.
.
.
19
1.2.1. Dcm - in Vivo and in Vitro Properties.
.
.
.
22
1.2.2. Vsr - in Vivo and in Vitro Properties.
.
.
.
25
CHAPTER
1.
.
.
1.2.3. Structural Data for Vsr
1.3.
.
.
.
.
.
.
28
1.2.4. CatalyticMechanismof Vsr .
.
.
.
.
34
Methyl-Directed Mismatch Repair - Lessons From Streptococcus
Pneumoniae
1.4.
36
Methyl-Directed Mismatch Repair in E. coli.
.
.
.
37
1.4.1. Dam - in Vivo and in Vitro Properties.
.
.
.
38
1.4.2. MutS - in Vivo and in Vitro Properties
.
.
.
39
1.4.3. MutL - in Vivo and in Vitro Properties
.
.
.
41
1.4.4. MutH - in Vivo and in Vitro Properties
.
.
.
42
1.4.5. Mechanism of Methyl-Directed Mismatch Repair
.
.
44
1.5.
Mismatch Repair in Eukaryotes
.
.
.
45
1.6.
Interaction Between the VSPR and MMR Systems .
.
.
46
1.7.
Mechanism of VSPR .
1.8.
1.9.
CHAPTER2.
.
.
.
51
Repair of T:G Mismatches in Eukaryotes
.
.
.
.
52
References
.
.
.
.
69
.
.
.
.
.
.
.
.
.
Reconstitution of the E. coli Very Short Patch Repair System with
6
Purified Proteins
87
2.1.
Abstract
88
2.2.
Introduction
89
2.3.
Experimental Procedures
91
2.4.
Results.
97
2.5.
Discussion
101
2.6.
References
115
CHAPTER 3.
The Size of Repair Tracts Created by E. coli Very Short Patch Repair
System.
118
3.1.
Abstract
119
3.2.
Introduction
120
3.3.
Experimental Procedures
122
3.4.
Results.
127
3.5.
Discussion
130
3.6.
References
144
06-Methylguanine - Formation, Consequences, and Repair .
146
4.1.
06-Methylguanine - A Biologically Important Adduct
147
4.2.
06-Methylguanine is Mutagenic
148
4.3.
06-Methylguanine is Cytotoxic
152
4.4.
The Adaptive Response to Alkylating Agents
153
4.5.
Ada - in Vivo and in Vitro Properties .
156
4.6.
Ogt - in Vivo and in Vitro Properties
161
4.7.
Protection Against Alkylation Induced Toxicity by Ada and Ogt
163
4.8.
Repair of 06-Methylguanine by Nucleotide Excision Repair.
164
4.9.
References
172
CHAPTER4.
CHAPTER 5.
Effect of Mismatch Repair Proteins on Methyltransferase Repair of
06-Methylguanine in Vivo
186
5.1.
Abstract
187
5.2.
Introduction
188
5.3.
Experimental Procedures
191
5.4.
Results.
197
7
5.5.
Discussion
200
5.6.
Future Directions
203
5.7.
References
224
CHAPTER6.
Cloning, Expression, and Purification of Truncated Hormone
Receptors
229
6.1.
Introduction
230
6.2.
Nuclear Hormone Receptors .
230
6.3.
Experimental Procedures
232
6.4.
References
242
CHAPTER
7.
A Rationally Designed Genotoxin that Selectively Destroys Estrogen
Receptor-Positive Breast Cancer Cells
244
7.1.
Abstract
245
7.2.
Manuscript
246
7.3.
References
253
CHAPTER8.
Design, Synthesis, and Evaluation of Estradiol-linked Genotoxicants
as Anti-cancer Agents.
256
8.1.
Abstract
257
8.2.
Introduction
258
8.3.
Experimental Procedures
260
8.4.
Results and Discussion
261
8.5.
Conclusion
264
8.6.
Acknowledgments
265
8.7.
References
270
273
CURRICULUM VITAE
8
ABBREVIATIONS
ImA
1-methyladenine
2D-NMR
two dimensional nuclear magnetic resonance
3mA
3-methyladenine
3mC
3-methylcytosine
3mG
3-methylguanine
5mC
5-methylcytosine
7mA
7-methyladenine
7mG
7-methylguanine
Aa
amino acid
A
adenine
Amp
Ampicillin
AP
apurinic
AR
androgen receptor
ATP
adenosine triphosphate
BCNU
1,3-bis(2-chloroethyl)- 1-nitrosourea
BER
base excision repair
,3ME
beta-mercaptoethanol
bp
base pair
C
cytosine
Cam
Chloroamphenicol
CCNU
N-(2-chloroethyl)-\P-cyclohexyl-N-nitrosourea
Da
Dalton
DBD
DNA binding domain
DMS
dimethylsulphate
DMSO
dimethylsulfoxide
DNA
deoxyribonucleic acid
DTT
1,4-dithio-DL-threitol
E. coli
Escherichiacoli
EDTA
ethylenediaminetetraacetic acid
9
EMS
ethyl methanesulphonate
EMSA
electrophoretic mobility shift assay
ENU
N-ethylnitrosourea
ER
estrogen receptor
FAD
flavin adenine dinucleotide
g
gravity
G
guanine
GR
glucocorticoid receptor
h
hour
HEPES
4-(2-hydroxyethyl)piperazine- 1-ethanesulfonic acid
HTH
helix-turn-helix
IDL
insertion-deletion loop
Kb
kilo-base
kDa
kilo Daltons
Kan
Kanamycin
KOH
potassium hydroxide
LB
Luria Broth
LBD
ligand binding domain
M
molar
MCS
multiple cloning sites
MGMT
0 6 -methylguanine DNA methyltransferase
Mel
methyl iodide
MMR
methyl-directed mismatch repair
MMS
methyl methanesulphonate
MNNG
N-methyl-N' -nitro-N-nitrosoguanidine
MNU
N-methlynitrosourea
MTase
methyltransferase
NaCl
sodium chloride
NEB
New England Biolabs (Ipswich, MA)
NEN
New England Nuclear
NER
nucleotide excision repair
10
NHR
nuclear hormone receptor
NTD
N terminal domain
0 2mC
L-methylcytosine
&OmG
06-methylguanine
O2mT
&-methylthymine
O4mT
0 4 -methylthymine
ODN
oligodeoxynucleotide
PAGE
poly-acrylamide gel electrophoresis
PCNA
proliferating cell nuclear antigen
PCR
polymerase chain reaction
PNK
polynucleotide kinase
PR
progesterone receptor
RM
restriction modification
RT
room temperature
Rpm
revolutions per minute
SAM
S-adenosylmethionine
SDS
Sodium Dodecyl Sulfate
T
thymine
T4PNK
T4 polynucleotide kinase
TAE
Tris-Acetate-EDTA
TBE
Tris-Borate-EDTA
TE
Tris-EDTA
TLC
thin layer chromatography
Tris
2-amino-2-(hydroxymethyl)- 1,3-propanediol
U
uracil
UV
ultraviolet
VSPR
Very Short Patch Repair
W
adenine or thymine
Xgal
5-bromo-4-chloro-3-indolyl-beta-D-galactoside
11
LIST OF FIGURES
Figure 1.1.
Organization of the dcm and vsr genes
53
Figure 1.2.
Repair of mismatches that arise through methylation and subsequent
deamination of cytosine
54
Figure 1.3.
Unmethylated and hemimethylated Vsr substrates
55
Figure 1.4.
Proposed mechanisms for the hydrolytic deamination of cytidine
56
Figure 1.5.
Chemistry behind C:G to T:A transition mutations .
57
Figure 1.6.
Model for how VSPR could be shaping the E. coli genome .
58
Figure 1.7.
Structure of vsr bound to its product - minor groove side
59
Figure 1.8.
Structure of vsr bound to its product - major groove side
60
Figure 1.9.
Geometric characteristics of C:G and T:G base pairs.
61
.
Figure 1.10. Methyl direction of mismatch repair .
62
Figure 1.11. Methyl-directed mismatch repair is bidirectional
63
Figure 1.12. ATP-hydrolysis dependent translocation model for MMR
.
64
Figure 1.13. Molecular switch model for MMR, part 1
65
Figure 1.14. Molecular switch model for MMR, part 2
66
Figure 1.15. Molecular switch model for MMR, part 3
67
Figure 2.1.
Unmethylated and hemimethylated Vsr substrates
Figure 2.2.
PhosphorImager results from Vsr nicking assays on unmethylated,
hemimethylated, and control substrates
106
Graphical representation of unmethylated and hemimethylated
substrate nicking by Vsr
107
PhosphorImagery results from incubating T:G containing duplexes
with VSPR proteins
108
Graphical representation of Vsr nicking results from reconstitution
reaction conditions
109
Figure 2.3.
Figure 2.4.
Figure 2.5.
12
.
105
Figure 2.6.
Methodology for assessing VSPR efficiency in vitro.
110
Figure 2.7.
Representative PhosphorImagery results from TLC separation of
mononucleotides at interrogation site.
.
.
.
111
Figure 2.8.
Percentage of VSPR events initiated that went to completion
112
Figure 3.1.
Method for detecting VSPR by monitoring the incorporation of
radioactivity into substrate DNA
135
Extracts from cells over-expressing recombinant Vsr incorporate
[a32P]-dCTP into short duplex substrates via VSPR .
136
Graphical representation of results from measuring VSPR via
radioactivity incorporation
137
Method for detecting VSPR by measuring the depletion of
radioactivity signal from substrate DNA
138
Figure 3.5.
Distribution of VSPR patch lengths created in vitro .
139
Figure 3.6.
Addition of surplus Ligase to VSPR reactions
140
Figure 3.7.
Addition of Ligase to VSPR reactions containing excess substrate .
141
Figure 4.1.
Sites of methylation on the DNA bases
165
Figure 4.2.
Conversion of MNNG to the active methanediazonium ion by
Figure 3.2.
Figure 3.3.
Figure 3.4.
cysteine
166
Figure 4.3.
The R and S stereoisomer of methylphosphotriesters.
Figure 4.4.
Base pairing models for O6mG:T and O6mG:C
168
Figure 4.5.
Genetic structure and regulation of the adaptive response in E. coli .
169
Figure 4.6.
Direct reversal of base damage by Ada
170
Figure 5.1.
Sequence of ogt and location of PCR primers
206
Figure 5.2.
Sequence of ada/alkB and location of PCR primers .
207
Figure 5.3.
Sequence of mutHand location of PCR primers
208
Figure 5.4.
Sequence of mutL and location of PCR primers
209
13
.
.
167
Figure 5.5.
Sequence of mutS and location of PCR primers
210
Figure 5.6.
Sequence of vsr and location of PCR primers
211
Figure 5.7.
Relative efficiencies of MTase repair in MR defective strains
212
Figure 5.8.
PCR mediated gene replacement
213
Figure 5.9.
In vivo assay for MTase repair
214
Figure 5.10.
Hypothetical mechanism for stimulation of MTase repair
by VSPR proteins
215
Figure 5.11.
Bacterial two-hybrid system utilizing domains of adenylate cyclase.
216
Figure 6.1.
Model for mechanism of bifunctional toxicants
236
Figure 6.2.
Restriction map of pSK278
237
Figure 6.3.
Restriction map of pSK278-hER-LBD
238
Figure 6.4.
Restriction map of pSK278-hAR-LBD
239
Figure 6.5.
Restriction map of pSK278-hPR-LBD
240
Figure 6.6.
Restriction map of pSK278-hGR-LBD
241
Figure 7.1.
(A) Piperidine treatment of 5'-d(AATATTGGCCAATATT) treated
with compound 1. (B) Retarded mobility of oligonucleotide-1
in the presence of ER-LBD illustrated by EMSA
251
Survival of MCF-7 (ER+) and MD-MB231 (ER-) cells treated with
1 or chlorambucil
252
Figure 8.1.
Structure and molecular features of lead compound .
266
Figure 8.2.
Survival of MCF-7 (ER+) and MDA-MB231 (ER-) breast cancer
cells after 2 h exposure to estradiol-linked toxicants .
269
Figure 7.2.
14
LIST OF TABLES
Table 1.1.
E. coli and S. cerevisiae Proteins Required for MMR
Table 2.1.
Oligodeoxynucleotides used for cloning Vsr into pET28a and
construction of nicking assay and reconstitution assay substrates
113
Quantitative results from reconstitution assays containing
Vsr, Pol I, and DNA Ligase protein .
114
Table 2.2.
68
Table 3.1.
Oligodeoxynucleotides used to construct duplexes containing internal
radiolabels at prescribed distances away from the mismatched T
142
Table 3.2.
DNA strands containing radiolabels at prescribed distances
away from the T to be mismatched
143
Relative amounts of DNA alkylated products following reaction of
double stranded DNA with MMS and MNU alkylating agents
171
Table 5.1.
PCR primers used for construction of recombination substrates
217
Table 5.2.
PCR primers used for analyzing strain genotypes
218
Table 5.3.
Sizes of genes and PCR amplification products
219
Table 5.4.
Initial plating results from MTase assay on C215 derivatives
220
Table 5.5.
Initial plating results from MTase assay on C216 derivatives
221
Table 5.6.
Initial plating results from MTase assay on C217 derivatives
222
Table 5.7.
Initial plating results from MTase assay on C218 derivatives
223
Table 8.1.
RBA= relative binding affinity for the rabbit uterine ER as compared
to estradiol (RBA=100)
267
Table 4.1.
15
LIST OF SCHEMES
Scheme 7.1.
Synthesis of compound 1
250
Scheme 8.1.
Synthesis of estradiol-linked aniline mustards
268
16
CHAPTER
1
Mismatch Repair in Escherichia coli
17
-ChapterI 1.1. MUTATION AVOIDANCE
The formation of a base pair other than the canonical Watson-Crick G:C or A:T
by a polymerase error, or by a chemical or hydrolytic reaction of an existing base pair,
will create a mismatch and lead to a mutation if not repaired.
Systems for mutation
avoidance are present in both prokaryotic and eukaryotic cells (1-15). These mismatch
repair (MR) systems are highly conserved from bacteria to humans and play an essential
role in maintaining the integrity of the genetic code and health of the host. Cells deficient
in MR exhibit a mutator phenotype, in which the spontaneous mutation rate is increased,
and the cells are hyper-recombinogenic.
In mammals, loss of MR is related to an
increased susceptibility to cancer (6,;16). Understanding mismatch avoidance is therefore
paramount to human health.
Mismatched base pairs can arise in DNA through multiple processes (17). Errors
in replication can cause an incorrect deoxynucleotide to be incorporated into the newly
synthesized strand. In fact, in Escherichia coli (E. col), MR contributes almost 1000fold to the fidelity of DNA replication (4;18). Though, under normal conditions of
replication, DNA polymerase III is responsible for proper base selection and insertion
(19) and has associated proofreading activity (20), mistakes are made with a frequency of
10-9 to 10-1°0 (21). Additionally, slippage during replication can result in the formation of
insertion-deletion loops (IDLs), which are composed of bases that are not matched at all.
IDLs of less than four nucleotides have been shown to be good substrates for MR (2224). IDLs of up to 4 or 5 bases can also be repaired by MR, albeit at lower efficiencies
(25). In contrast, heteroduplexes containing large regions of nonhomology are not MR
substrates (26). Recombination between two DNA strands that do not share complete
sequence homology can also result in the formation of IDLs and mismatches that are
subject to MR (27;28). Base pairs containing a chemically modified nucleobases can
also substrates for MR (29). In particular, one mismatched base pair that can arise in
DNA naturally is through the deamination of 5-methylcytosine (5mC). Small amounts of
5mC are present in both prokaryotic (30) and eukaryotic (31) DNA, and the increased
propensity of 5mC to hydrolytically deaminate elevates the level of T:G mismatches.
18
- Chapter1 -
The next sections will describe two mismatch repair pathways in E. coli, giving
particular attention to the very short patch repair (VSPR) pathway and the cellular
components it shares with the methyl-directed mismatch repair (MMR) pathway.
1.2. THE VERY SHORT PATCH REPAIR SYSTEM
In E. coli, the underlined cytosine of the pentanucleotide sequence 5'- CCWGG 3' (where W is an A or T) is methylated at the 5-position by a DNA cytosine
methyltransferase (32;33) which is the protein product of the dcm gene (34). The 5methylcytosines (5mC) present in Dcm methylation recognition sites are prone to
spontaneous deamination, which results in a T:G mismatch (35;36). If left unrepaired, a
C:G -- T:A transition mutation will arise following the next round of DNA replication.
To avoid the formation of these mutations, the very short patch repair system removes
and replaces the mismatched T with the proper base (37).
The first indication that such a system existed was observed in 1981 by Margaret
Lieb when, in multifactor crosses, the recombination frequencies of certain amber
mutations in the repressor (cI) gene of bacteriophage Xwere higher than predicted by the
physical distance separating them (38). Further study of the am6 mutation revealed that
these events arose from C -+ T transition mutations in a glutamine codon (5'- CAG -3')
located within the Dcm recognition sequence 5'- CCAGG -3' (39), and recombinants had
the C:G genotype (37;40). Additionally, Lieb demonstrated that a high frequency of
these unexpected recombinants included co-repair of markers within 10 nucleotides of
the mismatch but not in areas further than 20 nucleotides (39). Taken together, these
findings led Lieb to define a very short repair (VSPR) system that acts on the sequence
5'- CTWGG -3' (where T is mismatched with G) to preferentially restore it to 5'CCWGG -3'.
The significance of the 5'- CCWGG -3' sequence, being that for Dcm
recognition and methylation, was not overlooked.
19
- Chapter1 Additional evidence for the existence of VSPR and a proposed role in maintaining
Dcm methylation sites came from experiments with heteroduplex DNA (41-43).
Substrate containing a T:G mismatch in the sequence 5'- CTAGG -3', which could result
from the deamination of the Dcm product, was indeed subject to repair with a high (if not
complete) bias in favor of replacing the mismatched T. Increasingly, it was accepted that
VSPR functioned to reduce the mutation level associated with the spontaneous
deamination of 5mC created by Dcm.
This hypothesis was corroborated by Lieb in 1991 when she observed that
mutation at a particular hot spot was dependent upon Dcm and VSPR activity (44) In
lysogens where only Dcm was present, the mutation frequency at the hot spot was 10fold higher than in wt bacteria. This hot spot disappeared when both Dcm and Vsr were
removed, while addition of Vsr to wild type (wt) bacteria caused a 4-fold decrease in
mutation frequency. It is now accepted that a primary role of VSPR is to reduce the
number of mutations that result from cytosine methylation and deamination (45).
In the years following the discovery of the VSPR system, a number of genetic and
biochemical studies were conducted to implicate the proteins involved in the repair
process. In 1987, Lieb reported (46) that the protein products of mismatch repair genes
mutS and mutL were involved in VSPR on the basis that the frequency of in vivo crosses
in cells lacking either one of these genes was greatly diminished. Strains lacking mutHor
uvrD exhibited no difference from wt. Interestingly, the recombination frequency was
diminished in dcm6 cells lacking the Dcm methylase, leading Lieb to conclude that the
protein product of dcm had a repair function in addition to its methylase function. At the
time, however, there was no explanation for this mixed phenotype.
In a report by Dzidic and Radman in 1989 (47) the genetic requirements for
hyper-recombination by VSPR were extended to include polA (the gene for Pol I). In
fact, the recombination frequency was diminished in cells lacking the polA gene
sequences for either its 5' --+ 3' exonuclease or polymerase domains. This observation
20
- ChapterI led the authors to conclude that both activities of Pol I were paramount for effective
VSPR. In contrast, xth-, nth- and nfo- strains showed wt recombination frequencies.
In 1990, Sohail reported (48) that the two phenotypes of the dcm mutants, lacking
both methylation and repair activities, had been genetically pinpointed by mapping.
Additionally, they cloned genes that complemented the chromosomal mutation for
methylation, but not repair, and vice versa. The gene essential for methylation, dcm, was
predicted to code for a 473-amino-acid protein and was not required for VSPR, while the
second gene, vsr, was predicted to code for a 156-amino-acid protein which was required
for VSPR but not for methylation. The two genes were transcribed from a common
promoter even though they were found in different translational registers. The 5' end of
vsr overlaps the 3' end of dcm by 7 codons in a +1 reading frame (Figure 1.1). It was
hypothesized that this genetic arrangement assures Vsr is always produced along with
Dcm to minimize the mutagenic propensity of cytosine methylation. Sohail also noted
(48) that although the two genes are transcribed from a common promoter, two separate
proteins are produced. Thus, the genetic arrangement of dcm and vsr was the source of
confusion regarding initial experiments conducted on the dcm6 strain, as they lack vsr in
addition to dcm. This misunderstanding led to the incorrect inference (46) that the dcm
product is required for VSPR.
One year later, Hennecke showed that the vsr gene product was an 18 kDa
mismatch endonuclease that produced a single strand nick 5' to the underlined T within
duplex DNA sequences 5'- CTWGN -3' or 5'- NTWGG -3' (where N is A, T, C, or G)
when the T was opposite a G (49). The incision was mismatch-dependent and strandspecific. These results therefore illustrated that Vsr was the protein that initiates VSPR.
Consistent with these observations, strains lacking vsr are completely defective in VSPR
and have a very high frequency of C -- T transitions at 5mC sites (38;50).
Altogether, these experiments provided a model for the mechanism of VSPR
(Figure 1.2). Evidence suggested that Vsr initiates repair by producing a nick 5' to the
mismatched T. The previous implication of MutS and MutL in mismatch recognition
21
- ChapterI suggested these proteins stimulated VSPR by assisting Vsr in finding or incising substrate
mismatches. The exact mechanism of Vsr stimulation, however, is still unknown (see
below). Subsequent to incision, Pol I removes the T residue with its 5' -, 3' exonuclease
activity and commences repair synthesis in the usual template-directed fashion, with the
short synthesis tracts typical for the enzyme (51). DNA Ligase is reasonably assumed
seal the nicked strand on which repair has been performed.
1.2.1. DCM- IN VIVOAND IN VITROPROPERTIES
Dcm is the 53 kDa product of dcm, which is located at 43 minutes on the E. coli
chromosome (34).
The protein methylates the underlined cytosine residue in the
sequence 5'- CCWGG -3' (where W is A or T) at the C5 position of the pyrimidine ring
(32). The recognition sequence is palindromic and thus also results in methylation at the
corresponding position of the opposite strand, as well (32). The deamination of a 5mC in
fully dcm methylated DNA gives rise to a T:G mismatch in a hemimethylated duplex. If
the G containing strand contains an unmethylated C, the mismatched duplex is said to be
unmethylated (Figure 1.3). In E. coli, approximately 1% of all cytosine residues are
methylated by Dcm (52). This translates roughly into approximately 10,000 Dcm
recognition sites in the E. coli genome, or on average one recognition site every 400-500
base pairs.
Not surprisingly, the role of Dcm has been a topic of much discussion. However,
no obvious biologically interesting phenotype of dcm mutants has been reported (52), and
the reason for the existence of the dcm-vsr operon remains unclear (53). The closest
characteristic to a phenotype for dcm mutants is the ability of their isolated genomic
DNA to be cut by EcoRII endonuclease, which is part of a restriction modification (RM)
system and
recognizes
the same sequence
as Dcm
(54;55).
The EcoRII
methyltransferase and Dcm share a high degree of sequence similarity (48), are
functionally very similar, and are likely derived from a common ancestor.
22
- ChapterI An EcoRII RM system was found in drug resistance transfer plasmids (56), and
therefore the function of Dcm could be to protect the host's DNA from invasion (53). In
support of this theory, cells bearing Dcm were shown to be resistant to the virulence of a
parasitic EcoRII gene complex (53). It is also interesting to note that 5'- CCWGG -3' is
one of the most commonly utilized recognition sequences for RM systems (53). As of
March 20th , 2006, the REBASE database (57) identified 169 of the total 3773 RM
systems as recognizing the 5'- CCWGG -3' sequence.
It has also been suggested that methylation by Dcm could be involved in gene
regulation (52). A Dcm methylation site has been identified in the promoter region for
lexA (58), however no relationship between the Dcm methylation status and expression
has been established. Interestingly, some Dcm sites were found to remain unmethylated
in dcm+cells (59), suggesting that some sites are protected by DNA binding molecules or
by DNA topology, and could play a role in a yet to be discovered biological process.
The resultant 5mC residues are subject to "spontaneous" hydrolytic deamination
(60;61), analogous to the process that converts cytosine to uracil (62). Two chemical
mechanisms have been proposed for the hydrolytic deamination of cytosine (62;63). One
mechanism involves direct attack at the 4-position of the pyrimidine ring by a hydroxyl
ion, and the other involves an addition-elimination reaction with the formation of
dihydrocytosine as an intermediate (Figure 1.4). While the mechanism involving direct
attack has the advantage of simplicity, relatively little direct evidence has been reported
to support its occurrence in reactions involving nucleophilic displacement of the amino
group of cytosine. On the other hand, the route involving addition to the C=C double
bond is similar to that established for the reaction between hydroxylamine and cytosine
derivatives (64). Additionally, it has been directly demonstrated that nucleophiles can
add reversibly to the 5,6 double bond of a uracil ring (65). The deamination of
dihydrocytosine derivatives (resembling intermediate 2 in Figure 1.4) was also shown to
be a rapid process (66). Therefore, evidence suggests that an addition-elimination
reaction is a probable mechanism for deamination of cytosine and 5mC in DNA.
23
- Chapter
-
Deamination of either cytosine or 5mC gives rise to products that yield C:G -T:A transition mutations upon the next round of replication (Figure 1.5).
It is
noteworthy, however, that the extent of cytosine and 5mC deamination is not equivalent.
The higher rate of deamination for 5mC as compared to C (60;61), coupled to the finding
that T:G mismatches are repaired less efficiently than U:G mispairs (67) results in
cytosine methylation sequences being hot spots for C -, T mutations. Interestingly, the
accumulation of these hotspots in E. coli, depends on the phase of growth.
In exponentially growing cells (67), VSPR reduced the 5mC
-
T mutations by a
factor of four. However, cells deficient in Dcm displayed more than 10-fold decrease in
the 5mC -- T mutation level. Therefore, the inefficiency of VSPR during exponential
growth allows Dcm sequences to be hotspots for C -+ T mutations. In contrast, cells in
stationary phase (68) are completely protected by VSPR from Dcm sequences being
hotspots. It was observed that the 5mC -
T mutation rate in cells that were stored for
long periods of time was equivalent to that for cells lacking cytosine methylation. It was
also observed that the 5mC
-
T mutation rate for cells lacking VSPR was consistent
with the rate of 5mC deamination at 37 C.
Subsequently, it was shown that the
concentration of Vsr in the cell is low during exponential phase and increases when cells
start to enter stationary phase (69). To summarize, the ability of VSPR to protect against
5mC
-+
T mutations is linked to the amount of Vsr that is available to initiate repair.
Possible explanations for how the amount of Vsr is controlled in the cell (70;71), and
why VSPR was designed to be less efficient in exponential phase have been proposed
(72), and are discussed below.
In eukaryotes, methylation of cytosine occurs within CpG dinucleotides (73), and
has been implicated, among other functions, in the regulation of gene expression (74). A
large number of genetic diseases involve C to T changes at CpG sites (75), including 13thalassemia, hemophilia, and cancer (76-79), therefore establishing the maintenance of
the sites as critical.
24
- Chapter1 1.2.2. VSR- IN VIVO AND IN VITRO PROPERTIES
Vsr is the 18 kDa product of vsr, located at 43 minutes on the E. coli chromosome
(48). This 156 amino acid protein functions as a strand-specific endonuclease to initiate
repair of T:G mismatches in the sequence 5'- CTWGG -3' (where T is mismatched with
a G) (49). These T:G mismatches arise through the methylation of the underlined C in
the sequence 5'- CCWGG -3' by Dcm, followed by deamination of that methylated
residue.
Vsr, therefore, helps to reduce the mutation level associated with the
spontaneous deamination of 5mC created by Dcm.
In 1995, Lieb and Rehmat (80) reported recombination experiments conducted on
derivatives of Dcm sequences. Their findings showed that VSPR was most efficient in
the pentanucleotide sequence 5'- CTAGG -3', however there was also significant
correction when either the 5' most C or the 3' most G was replaced by another base. In
fact, some VSPR activity was observed in all of the sequences tested.
The same year, Glasner confirmed the broad sequence specificity of Vsr
biochemically (81). Using fluorescence-labeled oligodeoxynucleotide (ODN) substrates,
14 substrates were found to be processed by the enzyme, with varying efficiencies. The
substrates differed at one or two positions from the canonical pentanucleotide substrate
sequence 5'- CTWGG -3'.
Not surprisingly, the sequences 5'- CTAGG -3' and 5'-
CTTGG -3' were the most reactive substrates with relative rates of 100 and 68%
respectively. Sequence derivatives where the first or last base was varied (5'- NTWGG 3' or 5'- CTWGN -3') were all repaired with relative rates of 22% or higher. Therefore,
Vsr will act efficiently on T:G mismatches in many sequence contexts.
This same study (81) investigated potential relationship between the efficiency of
VSPR on individual sequences and their underrepresentation in the genome.
It was
previously hypothesized (82;83) that if a T:G mismatch resulted from misincorporation
of a G opposite a T in the template DNA strand, it would be a potential substrate for both
the methyl directed mismatch repair MMR and the VSPR systems. Further, if such a
25
- Chapter1 mismatch evaded correct repair by MMR, VSPR might incorrectly carry out "repair" by
removing and replacing the T. Therefore, over time, VSPR could operate on mismatches
not intended for the system and would provide a driving force in depleting the genome of
the T-containing sequences (for example 5'- CTAG -3') and enriching it for the
corresponding C-containing sequences (for example 5'- CCAG -3'). Ultimately, VSPR
could have a hand in modifying the E. coli genome (Figure 1.6).
Interestingly, Glasner's data (81) reflected a high quality of positive correlation
between the relative second-order rate constants on VSPR removal of mismatched T and
the underrepresentation rank for the 14 substrates studied. It was hypothesized that this
process would lead to corresponding sequence polymorphisms in the total E. coli K-12
population. In a case where two genomes that differ in one or more such polymorphic
sites combine, the T:G mismatches that are substrates for Vsr would be repaired in favor
of G and thereby transfer the polymorphism to the other genome involved. Additionally,
because VSPR replaces a short stretch of DNA downstream of the T:G mismatch, any
mismatches within the reach of the DNA synthesis tract would also be repaired in favor
of the G containing strand. In conclusion, Glasner postulated that VSPR might assist in
the patchwise unidirectional transfer of genetic information from one DNA molecule to
another, and contribute to the overall fitness of E. coli K- 12.
Another function for VSPR appears to be to repair U:G mismatches in Dcm
sequences that arise by deamination of cytosine (84). It was found that deamination of
cytosine can be promoted by the Dcm methylase.
In the absence of S-
adenosylmethionine, DNA cytosine methylases promote the formation of a 5-dihydro
intermediate that is much more susceptible to deamination than cytosine (85-88).
Normally, uracils that arise by the deamination of C are recognized by uracil DNA
glycosylase and are repaired by the base excision repair pathway. However, genetic
reversion assays (84) revealed that although VSPR is not as efficient as the glycosylase
system at correcting U:G mismatches to C:G, the VSPR system does contribute to the
correction of U:G mismatches in Dcm sites.
26
- Chapter
-
The ability of Vsr to nick 5' to U:G mismatches in Dcm sequences was also
studied biochemically (89). Results indicated that the single turnover rate for the U:G
mismatch was about half that for the T:G mismatch. These biochemical parameters,
however, were determined using protein that was not fully active. Turner observed that
much of the Vsr purified from over-expressing E. coli strains is catalytically inactive, and
less than 1% of the protein bound DNA (89). In fact, it was not until direct titration
experiments were conducted (89;90), in which increasing amounts of Vsr were added to a
constant amount of substrate DNA, that the activity of an individual protein preparation
could be determined. Ultimately, the authors concluded that the U:G mismatch had a
specificity constant (kst/KD) 10-fold lower than that for a T:G mismatch, due to an
increase in equilibrium dissociation constant (KD) and a reduction in the rate constant for
single turnover (kst). Similarly, the specificity constant for a T:G mismatch in an
unmethylated duplex (Figure 1.3) was 10-fold less than that for a T:G mismatch in a
hemimethylated duplex (Figure 1.3). Crystallographic data containing Vsr bound to
duplex DNA (91;92) later suggested that the methyl group present in the hemimethylated
duplex may pack against an aliphatic region of the protein (Phe-77, Lys-78, Val-79), and
assist binding at the protein DNA interface. These findings support the notion that the
biological role of Vsr is to initiate repair of T:G mismatches that arise from deamination
of the Dcm product. Even so, the ability of Vsr to incise unmethylated substrates has also
been confirmed in vivo (46).
To obtain the highest activity from Vsr protein, it has been reported (93) that
current preparations should include freshly transforming E. coli with the plasmid that
directs Vsr over-expression, purifying Vsr from the over-expressing strains, and using the
purified protein within two weeks of preparation. Furthermore, loss of Vsr activity due to
aggregation of protein during storage has been reported (94). Therefore, for best results,
Vsr protein must be prepared fresh each time. While many changes have been made to
the current methods for preparing Vsr, the reasons for the discrepancy in activity are not
understood. Perhaps, as discussed below, the low activity of some Vsr preparations can
be attributed to the loss of 14 residues from N-terminus of Vsr upon storage. Whatever
27
- Chapter
-
the case, adhering to these precautions has allowed fully active Vsr to be isolated and
used to measure the binding and kinetic constants for its interaction with substrate DNA.
The binding affinity of Vsr for its hemimethylated substrate (Figure 1.3) was
measured using competition gel-retardation assays (93). Complexes of Vsr and a 32p_
labeled duplex were formed (in the presence of Ca2 +) and increasing amounts of
unlabelled competitor DNA were added. Utilizing these methods, a KD of 10.9 nM was
determined. Single turnover reaction conditions were utilized to saturate the substrate
and reflect rates of turnover after substrate binding yet before product release. Under
these conditions, the kst was 7.5 min ' . For the unmethylated substrate (Figure 1.3), a kst
of 2.8 min -' for Vsr was independently reported (94). A comparison of the kinetic
measurements between the hemimethylated and unmethylated substrates suggests that the
activity of Vsr is roughly 2.5-fold more efficient on the hemimethylated substrate. The
kinetics of Vsr on these two substrates, however, have never been tested in parallel with
fully active protein.
Determination of both KD and kt values (93) allowed the computation of the
single turnover specificity constant (kst/KD),which was 0.7 min -' nM -1 for Vsr. This
constant is generally agreed to be the most suitable metric for comparing the ability of
different proteins to bind DNA substrates (95-97). In reactions where the steady state
rate of hydrolysis is measured, the specificity constants oftentimes reflect the product
release step as this step is commonly rate limiting.
In such cases, the steady state
constants are not particularly informative for comparisons. In light of the observation
that Vsr produces a very stable enzyme product complex (89), the single turnover
constants are indeed more useful than the steady state values.
1.2.3. STRUCTURAL DATA FOR VSR
To date, three structures of Vsr have been solved by X-ray diffraction and are
available on the RCSB Protein Data Bank. The first structure, IVSR, (98) was solved to
28
- Chapter1 1.80 A and contains residues 21 to 156. The second structure, 1CWO, (91) was solved to
2.30 A and contains residues 2 to 156 of the Vsr protein bound to product duplex DNA.
The third structure, 10ODG,(92) was solved to 2.80 A and contains residues 22-155 of the
Vsr protein bound to its reaction product site in duplex DNA. Both of the Vsr-DNA
complexes contain hydrolyzed substrate. The following discussion is derived from the
data presented in these three publications, and will go into the finer details of the
structures and what can be learned upon their inspection.
Vsr has a three-layered a3/la fold that is stabilized by a structural zinc site (98).
The Zn2+ is coordinated by four protein residues (Cys-66, His-71, Cys-73, Cys-117) in a
tetrahedral arrangement, three of which are on one loop. This coordination is distinct
from the majority of zinc sites found in proteins.
Typically, the four coordinating
residues are divided equally on two strands (99). While Vsr shares no sequence
homology with other endonucleases (49), the overall topology of Vsr resembles the type
II restriction endonucleases, which have been extensively reviewed elsewhere
(100).
Members of this family include a growing list of restriction enzymes, as well as the DNA
repair nucleases MutH (101), X exonuclease (102), and archeal Holliday junction
resolvase (103). However, more specific comparison reveals that Vsr is not a standard
type II endonuclease, as previously expected.
The central f3 sheet of Vsr is composed of two short and four long P strands, and
braced on either side by a-helicies.
The active site of this type II restriction
endonuclease family is located in a niche formed in the
catalytic motif PDX( 6_30)(D/E)XK (104;105).
sheet and is composed of the
For this family of enzymes, these
conserved residues are superimposable from enzyme to enzyme and are essential for
catalytic activity.
Typically, the first two residues are acidic and bind an essential
divalent cation. The last residue, typically Lys, is the most variant amino acid between
enzymes, and has been proposed to stabilize the transition state (106;107) or orient the
attacking water (108;109). Superimposition of the 13sheet of Vsr onto members of the
type II restriction enzyme family shows that Asp-51 (D) of Vsr (a catalytically essential
residue) is the first conserved Asp of the catalytic motif. In keeping with the motif, this
29
-Chapter
Iresidue does bind an essential divalent cation. The rest of the catalytic motif for type II
restriction enzymes, however, is not conserved in Vsr.
Phe-62 in Vsr superimposes onto the second catalytic residue of the motif,
however this residue can not engage in metal coordination. The third residue in the motif
correlates to a partially conserved His or Asp 64 in Vsr, and while this residue is
important for activity, it is not essential. Additionally, His 69, which is close in vicinity
to His 64 but positionally distinct from the catalytic residues of type II restriction
enzymes, was found to be absolutely required for Vsr activity.
These findings have
disqualified Vsr from strict conservation with the type II restriction enzyme catalytic
motif, and imply that the mechanism for Vsr differs from that of typical type II restriction
enzymes. These structural data helped to clarify the discrepancies between how the motif
is used for typical type II restriction enzymes and Vsr.
In contrast to typical type II restriction enzymes which utilize highly coordinated
metal ions, Vsr coordinates two Mg 2+ ions through extensive metal water clusters and has
very little direct coordination with the Mg2+ ions. Besides Asp-51, the only other direct
metal coordination includes the main chain carbonyl of Thr-63, one residue away from
the predicted Phe-62. His-64 and Glu-25 coordinate water molecules in the water metal
clusters. Alanine scanning has shown each of these residues to be important for Vsr
activity. The crystal structure also revealed that the product DNA duplex, containing a 5'
phosphate on the mismatched thymine and a 3' hydroxyl on the neighboring cytosine,
was involved in Mg2 + coordination. The oxygen on the C3' of the cytosine coordinates
one metal ion.
Additionally, the 5' phosphate of the Vsr reaction product was
coordinated with both metal ions. Interestingly, His-69 was coordinating one of the
terminal phosphate oxygens. This coordination is atypical since Mg2 + generally prefers
the harder oxygen atoms of carboxyl residues as ligands.
In addition to the structural involvement of zinc, Vsr requires Ca2+ or Mg2+ for
substrate binding (90). This observation is in contrast to a number of restriction enzymes
including EcoRI and BamHI, which require metal ions for catalysis but not for binding
30
- Chapter1 (100). This distinction implies that the metal ions in Vsr play a structural role in addition
to a catalytic role. Consistent with this idea, it was observed that Vsr can bind substrate,
but not cleave it, in the presence of Ca2+ (89). This property allows the binding and
cleavage activities of Vsr to be separated, and has been useful for a number of
biochemical studies (89;90;93).
In conclusion, the first structure of the Vsr protein alone revealed many important
features of the molecule including its topology in relation to type II restriction enzymes,
certain conserved elements in its catalytic site, its novel metal binding coordination, and
why certain residues are important or critical to catalysis. The second structure of Vsr
contained the protein bound to duplex DNA product, and provided valuable information
on how the two molecules interact.
The N-terminus of Vsr, which was partially removed in the truncated structure,
does not form a single alpha helix as was predicted by secondary structure analysis of the
primary sequence (91). Rather, the N-terminus is composed of two short alpha helices
separated by a flexible linker (Figure 1.7). These results are in good agreement with
limited proteolysis studies which suggested that Vsr had exposed turn-arounds at amino
acids 14 and 20 (91;98). The main core of Vsr binds the DNA duplex from the major
groove while the most N-terminal helix (Lys-7 to Arg 15) pins down the DNA from the
minor groove side. A large opening has been formed in the center of the DNA duplex via
intercalation of multiple side chains between the DNA strands.
The protein surface
shows a strong electrostatic and steric complementarity to the DNA duplex. The basic
regions follow along the phosphate backbone of the DNA and a strong acidic patch is
found close to the metal binding sites in the catalytic center. Met-14 and Ile-17 project
from the minor groove side and leave only enough space for the phosphate backbone to
pass on either side.
Remarkably, three aromatic residues (Phe-67, Trp-68, and Trp 86) insert into the
DNA duplex from the major groove side (Figure 1.8). Phe-67 stacks with the A:T base
pair in the recognition sequence, Trp-68 stacks with the mispaired T, and Trp-86 stacks
with two sugars on the strand opposite the mismatched T. The major groove is opened
31
- Chapter
-
up, the DNA is significantly unwound, and the duplex is kinked by approximately 44
degrees. It is surmised then that these residues recognize and enhance the structural
distortion in DNA resulting from the T:G mismatch (99). Not surprisingly, these three
residues are invariantly conserved in the Vsr family (91).
Two of the intercalating residues (Phe-67 and Trp-68), are adjacent to His-69,
which is absolutely required for activity and proposed to activate the attacking water.
Additionally, all these residues are part of the loop that contains three (Cys-66, His-71,
and Cys-73) of the four residues that coordinate the structural Zn2+ ion. Therefore, these
residues are constrained conformationally and precisely aligned for mismatch recognition
and DNA strand incision.
Vsr recognizes its mismatch within the DNA helix, primarily by the structural
distortion in DNA caused by wobble pairing between T and G (Figure 1.9). In a T:G
wobble pair, the 06 and N1 of guanine hydrogen bond to the N3 and the 02 of thymine,
respectively.
This pairing is preferred by Vsr for a number of reasons.
Firstly, the
geometry of a T:G mispair is such that it can enter the active site of Vsr. A normal G:C
base pair would encounter steric clash between the O2 of cytosine and Thr- 19. However,
because the 02 of T in a T:G mispair is pushed into the major groove, these mismatches
can fit into the Vsr active site. Secondly, the wobble pairing achieves a shape such that
the scissile phosphate of the mismatched thymine is in proximity to the catalytic residues
in the active site. In other words, normal base pairs would be situated such that their
phosphate group would not be proximal to the Vsr catalytic residues. Thirdly, the wobble
pairing disrupts the DNA stacking between the mismatched G and the base on the 5' side
such that it causes (1) an opening of the DNA by unwinding, (2) increased distance
between G and its 5' neighbor, and (3) the ability of Vsr to recognize and bind the G:T
mismatch by intercalating within the recognition site. In addition to the ability of Vsr to
recognize T:G mismatches via the drastically different geometry, there are a few
hydrogen bonding contacts between the mismatched base pair and Vsr residues.
The
invariant Asn-93 binds to the 04 of thymine and presumably plays a role in cytosine
discrimination. Guanine is bound at its 06 position by Lys-89 and at its N\ by the main
32
- Chapter1 chain of Met-14. Biochemical analysis of mutants in these residues will be necessary to
determine the extent to which these interactions are required for recognition.
The five base pair recognition sequence occurs through an interlaced network of
direct and water mediated hydrogen bonding interactions. Two notable features are that
(1) Arg-120 has the only direct contact with the major groove, and it binds to the G:C
base pair that is required for recognition, and (2) the third base pair in the middle of the
recognition sequence (which can be A:T or T:A) has the most interactions. The sequence
specificity of Vsr may therefore lie primarily in the ability of duplex DNA to tolerate
intercalation.
The third structure of Vsr was obtained using a DNA duplex that generated a site
corresponding to the hemimethylated cleavage product of Vsr (92).
This second
protein/DNA structure is quite similar to the co-crystal structure previously reported (91),
but some differences are worth mentioning. The structure shows a 60° angle along the
minor groove as compared 44 ° in the previous structure solved by Tsutakawa.
Additionally, the middle A:T base pair (in the sequence 5'- CTAGG -3') was in a
Hoogsteen conformation, with the A being in the syn conformation. This pairing is
believed to exist in bulk DNA at low levels (in equilibrium with the Watson-Crick form)
but has not, until recently, been observed in a protein-DNA complex (110). To explain
why this Hoogsteen pairing was first observed in this structure, it is relevant to note that,
in this structure, Vsr is bound to a hemimethylated DNA substrate. The methyl group on
the C5 position of C would potentially clash with the C5 methyl group of T if the
thymine residue was not positioned closer to the minor groove through a Hoogsteen pair.
Therefore, it was hypothesized that, in this structure, the Hoogsteen A:T base pair is more
energetically favorable than a normal A:T base pair that would introduce methyl-methyl
clash.
33
- Chapter
I1.2.4. CATALYTIC MECHANISM OF VSR
Taken together, these structural data (91;98) lead Tsutakawa to propose a
catalytic mechanism for Vsr (99). When Vsr binds the mismatch-containing DNA
duplex, the scissile phosphate is coordinated and stabilized by both metal water clusters.
His-69 abstracts a proton from one of the waters in the metal water clusters and the
resulting activated water attacks the phosphate group between the mismatched thymine
and its 5' cytosine neighbor.
The orientation of the His-69, terminal phosphate, and
deoxyribose oxygen groups suggests an inline attack. The lack of significant side chain
conservation in the area surrounding the deoxyribose oxygen suggests that its protonation
is from a water molecule and not the protein.
Additional support for His-69 acting as a catalytic base comes from the
observation that replacement of this residue with alanine results in a variant with
dramatically reduced activity (91;98). Additionally, a hydrogen bond between His-69
and Asp-97 would increase the pKa of His-69, facilitating its role as a base (99). Several
endonucleases including DNaseI (111), exonuclease III (112), and HAP1 (113) use a
similar Asp-His pair to provide basic catalysis.
The two magnesium water clusters are also probably required for stabilizing the
additional negative change that develops on the scissile phosphate in the transition state.
It has also been hypothesized that one of the magnesium ions may have a role in correctly
aligning the activated water molecule for attack, while the other magnesium ion may
provide Lewis acid catalysis by coordinating to the leaving group and neutralize the
developing negative charge on the scissile phosphate (93).
Furthermore, the possibility of His-69 involvement with direct attack of the
scissile phosphate to form an enzyme-DNA covalent intermediate, subsequently
hydrolyzed by water, was excluded by Elliot and coworkers on the basis of
stereochemical analysis of the phosphate group (93;114,115). They deduced that the
formation and subsequent hydrolysis of an enzyme-DNA intermediate would proceed
34
- Chapter1 with two displacements at the phosphorus atom and therefore two inversions of
configuration.
Ultimately, the resulting product would have overall retention of
stereochemistry (115;116). Conversely, if His-69 deprotonates a water molecule which
then attacks the scissile phosphate directly, only one displacement occurs at the
phosphate and the resulting product would have overall inversion of stereochemistry.
Experiments conducted using H2 18 0 and Vsr substrates that contained the Rp
phosphorothioate isomer at the scissile phosphate site showed that the Vsr reaction
proceeds with inversion of configuration at the scissile phosphate, and, therefore, His-69
functions as a base to deprotonate the attacking water molecule.
The final catalytic requirement for nucleases is either a Lewis or a proton acid to
interact with, and thereby stabilize, the developing negative charge on the leaving group
(114;117). In the case of Vsr, the leaving group is the 3'-bridging oxygen atom at the
scissile phosphate. To probe if stabilization of the leaving group is achieved through
coordination with a metal center, this group was modified to contain a 3'phosphorothioate (which coordinates Mn2+better than Mg2+ ) and binding and hydrolysis
rates were measured (93).
While binding of Vsr to this modified substrate was
unchanged, hydrolysis in the presence of Mg2 + was strongly reduced. However, cleavage
was fully restored to wild type levels when Mg2+ was replaced with Mn2+ , strongly
suggesting that one Mg2+ serves as a Lewis acid and stabilizes the negative charge on the
leaving group as it develops.
In summary, when Vsr binds to a T:G mismatch in duplex DNA, the scissile
phosphate is coordinated and stabilized by two Mg2+ water clusters. His-69, acts as a
general base and abstracts a proton from a water in the metal water cluster. The resulting
activated water conducts a direct inline attack on the phosphate group between the
mismatched thymine and its 5' cytosine neighbor. The increased negative charge in the
transition state is localized on the two non-bridging oxygen atoms and is stabilized by the
two Mg2 + water clusters. One of the Mg2+ ions also acts as a Lewis acid to labilize the
leaving group by neutralizing the negative charge that develops as the reaction proceeds.
35
- Chapter
-
1.3. METHYL-DIRECTED MISMATCH REPAIR LESSONS FROM STREPTOCOCCUS PNEUMONIAE
Evidence of the existence of mismatch repair in prokaryotes and the idea of strand
discrimination
originated from early work involving transformation
studies in
Streptococcus pneumoniae (S. pneumoniae) (118). In a S. pneumoniae wt strain, the
efficiencies to which different transformants integrated was varied. Mismatch repair was
proposed to be responsible for this phenomenon because heteroduplexes formed during
integration would have contained
mismatches,
since the regions involved in
recombination were not completely homologous.
It was also observed that the
transformation efficiency was dependent not only upon the system's preference for
undertaking repair on the donor strand, underscoring strand discrimination, but also on its
preference for the type of mismatch on which repair was carried out. In other words,
different mismatches were repaired with different efficiencies.
Treatment of S. pneumoniae wt cells with mutagens led to the isolation of mutants
that did not discriminate between high and low efficiency markers (119). The mutations
responsible for this phenotype were designated hex (originally for "high efficiency
unknown [x]", now for heteroduplex repair deficiency) (118). Interestingly, hex mutants
showed an elevated level of mutation accumulation (120), suggesting that the hexdependent MMR system could play a role in mutation avoidance.
It was hypothesized that strand discrimination was established by the ability of the
system to recognize that donor strands possessed strand breaks at their ends (121). In
support of this hypothesis, low efficiency markers were sensitive to donor DNA that had
been UV irradiated (and thus contained a high number of strand breaks) in a fashion that
required the hex genes (118;122). Therefore, the mutator phenotype of the hex mutants
was likely due to their inability to recognize strand breaks and discriminate between
strands (123-125). Since the lagging strand of DNA is continuously forming breaks at
the ends of its Okazaki fragments during replication (124;126) it is believed that hex
36
- Chapter
-
dependent MMR is programmed to carry out repair on errors that arise during replication
to the newly synthesized strand.
Methyl-directed mismatch repair in E. coli was believed to be similar to that for S.
pneumoniae. Corroboration for that hypothesis was two-fold. It was observed that HexA
from S. pneumoniae is homologous to the MutS mismatch recognition protein from
Salmonella typhimurium, and could complement an E. coli strain deficient in MutS (127).
Additionally, the long standing hypotheses that repair was directed to the newly
synthesized strand via the methylation state of d(GATC) sites (128;129) was confirmed
(130).
1.4. METHYL-DIRECTED MISMATCH REPAIR IN E. COLI
Methyl-directed mismatch repair (MMR) in E. coli (131) is initiated when the
mismatch recognition protein MutS finds a substrate in DNA.
Upon mismatch
recognition, MutS interacts with its canonical partner MutL to activate the latent
endonuclease activity of MutH (132). MutH is a type II restriction endonuclease that
cleaves the newly synthesized strand, containing the incorrect base, adjacent to the A at
the nearest unmethylated d(GATC) site (132).
The ability of MutH to incise the correct
strand in the palindromic d(GATC) sequence is achieved by the methylation status of the
two strands (Figure 1.10). For a short time after replication, the d(GATC) sequence on
the nascent DNA strand is not methylated
(133;134).
MutH can only incise
unmethylated d(GATC) sequences (132). Thus, repair is directed by the methylation
status of the DNA, and, therefore, this characteristic has been reflected in the naming of
the repair system.
The incision site, which can be either 5' or 3' to the error, is the entry point for
removing the strand which contains the mismatch.
MutL has been shown to play a
pivotal role in loading DNA helicase II and recruiting single-stranded DNA binding
protein (135;136).
Additionally, contacts between MutL (as part of a MutL-MutS
37
- ChapterI complex) and DNA helicase II have been shown to activate the helicase (135;136).
Therefore, MutL plays an important role in communicating the direction of the error.
Models for how this communication is accomplished are discussed below.
Working in concert, the helicase and single-stranded binding protein expose
single stranded DNA that is subsequently digested by either 3' or 5' exonucleases,
depending on the orientation of the error (17;137). Originally, Exo I was implicated in
the degradation of DNA in the 3' direction (138). It was later shown that Exo X (139)
and Exo VII (140) could provide this function as well (141;142). On the other hand,
degradation of DNA in the 5' direction has been shown to involve RecJ or ExoVII (143).
Subsequent to excision of the error containing strand, Pol III and DNA Ligase to
resynthesize and ligate the DNA to restore the native structure.
Remarkably, the
resynthesis of a significant tract of DNA can be mandated by an incision site located up
to a couple of thousand bases away from the replication error (128;144). Though all
repair events are not necessarily long (145), MMR is sometimes referred to as long-patch
repair.
1.4.1. DAM - IN VIVO AND IN VITRO PROPERTIES
Methylation of the exocyclic AP position of adenine in d(GATC) sequences is
entrusted to the DNA adenine methylase, Dam (146), which is a 278 amino acid protein
that has an approximate molecular mass of 32 kDa (147). The importance of methylation
was confirmed by the observation that dam- strains of E. coli have increased rates of
spontaneous mutation (148;149). The responsibility of dam+ dependent methylation in
providing the strand discrimination signal was later established in experiments with
heteroduplexes made from two strands of different methylation status (129;146;150-152).
In hemimethylated sequences, repair was highly based towards the unmethylated strand,
with the methylated strand serving as the template during repair.
Experiments on
heteroduplexes showed that MMR will not act unless an unmethylated d(GATC) site is
present (153). Similarly, reduced expression of Dam results in reduced repair and hyper-
38
- Chapter1 mutation (33;149).
Over-expression of Dam also results in an increase in the
spontaneous mutation rate (154;155) because the window in which MMR has to act is
shortened (156) and, as a consequence, more mutations evade repair.
1.4.2. MUTS - IN VIVO AND IN VITRO PROPERTIES
The mutS gene codes for an 853 amino acid protein (157) with a monomer
molecular mass of approximately 97 kDa (158). The protein belongs to the Walker A/B
sequence motif of the NTPase families (159). Sedimentation experiments revealed that a
MutS homodimer acts as the recognition component of MMR by distinguishing
mismatches (160) and IDLs (23). Binding of MutS to mismatches contained in stretches
of up to 20 nucleotides was first demonstrated by DnaseI footprinting analysis (160).
The MutS footprint was also found to expand in the presence of MutL and ATP (161).
These conditions also activated a MutH dependent endonuclease activity at a
hemimethylated d(GATC) site (162).
MutS itself demonstrates a weak ATPase activity (163), and proper functioning of
this enzyme has been attributed to the essential need for ATP hydrolysis (132). Indeed,
introduction of mutations in the phosphate binding loop (P-loop) motif of the MutS ATPbinding site results in a dominant negative mutator phenotype (164). However, the role
of hydrolysis and the mode of signal transduction between mismatched nucleotides and
the excision machinery is under debate (165).
MutS has varying affinities for the eight possible mispairs, ranging from 10 to
more than 1,500-fold higher for mismatched DNA as compared with perfectly matched
DNA (166;167). The relative affinities of MutS for mismatches ranked in decreasing
order are T:G > C:A > G:G, A:A > T:T, C:T, G:A > C:C (150;158;160). The C:C
mismatch is recognized by MutS (158) but has not been reported as subject to MMR.
Although mispairs leading to transition mutations (purine to purine or pyrimidine to
pyrimidine) are, for the most part, better repaired than those leading to transversion
39
- Chapter1 mutations (purine to pyrimidine or visa versa), increased efficiency of repair may be
dependent upon sequence context. It has been reported that an increase in GC content in
the area surrounding the mismatch may increase repair efficiency (168).
Structures of MutS from Thermus aquaticus (169) and E. coli (1 70;170-1 75) have
been solved. The overall structure of a carboxyl-truncated E. coli MutS bound to a T:G
mismatch shows an asymmetric MutS dimer, where only one monomer is bound to ADP
and is recognizing the T:G mismatch. The ATPase domain of the other monomer is
unoccupied and reinforces the notion that the two subunits do not share equal roles in
mismatch recognition. The monomer units make contacts at the top and bottom to form a
dimer that has two large holes in the middle. The DNA is bound at one of these holes
and the other is unoccupied. It has been proposed that this hole could serve as a second
DNA binding site to allow for MutS to simultaneously bind DNA at the mismatch site
and at the MutH cleavage site by looping the DNA through this second hole (169).
Interestingly, the ATPase sites are located at the end of the complex opposite that
which binds DNA, invoking questions as to how ATP hydrolysis is coupled to mismatch
recognition.
While it has been observed that the binding of DNA by MutS stimulates
ATP hydrolysis (176), which results in a decrease in DNA binding affinity (177), the
physical basis for how DNA binding and ATPase activity is coupled is not understood.
The duplex DNA is in the B-form and has a strong kink of approximately 60 at
the mismatch site. The bases at the mismatch site are unstacked and have atypical roll,
tilt, and twist angles. The kink in the DNA is manifested in sugar pucker rearrangements
from C2'-endo (typical for B-form) to C3'-endo (typical for A-form) for a stretch of six
nucleotides.
The N-terminal mismatch binding domain of one subunit inserts into the minor
groove and makes direct contacts with the mismatch. At the center of the DNA binding
interface, a conserved Phe-36 residue is stacked against the unpaired thymine, and the
protein forms hydrogen bonds with a number of the unpaired bases. Additionally, Glu-38
40
- ChapterI is engaged in hydrogen bonding with the mismatched thymine residue. In normal base
pairs, these groups contacted by the protein would not be exposed. Therefore, it appears
as though MutS probes for the mismatch by looking for contacts that would not exist in
perfectly matched DNA.
Although MutS readily forms tetramers (160;176;177) the carboxyl-truncated
fragments of MutS do not crystallize as such (169;171) and the orientation of the two
dimers is therefore unknown. Perhaps such a structure would reveal more about how
signaling between MutS and other mismatch proteins is achieved. Regardless, the fact
that bacterial MutS dimers form structural heterodimers is interesting because of the use
of true heterodimers by eukaryotes (178). This asymmetry could be involved in strand
specific signaling of mismatches or evidence that the MutS monomer units operate
through conformational switching between the two units (1 70).
There is recent evidence that MutS, Pol I, and DNA Ligase all interact with the f3clamp accessory protein (179). This protein is typically associated with Pol III and is
responsible for processive DNA replication. The observation that the ,-clamp associates
with DNA repair proteins suggests that it is involved in a variety of reactions in addition
to chromosomal replication, and that certain instances of repair could be highly
orchestrated involving multi-component repairosomes.
1.4.3. MuTL - IN VIVOAND IN VITRO PROPERTIES
The mutL gene has been cloned from E. coli (180) and encodes a 70 kDa protein
monomer that can be over-expressed in E. coli and purified (161). MutL exists as a
homodimer through interactions in its C-terminus (161;181).
The MutL dimer is
recruited to the ATP bound MutS complex (161), in a manner that does not require
binding or hydrolysis of ATP by MutL (182).
41
- Chapter1 MutL, like MutS, has an ATPase domain that is critical to MMR (183). Binding
of ATP by MutL occurs at the N-terminus, causing conformational changes that induce
additional dimer contacts at its N-terminus (184). N-terminal dimerization, however,
does not require ATP hydrolysis, as nonhydrolyzable ATP analogs also produce Nterminal contacts (184). Mutations that inactivate hydrolysis, but not binding of ATP by
MutL, are strongly dominant negative to MMR in vivo (185) and in vitro (186).
Hydrolysis of ATP is therefore a critical for a step of MMR after the interaction between
MutS and MutL. Possibly, hydrolysis of ATP by MutL serves to activate MutH (101)
and assists the actions of DNA helicase II (186). In any case, the changes that take place
upon hydrolysis of ATP by MutL help to coordinate and regulate interactions critical for
MMR (184). In this regard, MutL has been deemed a molecular matchmaker (187).
Structural data for MutL (181;183;184;188) reveal that the MutL ring-like dimer
contains many residues for DNA binding on the inner surface of the ring. These data
suggest that MutL, like MutS, encircles DNA and possibly assists strand specific excision
by looping out the DNA to bring the mismatch site and MutH incision site within close
proximity (181).
1.4.4. MuTH - IN VIVO AND IN VITRO PROPERTIES
The mutH gene codes for a 229 amino acid protein with a molecular mass of
approximately 25 kDa (162).
MutH is structurally similar to a variety of type II
restriction enzymes (101). The protein consists of two globular domains connected by a
flexible junction, and a comparison of multiple MutH structures reveals that this junction
allows for multiple protein conformations (189). This flexibility could explain, in part,
why association with MutL is required for activity. MutL might assist in stabilizing
MutH in an active conformation. The active site of MutH is located in a shallow cleft
between the two globular domains and is coated with a number of basic residues that
likely attract the negatively charged phosphate backbone of the DNA substrate.
42
- Chapter1 MutH has an extremely weak Mg2+ dependent endonuclease activity that nicks
hemimethylated DNA, 5' to the G in the sequence d(GATC) (162). Initial measurements
of the endonuclease activity for the purified enzyme demonstrated less than one cleavage
event per hour per MutH monomer, leading to the hypothesis that the activity of MutH
was somehow stimulated by other components of MMR (131;162). This notion was
strengthened by the observation that the activity of the same protein preparation was
approximately 20 to 70-fold higher when added to a reconstituted methyl-directed
mismatch repair system. It was later demonstrated that a MutS-MutL mismatch ternary
complex, and ATP hydrolysis, allowed for full activation of MutH (132). Interestingly,
high concentrations of MutL have been shown to be sufficient for activating MutH in an
ATP-dependent manner (101). The need for the MutH and/or the d(GATC) incision site
can be bypassed altogether if a nick is present in the undermethylated strand (131;153).
An important observation is that MutH can cleave both DNA strands if the
d(GATC) sites are completely unmethylated (132). Decades earlier, it was observed that
dam mutants, which contain fully unmethylated d(GATC) sites, were killed by growth in
the presence of base analogs that form substrates for MMR in DNA, such as 2aminopurine (190). This observation lead to the suggestion that, in the absence of
methylation, MMR would initiate repair on either strand, leading to the formation of
double strand breaks (190).
This hypothesis received criticism on the grounds that
mismatches did not occur frequently enough to produce excision tracts that would
overlap (191). Therefore, the finding that MutH can create a double strand break from
one mismatch, assists in explaining cell death in dam- strains.
Nicking of the undermethylated strand by MutH can take place either 3' or 5' to
the mismatch (Figure 1.11) (132). The choice of which single stranded exonuclease will
carry out the excision and degradation of the DNA is contingent upon DNA helicase II
unwinding the DNA at the nick site in the appropriate direction towards the mismatch
(143). Though the appropriate direction for DNA unwinding is coordinated by MutL
(186), the precise method of signaling is not understood. The next section describes two
prominent models for how MMR is orchestrated.
43
- ChapterI 1.4.5. MECHANISM OF METHYL-DIRECTED MISMATCH REPAIR
One model (18;145) which is commonly referred to as the Hydrolysis Dependent
Translocation Model (Figure 1.12) postulates that the assembly of a MutS/MutL complex
at the mismatch site uses ATP hydrolysis to motor bidirectionally and to mediate the
formation of an a-shaped loop structure containing the substrate mismatch (177). MutL
enhanced the rate of MutS-mediated DNA loop formation and both MutS and MutL
appeared to be bound at the base of the a-loop. Loop formation required a mismatch and
ATP. Hydrolysis of ATP (and not just binding) was necessary for translocation, as
nonhydrolyzable ATP analogs were not sufficient to promote loop growth. The original
model was modified (192) to accommodate the observation that MutS appears to move a
large distance on the DNA per ATP hydrolysis event. Such a DNA tracking process was
envisioned to link MutS mismatch recognition with MutH endonuclease activity at a
nearby d(GATC) site as well as to provide directionality for subsequent loading of the
UvrD helicase and the appropriate ssDNA exonuclease.
An explanation as to how MutS can translocate bidirectionally (193-195), is based
on the idea that the nucleotide binding sites of MutS can act as "latch (L) sites" that can
be in the open or closed conformation depending on their occupancy. If ADP is bound to
one L site, it will be in the closed conformation and not allow translocation at that site.
Simultaneously, however, the other L site will be ATP bound and open, therefore
allowing translocation in that direction.
Hydrolysis of ATP in the second site,
accompanied by replacement of ADP with ATP in the first site, allows translocation to
occur in the other direction. This process repeats, so long as ADP and ATP are around,
to achieve bidirectional translocation.
Another model (165) commonly referred to as the Molecular Switch Model
(Figure 1.13, 1.14, and 1.15) is based on the observations (182) that ADP-bound MutS
binds to mismatched DNA and then exchanges ADP for ATP. Nucleotide exchange is
said to result in conformational changes that destabilize mismatch binding and give rise
to an ATP-bound MutS sliding clamp that diffuses along the helix, independent of ATP-
44
-Chapter
Ihydrolysis (182;196;197). Interestingly, ATP binding by the human MutS homolog,
hMSH, has been shown to result in the formation of a DNA sliding clamp capable of
hydrolysis-independent diffusion for several thousand nucleotides (198).
It is also
noteworthy to mention that only the nucleotide-free and ADP bound structures of MutS
have been solved because infusion of ATP or ATPyS disintegrates the MutS crystal
(172).
It appears that ATP binding by MutS invokes conformational transition(s).
Iterative loading of multiple clamps was responsible for marking the mismatch and
creating a gradient of clamps, providing directionality, along the duplex surrounding the
mismatch site (198;199). MutL is said to interact with ATP-bound MutS clamps to
produce complexes that can interact with, and activate, MutH endonuclease.
Observations indicate that interaction between the MutL complex and MutH increases the
ability of MutL to bind ATP, and in turn the ATP-MutL complex activates the
endonuclease activity of MutH (182).
MutL, in this sense, is a protein activated
molecular switch. Subsequent to incision, UvrD helicase begins unwinding the DNA
with an activity that is enhanced by MutS and MutL. This property was hypothesized to
direct the helicase towards MutS-MutL complexes and therefore towards the mismatch.
1.5. MISMATCH REPAIR IN EUKARYOTES
Eukaryotic mismatch repair (6;8;16) is functionally homologous to that of E. coli,
plays an important role in mutation avoidance, and is critical to health. Defects in human
mismatch repair have been linked with a predisposition to hereditary nonpolyposis colon
cancer and ovarian cancer. Eukaryotic MMR proteins also have been associated with the
triggering of programmed cell death.
Many eukaryotic DNA repair enzyme homologs have been identified and their
functions characterized (Table 1.1) (18;200;201). However, to date, no MutH homologs
have been assigned.
Additionally, the mechanism by which strand discrimination is
achieved is not thought to include methylation, as in E. coil.
Rather, strand
discrimination is possibly directed by nicks which exist at the ends of Okazaki fragments
45
- Chapter
-
in lagging strand synthesis and at the growing 3' end of the leading strand (202;203).
Another possibility is that the proliferating cell nuclear antigen (PCNA) could assist in
discriminating the parental strand from newly synthesized strand through its interactions
with various MMR proteins (204;205).
Interestingly, the homologs to E. coli MutS do not form asymmetrical
homodimers as described above.
Rather, they form a variety of protein-protein
heteroduplexes depending on the substrate.
It is hypothesized that subsequent to
recognition of the MMR substrate, the MutS complex will form a clamp-like structure
and slide along the DNA. Formation of a repairosome proceeds by association of MutLlike heterodimers and PCNA to the MutS-like complex. This complex then identifies the
nascent strand and initiates repair by some mechanism yet to be described. Repair is then
carried out by proteins including DNA polymerase 6, replication protein A (RPA),
replication factor C (RFC), and exonuclease EXO I/HEX 1.
1.6. INTERACTION BETWEEN THE VSPR AND MMR SYSTEMS
A comparison between the long patch (MMR) and short patch (VSPR) mismatch
repair systems discussed above reveals that they vary in a number of ways.
As
highlighted by the names of the repair systems, the average length of the repair tract
associated with each is quite different. While a VSPR repair tract rarely exceeds 10
nucleotides in length, that for MMR has been observed to reach thousands of nucleotides.
MMR repairs a large number of substrates including a multitude of base-base
mismatches, IDLs, and chemically modified bases. VSPR, on the other hand, repairs T:G
and U:G mispairs to yield C:G in all cases. MMR has the requirement that the DNA
must contain an unmethylated d(GATC) sites for MutH incision.
In contrast, the
methylation status of d(GATC) sites has no effect on VSPR. However, VSPR does have
sequence context specificity. Therefore, many differences exist between these two repair
pathways.
46
- Chapter1 One thing in common between these two repair pathways is that they share the
ability to act on T:G mismatches.
Furthermore, VSPR has the opportunity to act
incorrectly on T:G mismatches that arise from the misincorporation of G opposite T.
These types of mismatches should be processed correctly by MMR before VSPR gets a
chance to act, thus avoiding a fixation of the mutation. Therefore, given the intention of
cellular repair systems to avoid DNA mutations, the VSPR and MMR systems should be
coordinated with each other. This section discusses how the two systems may interact.
As highlighted above, both the VSPR and MMR systems use MutS and MutL for
initial mismatch recognition. It was originally thought that VSPR could not function in
cells lacking the muts or mutL gene (41-43;46). However, it was subsequently shown
that the level of VSPR in mutS and mutL mutants was only reduced, not eliminated (70).
The reduction of VSPR was also observed upon over-expression of MutS from a
multicopy plasmid (80;206), suggesting that the level of MutS maintained by the cell is
optimal for its functions in VSPR. In contrast, introducing a MutS producing plasmid
into mutS cells does not affect MMR (164). These observations led Lieb (206) to study a
number of mutS
alleles, that were dominant negative for MMR, for their effects on
VSPR. Lieb found that many of the MutS mutants (Class II) did prevent VSPR. These
alleles had previously been found to reject complementation by muts+, mutL, or mut
plasmids (164), indicating that the host mutant allele products bind to mismatches and
prevent further access.
Some mutants (Class I), however, were found to not disrupt
VSPR in mutS* cells and actually had the effect of stimulating VSPR in mutS cells.
These results indicated that specific MutS alleles, lacking functions for MMR, can still
participate in VSPR.
A study of these Class I alleles revealed that some were complemented by overexpression of MutL and/or MutH from plasmids (164), indicating that the nature of the
host mutS mutations affected the way in which the alleles interacted with the MutL and/or
MutH proteins. While VSPR is compromised in cells lacking MutL (41-43;46),mutH
strains retain full repair activity (46). Therefore, these alleles were capable of interacting
with MutL for VSPR activity but not for MMR.
47
Not only did these observations
- Chapter
-
highlight that the roles of these recognition proteins in VSPR and MMR are different, but
also that MutL plays an integral role in each repair system.
Evidence that MutL actually balances the two repair systems came from
experiments done in the Cupples Lab.
Mutational studies in cells that were over-
expressing Vsr revealed that not only did high levels of Vsr result in T -- C mutations,
but also a large number of other unexpected mutations, including frameshifts (207,208).
It was unlikely that this mutator phenotype resulted from competition between Vsr and
MutS for substrates. Rather, a more likely explanation was that Vsr was competing for a
component of, and preventing, MMR.
In vitro studies revealed that binding of Vsr to its substrate is promoted by MutL
catalytically (209). In contrast, stimulation of MutS binding to the Vsr substrate by MutL
was increased by 6-fold stoicheometrically. In agreement with these findings, bacterial
two-hybrid studies have confirmed in vivo interaction between Vsr and MutL (210).
Additionally, interaction between Vsr and MutL inhibited the ability of MutL to
dimerize, interact with MutS and/or MutH, and to mediate a previously unknown
interaction between MutS and MutH. No interaction between Vsr and MutS and/or
MutH was observed. All these data together strongly suggested that Vsr makes critical
contacts with MutL and that these interactions can detract MutL from its role in MMR.
Further investigation (94) into the nature of the interaction between Vsr and MutL
revealed that a N-terminal truncation mutant of Vsr (Vsr-A14), cannot be stimulated by
MutL in vivo, and promotes diminished mutagenicity. In contrast, four derivatives of
full-length Vsr that were enzymatically inactive (F67A, W68A, H69A, and H71A), were
as mutagenic as the wt protein. Therefore, the ability of Vsr activity to be stimulated by
MutL, and the ability of the Vsr-MutL interaction to be detrimental to MMR, resides in
the N-terminal domain (NTD) of Vsr and not in the proteins incision activity.
Interaction between Vsr and MutL was shown to be independent of the first 19
residues of Vsr (210). Therefore, the observation that the Vsr-A14 mutant has diminished
mutagenicity is not simply because it does not bind MutL. How then does Vsr inhibit
48
- Chapter1 MutL from participating in MMR? To better understand how inhibition might occur, it
will be useful to revisit the role of ATP hydrolysis by MutL in MMR.
Studies of MutL revealed that the protein must bind and hydrolyze ATP to be
functional in MMR (183).
Further investigation (described above) revealed that
hydrolysis of ATP by MutL stimulates the endonucleolytic activity of MutH, and results
in loss of N-terminal contacts in the MutL dimer (184). Additionally, re-dimerization of
the N-termini is thought to be the rate limiting step in the ATPase cycle of MutL (184).
Therefore, after stimulating the protein responsible for nicking DNA, MutL needs to bind
another ATP and dimerize at its N-terminal before it can take part in another repair event.
It is plausible that MutL goes through a similar cycle with Vsr. That is, interaction
between Vsr and MutL, invokes MutL to hydrolyze ATP to convert Vsr into its active
form. In the process, MutL is rendered incapable of assisting another repair event for a
short time. Given the observation that the Vsr-A14 mutant has diminished mutagenicity,
as compared with the wt enzyme, it is plausible that the NTD of Vsr is involved in MutL
dependent stimulation of the endonuclease. Temporary incapacitation of MutL would
explain how over-expression of the full length Vsr is mutagenic. It also explains how the
Vsr-A14 is not mutagenic. If the NTD of Vsr is not present for stimulating the activity
of Vsr, MutL will not give up its activated state by hydrolyzing its ATP. To understand
how MutL interacts with the NTD of Vsr, to stimulate endonuclease activity, we must
look at how the NTD of Vsr is ordered.
Early on, trypsin and chymotrypsin proteolysis studies revealed that the NTD of
Vsr is unstructured (98), and it had to be removed before the free protein would
crystallize (98). It was also observed that the tail was labile in vitro (see below) (81).
However, crystallographic studies revealed that, at some point, the NTD of Vsr becomes
ordered and forms an a-helical structure that lies across the minor groove of DNA (91).
In purified protein, ordering of the NTD of Vsr appears to be very slow (211). It was
therefore hypothesized that the interaction between MutL and full length Vsr in vivo
helps the NTD of Vsr achieve an ordered conformation that stabilizes Vsr binding to
DNA (94). This notion is consistent with data showing that MutL can stimulate Vsr
directly (89).
49
- Chapter
-
It is interesting to note that a N-terminally truncated form of Vsr, missing 12 to 14
residues, was observed to form upon storage (81). This finding shed light on the lability
of the NTD, and also complicated initial attempts to characterize Vsr biochemically (89).
Early preparations of Vsr were reported to contain enzyme with only 1% activity (89).
Interestingly, reliable biochemical studies have reported a kcat of 2.8 min' and 0.09 min -'
for wt Vsr and Vsr-A14 respectively (94). These data indicate an activity for the Vsr-A 14
mutant of approximately 3%, as compared with the wt Vsr. Therefore, early kinetic
studies on Vsr (89;90) were likely conducted on the truncated form of Vsr and might not
accurately reflect behavior in vivo.
Given that the VSPR and MMR pathways are balanced by the availability of
MutL, a mechanism by which they avoid competition with each other is imperative. As
mentioned earlier, the level of Vsr in the cell is dependent on the phase of growth. Vsr
was found to be present in the cell at low levels during exponential phase and at elevated
levels as the cells entered stationary phase (69). This trend is consistent with the notion
that VSPR utilizes the MutL protein after it has carried out its duties in MMR.
MMR is active just after replication, during the time the newly synthesized strand
is unmethylated at its d(GATC) sites (133;134), strand discrimination can occur (152),
and MutH is capable of initiating a repair event by incising the DNA (153). Once Dam
has fully methylated the d(GATC) sites in DNA, MutH cannot initiate MMR (162), and
therefore relieves MutS and MutL from their commitment to that repair pathway.
Accordingly, like Vsr, the level of MutH in the cell is dependent on the phase of
cell growth, and has been observed to decrease by 3-fold when cells are in nonreplicating conditions (212;213). Interestingly, in stationary cells, the level of MutS is
decreased by approximately 10-fold whereas the level of MutL remains constant (212). It
therefore appears that the ability of the cell to balance MMR and VSPR relies on
regulating the levels of Vsr and MutH so that they do not compete for MutL. This system
explains why, during stationary phase, MMR is relatively inefficient (214) and VSPR is
most efficient (68). Given that replication errors produce substrates for MMR during the
growth phase, and accumulation of deaminated 5mC residues occurs in stationary phase,
50
- Chapter
-
it is reasonable that E. coli has evolved to balance two distinct repair pathways and
maximize the efficient use of the proteins they share in common.
A number of mechanisms for how the level of Vsr is controlled in the cell have
been proposed. One posits that the expression of Vsr is regulated by overlap with Dcm.
The 5' end of vsr overlaps the 3' end of dcm by 6 codons in a +1 reading frame (Figure
1.1) (48). The two genes are transcribed from a common promoter, and therefore the
ribosome must frameshift to produce two proteins from the same transcription product.
The shift in translational reading frames has been studied and found to be approximately
4-fold more efficient as cells enter stationary phase (215). The increase of Vsr in cells as
enter stationary phase (69) might therefore be regulated by the cells efficiency at
translational frameshifting.
Another theory for how the level of Vsr might be controlled in the cell came from
in vivo VSPR assays in dam- strains (70). Dcm-induced CCAGG to CTAGG mutations
were found to be increased in dam- strains more than in mutS strains. The mutation level
was dependent on the concentration of Dcm methylase, and therefore resulted from a
deficiency in VSPR, not MMR.
Interestingly, western analysis revealed that the
regulation of Vsr in dam- strains was found to take place post-transcriptionally. While
the level of vsr transcript was the same in wt and dam- strains, the absolute amount of Vsr
in the damnistrains was reduced. Therefore, though the Dam methylase does not affect
the growth phase-dependent level of Vsr, it has a significant effect in maintaining the
absolute level of Vsr in the cell.
1.7. MECHANISM
OF
VSPR
The mechanism by which MutS and MutL stimulate Vsr is not fully understood.
The observations (as discussed above) that a slow conformational change of the Vsr NTD
occurs upon DNA binding (211), that ordering of the NTD is essential for forming a
stable protein DNA complex (91), and that MutL can stimulate Vsr directly (89), suggest
that MutL activates Vsr by ordering its NTD. It has also been hypothesized that MutS
51
- Chapter
-
and MutL assist MMR by presenting mismatches within a a-shaped loop (177). As a
result, the location of the mismatch is more defined. Also, depending on the size of the
loop, it could have the potential to increase negative supercoiling of the DNA and cause
unwinding, and thus more exposure, to the mismatch site.
Altogether a likely explanation for how MutS and MutL stimulate Vsr is as
follows. Initial recognition of the mismatch is carried out by MutS, which then recruits
MutL to assist in presenting the mismatch in a looped DNA structure.
Vsr is then
recruited to the complex because fully methylated d(GATC) sites render MutH incapable
of DNA incision. Contacts are made between MutL and Vsr which activate Vsr by
ordering its NTD so that it forms a stable complex with its DNA substrate. Vsr then
identifies its substrate DNA mismatch located within the loop and incises the T
containing strand to initiate VSPR.
1.8. REPAIR OF T:G MISMATCHES IN EUKARYOTES
At least two proteins exist for the specific repair of T:G mismatches in
eukaryotes. In 1993, Nedderman and Jiricny isolated a human enzyme that excises T
from T:G mismatches (216). The repair protein was found to be a glycosylase (TDG)
that also initiates BER on uracil sites in DNA (217). However, unlike previous uracil
glycosylases (Family 1) (218), TDG only excises uracil when it was opposite a guanine.
Uracils opposite adenine or in single-stranded DNA are not excised by this enzyme. A
structural homolog to TDG does exist in bacteria; however this enzyme, Mug, only acts
efficiently on uracil and does not share sequence homology with TDG (219). Multiple
reports are consistent in the conclusion that Mug does not excise T opposite G (220-222).
The second protein present in eukaryotes for the repair of T:G mismatches is the 5methylcytosine-binding domain glycosylase 4 (MBD4) (223). This protein also initiates
BER by recognizing and hydrolytically removing T opposite G in the context of a
methylated CpG site.
52
-Chapter
IFigure 1.1. Organization of the dcm and vsr genes. The 5' end of vsr overlaps the 3' end
of dcm by 7 codons in a +1 reading frame. This genetic arrangement was hypothesized to
assure that Vsr is always produced along with Dcm to minimize the mutagenic effects of
cytosine methylation. Although the two genes are transcribed from a common promoter,
two separate proteins are produced.
E A Q H G R R S R u
dcm
dcmACAACATGGCCGACGTTCACGATAAGGCCACTRRSR
5' ..
AAGAGGCACAACATGGCCGACGTTCACGATAAGGCCACT
vsr
MA D
53
V
H
D
K
A
T
- Chapter1 Figure 1.2. Repair of mismatches that arise through methylation and subsequent
deamination of cytosine. Dcm methylates the C5 of the pyrimidine ring of the underlined
C in the sequence 5'- CC(A/T)GG -3'. Subsequent deamination of that residue gives rise
to a T:G mismatch, which is a substrate for very short patch repair. Vsr initiates repair by
producing a single strand nick 5' to the mismatched T. This activity is stimulated by, but
does not require, MutS and MutL. Pol I carries out nick translation by removing and
replacing the mismatched T and a small number of nucleotides downstream of the
mismatch. DNA Ligase seals the nick to complete the repair event.
CH3
5'-CC(A/T)GG-3'
3'-GG(T/A)CC-5'
I
Dcm
5'-CC(A/T)GG-3'
3'-GG(T/A)CC-5'
I
Spontaneous
Deamination
1. Pol I
2. Ligase
,G T(A/T)GG-3
.GG(TIA)CC-5'
e
Is also
Methylated
of 5mC
Vsr
(MutS/ MutL)
54
5'-CT(A/T)GG-3'
3'-GG(T/A)CC-5'
- Chapter
-
Figure 1.3. Unmethylated and hemimethylated Vsr substrates
CH 3
5'-
CCAGG
3'- G(GTCC
-3'
Dcm
-5,
CCAGG -3'
GGTCC -5,
5'
3,-
I
CH3
Spontaneo us
Deaminati( on
of 5mC
r
5'- CTAGG
-3'
5 -
3'- GGTCC
-5,
3
-
CTAGG -3,
GGTCC -5,
I
CH3
Unmethylated
Vsr Substrate
Hemimethylated
Vsr Substrate
55
- Chapter
-
Figure 1.4. Proposed mechanisms for the hydrolytic deamination of cytidine. One
mechanism involves direct attack at the 4-position of the pyrimidine ring of cytidine (1)
by a hydroxyl ion followed by loss of ammonia to yield uridine (4). The other
mechanism involves an addition-elimination reaction where water is added to the 5,6
double bond of cytidine (1) to form a dihydrocytidine intermediate (2). This intermediate
is then further attacked by water to release ammonia and yield uridine hydrate (5), which
then dehydrates to uridine (4). In DNA, R symbolizes the deoxyribose-phosphate
backbone.
NH.
H2 0
1.
2.
H20j
H2 0
NH 4 +
8
H3 N
HN
0
3.
O
OH
H
HNH
RN H
5.
R
R
H2
NH4
H
H
4.
56
- Chapter1 Figure 1.5. Chemistry behind C:G to T:A transition mutations. Modification of cytosine
(A) can occur via deamination and methylation. Deamination of cytosine gives rise to
uracil (B), and methylation of cytosine produces 5-methylcytosine (C). Subsequent
deamination of 5-methylcytosine produces thymine (D). Replication past either uracil or
thymine will give rise to a C:G to T:A mutation.
O
NH2
Deamination
%N
NH
N
) ®./
HO
0:- O
Repair
o
I
C.
>>I,
CG to TA Mutation
I
t
Deamination
O
NH
I
H
I
HY
N
57
-
1U
- Chapter1 Figure 1.6. Model for how VSPR could be shaping the E. coli genome. If a replication
error occurred such that a guanine was misincorporated opposite thymine, the resulting
mismatch could be a substrate for both MMR and VSPR. While MMR would repair the
error correctly in favor of thymine, VSPR would repair the mismatch in favor of guanine.
Over time, this process could lead to the underrepresentation of 5'- CTAGG -3' and overrepresentation of 5'- CCAGG -3' sequences.
5' -CTAGG-3'
3' -GATCC-5'
IReplication
Error
5' -CTAGG-3
I
3'-GGTCC-5I
VSPR
MMR
5'-CTAGG-3'
3'-GATCC-5'
5' -CCAGG-3'
3'-GGTCC-5'
58
- Chapter 1 Figure 1.1. Structure ofVsr
of Vsr forms two a-helicies
DNA from the major groove
ions are bound by residues
water mediated contacts.
bound to product DNA, minor groove side. The N-terminus
that pin down the DNA. The main core of Vsr binds the
side, where the chemistry takes place. The two magnesiUlll
from the ~-sheet and assist DNA binding through multiple
59
- Chapter 1 Figure 1.8. Structure ofVsr bound to product DNA, major groove side. Vsr has a three
layered a/~/a fold that is stabilized by a structural zinc. Vsr also has two magnesium
ions that participate in water clusters to bind DNA. The DNA is significantly unwound
and bends off axis by 44°. Three residues insert themselves into the opening and help
orient His-69 (shown in ball and stick) to activate a water for attack of the scissile
phosphate.
60
Chapter1
o ntr a
cs
triC
Figure 1.9. Geonme chirasraectiteri s tics ooff C G
a ndc T:G base pairs In contrast
a
p
d
ll
e
d
s
a
:Gm inbeato
TY
the ar,
is:G
aihrse, th
oPes
balIssemaptc
th
m
d. P r r e 2g
oemetry PpushieeTshCte
the
ajor groove andydistorte
D)NA duplex Ae
to the m
.
ferenc ,-,cnc
e
. Adapted from re
ex
le
p
u
d
A
N
D
10o
I .
io10.3jit
61
rte
king With
base pair stac
This4j
- Chapter1 Figure 1.10. Methyl direction of mismatch repair. Replication can cause mismatches to
form. The mismatched base in the newly synthesized unmethylated strand is targeted for
repair. MMIR excises the mismatched base and carries out resynthesis. The newly
synthesized strand is then methylated by Dam.
CH 3
3'
3'
3
G GATC
3'
C C--CTAG
5'
I
CH 3
5'
3'
CH 3
CH33
C
3'
5'
3'
-C
GGATC
CTAG
5
3'
5
CH
I 3
5'
G
GATC
3'
3'
C
CTAG
5'
CH 3
CH3
62
-Chapter
IFigure 1.11. Methyl-directed mismatch repair is bidirectional. The point of incision by
MutH can be upstream or downstream of the mismatch. Depending on the orientation of
the mismatch with respect to the nick, either 3' exonuclease (Exo I) or 5' exonuclease
(Exo VII or RecJ) activity is appropriate for removing the intervening nucleotides.
Loading of the helicase in the correct orientation, for subsequent exonuclease action, is
accomplished by the MutS-MutL complex. Pol III fills in the repair patch and DNA
Ligase seals the nick to complete repair.
CH 3
CH 3
3'
5'
-
5'
3'
NV,
MutH, MutL
MutS, ATP
CH 3
CH 3
I
A
3 -
5, -
CH
5'
3'
3'
5'
I
CH
3
I
3
A1
64.
i-L1
5'
,3,
1\/111111
Exo
or Exo
3'-
I
MutL, MutS,
x
helicase II, ATP
CH 3
I
Exo VII
or RecJ
CH 3
kA5'
3''
3'
5'
0
5-
CH 3
CH 3
·
5'
5,
,
.
CH 3
.
.~~~~~~~~~
_i
CH 3
CH 3
5'
3'
3'
5'
63
I
3'
I
SSB, ligase
I
I'
-
DNA pol III holoenzyme,
CH 3
II
-9f
I
5'
3'
- Chapter 1 Figure 1.12. ATP-hydrolysis dependent translocation model for MMR. MutS binds to
the mismatch and recruits MutL to the site.
This complex uses ATP to motor
bidirectionally and mediate the formation of a loop structure.
MutH beCOlnes activated
by the MutS-MutL complex and makes an incision on the error-containing strand at the
unmethylated d(GATC) site (not shown). DNA helicase II then loads onto the DNA at
the incision site and unwinds around the DNA loop.
Hemimethylated d(GATC) sites
.
....,
DNA mismat~
/
CH3
~
CH3
-:.
MutS
64
MutL
DNA
- Chapter 1Figure 1.13. Molecular switch model for MMR, part 1. MutS bound to ADP acts as a
mismatch sensor. Mismatch binding promotes MutS to exchange ADP with ATP and
then diffuse along DNA, independent of hydrolysis. Diffusion of ATP-MutS re-exposes
the mismatch, allowing multiple clamps to load and slide. MutL interacts with ATPMutS clamps and slides with them as part of a ATP-MutS-MutL complex, independent of
hydrolysis.
Binding of MutL to MutS promotes unloading of the clamp and will
therefore abort sliding in the absence of key MMR cOlnponents.
MutS
MutS
~
.
Mutl
0
Iftl
Multiple MutS sliding clamps
~7Jn
[-=-] n
............. ~
Multiple MutS-MutL sliding clamps
+
Pi
65
- Chapter 1 Figure 1.14. Molecular switch model for MMR, part 2. MutL physically connects the
ATP-MutS-MutL complex to MutH at the unmethylated d(GA TC) site. MutH increases
the ability of MutL to bind ATP, which in turn activates the endonucleolytic activity of
MutH. In this regard, MutL is a protein-activated molecular switch.
MutH
r
.............. ~
hemimethylated
d(GATC) site
!
r
CH3
MutL+ATP
Nick on unmethylated strand!
at GATe site
............. ~
-G-fl----I--CH3
66
- Chapter 1 Figure 1.15. Molecular switch model for MMR, part 3. DNA helicase II is loaded onto
the DNA and initiates unwinding in a manner which requires ATP and is stimulated by
MutS plus MutL. Theories posit that the helicase unwinds in the appropriate direction
because MutL either assists in its loading or it orients the he Iicase correctly by interacting
with it from the direction of the mismatch. Upon unwinding, the DNA is degraded by an
exonuclease and resynthesized by Pol III.
DNA helicase II
6.
~ MutL + AlP
.............~
~ADP
"-f6--f¥:'
helicase loading
CH3
r-
~"'''ii-~
AlP
ADP
t+
helicase unwindin9..
I
CH3
r- AlP
t+AD P
exonuclease
de radation of DNA
67
- Chapter
-
Table 1.1. E. coli and S. cerevisiae proteins required for MMR. Adapted from Schofield
and Hsieh (200).
S. cerevisiae
E. coli
MutS - Binds to mismatches and small
IDLs
Msh2/Msh6 - Binds to mismatches and 1base IDLs
Msh2/Msh3 - Binds to IDLs; has a role in
double-strand break repair
Msh4/Msh5 - No role in repair; meiosisspecific MutS homolog
MutL - Molecular matchmaker; interacts
with MutS in a mismatch-specific
manner to activate MutH cleavage
activity
Mlhl/Pmsl - Primary MutL homolog for
postreplication repair
MutH - Nicks unmethylated strand at a
hemimethylated d(GATC) site, initiating
repair
No known homolog
[3-clamp - Processivity clamp for Pol III,
interacts with MutS in vitro, may recruit
MutS to the replication fork
PCNA - Processivity clamp for Pol 8 and
Pol ; interacts with Msh3 and Msh6,
increases mispair binding specificity and
also involved in repair resynthesis
Helicase II - (UvrD) delivered to nick by
MutS and MutL; unwinds DNA prior to
its excision
No known homolog
Mlhl/Mlh3 - Involved in repair of some
IDLs; also functions in meiosis
Mlhl/Mlh2 - Minor role in suppression of
frameshifts
RecJ, Exo VII - Required for 3' to 5'
Excision between nick and mismatch
Exo I, Exo X - Required for 5' to 3'
Exo I - 5' to 3' exonuclease, deletion of
which results in mild mutator phenotype
Excision between nick and mismatch
Pol III - DNA polymerase required
Pol5 - DNA polymerase required for repair
resynthesis
For repair resynthesis
SSB - Single-strand-DNA-binding
aids excision and resynthesis
protein;
RPA - Single-strand-DNA-binding
aids resynthesis
DNA Ligase - Seals nicks
DNA Ligase - Seals nicks
68
protein;
- Chapter
-
1.9. REFERENCES
(1) Fishel, R. (1998) Mismatch repair, molecular switches, and signal transduction.
Genes Dev. 12(14), 2096-2101.
(2) Grilley, M., Holmes, J., Yashar, B., and Modrich, P. (1990) Mechanisms of
DNA-mismatch correction. Mutat. Res. 236(2-3), 253-267.
(3) Harfe, B. D., and Jinks-Robertson, S. (2000) DNA mismatch repair and genetic
instability. Annu. Rev. Genet. 34, 359-399.
(4) Harfe, B. D., and Jinks-Robertson, S. (2000) Mismatch repair proteins and
mitotic genome stability. Mutat. Res. 451(1-2), 151-167.
(5) Hsieh, P. (2001) Molecular mechanisms of DNA mismatch repair. Mutat. Res.
486(2), 71-87.
(6) Jiricny, J. (1998) Eukaryotic mismatch repair: an update. Mutat. Res. 409(3),
107-121.
(7) Kolodner, R. (1996) Biochemistry and genetics of eukaryotic mismatch repair.
Genes Dev. 10(12), 1433-1442.
(8) Kolodner, R. D., and Marsischky, G. T. (1999) Eukaryotic DNA mismatch
repair. Curr. Opin. Genet. Dev. 9(1), 89-96.
(9) Li, G. M., and Modrich, P. (1995) Restoration of mismatch repair to nuclear
extracts of H6 colorectal tumor cells by a heterodimer of human MutL homologs.
Proc. Natl. Acad. Sci. U. S. A. 92(6), 1950-1954.
(10) Li, G. M. (2003) DNA mismatch repair and cancer. FrontBiosci. 8, d997-1017.
(11) Modrich, P. (1987) DNA mismatch correction. Annu. Rev. Biochem. 56, 435-466.
(12) Modrich, P. (1997) Strand-specific mismatch repair in mammalian cells. J. Biol.
Chem. 272(40), 24727-24730.
(13) Rasmussen, L. J., Samson, L., and Marinus, M. G. (1996) Dam-directed DNA
mismatch repair. In DNA Damage and Repair (Nickoloff, J. A., and Hoekstra, M.
F., Eds.) pp 205-228, Humana Press, Totowa, N.J.
(14) Yang, W. (2000) Structure and function of mismatch repair proteins. Mutat. Res.
460(3-4), 245-256.
(15) Yang, W., Junop, M. S., Ban, C., Obmolova, G., and Hsieh, P. (2000) DNA
mismatch repair: from structure to mechanism. Cold Spring Harb. Symp. Quant.
Biol. 65, 225-232.
69
- Chapter
-
(16) Jiricny, J., and Nystrom-Lahti, M. (2000) Mismatch repair defects in cancer.
Curr. Opin. Genet. Dev. 10(2), 157-161.
(17) Modrich, P. (1991) Mechanisms and biological effects of mismatch repair. Annu.
Rev. Genet. 25, 229-253.
(18) Modrich, P., and Lahue, R. (1996) Mismatch repair in replication fidelity, genetic
recombination, and cancer biology. Annu. Rev. Biochem. 65, 101-133.
(19) Welch, M. M., and McHenry, C. S. (1982) Cloning and identification of the
product of the dnaE gene of Escherichia coli. J. Bacteriol. 152(1), 351-356.
(20) Schuermann, R. M., and Echols, H. (1984) A separate editing exonuclease for
DNA replication: the E subunit of Escherichia coli DNA polymerase III
holoenzyme. Proc. Natl. Acad. Sci. U. S. A. 81, 7747-7751.
(21) Cox, E. C. (1976) Bacterial mutator genes and the control of spontaneous
mutation. Annu. Rev. Genet. 10, 135-156.
(22) Dohet, C., Wagner, R., and Radman, M. (1986) Methyl-directed repair of
frameshift mutations in heteroduplex DNA. Proc. Natl. Acad. Sci. U. S. A. 83(10),
3395-3397.
(23) Parker, B. O., and Marinus, M. G. (1992) Repair of DNA heteroduplexes
containing small heterologous sequences in Escherichia coli. Proc. Natl. Acad.
Sci. U. S. A. 89(5), 1730-1734.
(24) Fishel, R. A., Siegel, E. C., and Kolodner, R. (1986) Gene conversion in
Escherichia coli. Resolution of heteroallelic mismatched nucleotides by co-repair.
J. Mol. Biol. 188(2), 147-157.
(25) Fang, W., Wu, J. Y., and Su, M. J. (1997) Methyl-directed repair of mismatched
small heterologous sequences in cell extracts from Escherichia coli. J. Biol.
Chem. 272(36), 22714-22720.
(26) Dohet, C., Dzidic, S., Wagner, R., and Radman, M. (1987) Large non-homology
in heteroduplex DNA is processed differently than single base pair mismatches.
Mol. Gen. Genet. 206(1), 181-184.
(27) Radman, M., and Wagner, R. (1986) Mismatch repair in Escherichia coli. Annu.
Rev. Genet. 20, 523-538.
(28) Worth, L., Jr., Clark, S., Radman, M., and Modrich, P. (1994) Mismatch repair
proteins MutS and MutL inhibit RecA-catalyzed strand transfer between diverged
DNAs. Proc. Natl. Acad. Sci. U. S. A. 91(8), 3238-3241.
(29) Marra, G., and Schar, P. (1999) Recognition of DNA alterations by the mismatch
repair system. Biochem. J. 338(Pt 1), 1-13.
70
- Chapter
-
(30) Lindahl, T. (1979) DNA glycosylases, endonucleases for apurinic/apyrimidinic
sites, and base excision-repair. Prog. Nucleic Acid Res. Mol. Biol. 22, 135-192.
(31) Ehrlich, M., Zhang, X. Y., and Inamdar, N. M. (1990) Spontaneous deamination
of cytosine and 5-methylcytosine residues in DNA and replacement of 5methylcytosine residues with cytosine residues. Mutat. Res. 238(3), 277-286.
(32) May, M. S., and Hattaman, S.
(1975) Deoxyribonucleic acid-cytosine
methylation by host- and plasmid-controlled enzymes. J. Bacteriol. 122(1), 129138.
(33) Marinus, M. G. (1984) Methylation of prokaryotic DNA. In DNA Methylation
(Razin, A., Cedar, H., and Riggs, A. D., Eds.) pp 81-109, Springer-Verlag, New
York.
(34) Marinus, M. G. (1973) Location of DNA methylation genes on the Escherichia
coli K-12 genetic map. Mol. Gen. Genet. 127(1), 47-55.
(35) Coulondre, C., Miller, J. H., Farabaugh, P. J., and Gilbert, W. (1978) Molecular
basis of base substitution hotspots in Escherichia coli. Nature. 274(5673), 775780.
(36) Duncan, B. K., and Miller, J. H. (1980) Mutagenic deamination of cytosine
residues in DNA. Nature. 287(5782), 560-561.
(37) Lieb, M., Allen, E., and Read, D. (1986) Very short patch mismatch repair in
phage lambda: repair sites and length of repair tracts. Genetics. 114(4), 10411060.
(38) Lieb, M. (1981) A fine structure map of spontaneous and induced mutations in
the lambda repressor gene, including insertions of IS elements. Mol. Gen. Genet.
184(3), 364-371.
(39) Lieb, M. (1983) Specific mismatch correction in bacteriophage lambda crosses
by very short patch repair. Mol. Gen. Genet. 191(1), 118-125.
(40) Lieb, M. (1985) Recombination in the lambda repressor gene: evidence that very
short patch (VSP) mismatch correction restores a specific sequence. Mol. Gen.
Genet. 199(3), 465-470.
(41) Jones, M., Wagner, R., and Radman, M.
recombination in E. coli. Cell. 50(4), 621-626.
(1987) Mismatch repair and
(42) Jones, M., Wagner, R., and Radman, M. (1987) Mismatch repair of deaminated
5-methyl-cytosine. J. Mol. Biol. 194(1), 155-159.
71
- Chapter1 (43)
Zell, R., and Fritz, H. J.
(1987) DNA mismatch-repair
in Escherichia
coli
counteracting the hydrolytic deamination of 5-methyl-cytosine residues. EMBO J.
6(6), 1809-1815.
(44) Lieb, M. (1991) Spontaneous mutation at a 5-methylcytosine hotspot is prevented
by very short patch (VSP) mismatch repair. Genetics. 128(1), 23-27.
(45) Lieb, M., and Bhagwat, A. S. (1996) Very short patch repair: reducing the cost of
cytosine methylation. Mol. Microbiol. 20(3), 467-473.
(46) Lieb, M. (1987) Bacterial genes mutL, mutS, and dcm participate in repair of
mismatches at 5-methylcytosine sites. J. Bacteriol. 169(11), 5241-5246.
(47) Dzidic, S., and Radman, M.
(1989) Genetic requirements for hyperrecombination by very short patch mismatch repair: involvement of Escherichia
coli DNA polymerase I. Mol. Gen. Genet. 217(2-3), 254-256.
(48) Sohail, A., Lieb, M., Dar, M., and Bhagwat, A. S. (1990) A gene required for
very short patch repair in Escherichia coli is adjacent to the DNA cytosine
methylase gene. J. Bacteriol. 172(8), 4214-4221.
(49) Hennecke, F., Kolmar, H., Brundl, K., and Fritz, H. J. (1991) The vsr gene
product of E. coli K-12 is a strand- and sequence-specific DNA mismatch
endonuclease. Nature. 353(6346), 776-778.
(50)
Petropoulos,
L., Vidmar, J. J., Passi, E., and Cupples, C. G.
(1994) A simple
assay for monitoring the mutagenic effects of 5-methylcytosine deamination in
Escherichia coli. Mutat. Res. 304(2), 181-185.
(51) Bambara, R. A., Uyemura, D., and Choi, T. (1978) On the processive mechanism
of Escherichia coli DNA polymerase I. Quantitative assessment of processivity. J.
Biol. Chem. 253(2), 413-423.
(52) Palmer, B. R., and Marinus, M. G. (1994) The dam and dcm strains of
Escherichia coli--a review. Gene. 143(1), 1-12.
(53) Takahashi, N., Naito, Y., Handa, N., and Kobayashi, I. (2002) A DNA
methyltransferase can protect the genome from postdisturbance attack by a
restriction-modification gene complex. J. Bacteriol. 184(22), 6100-6108.
(54) Hattman, S., Schlagman, S., and Cousens, L. (1973) Isolation of a mutant of
Escherichia coli defective in cytosine-specific deoxyribonucleic acid methylase
activity and in partial protection of bacteriophage lambda against restriction by
cells containing the N-3 drug-resistance factor. J. Bacteriol. 115(3), 1103-1107.
(55) Schlagman, S., Hattman, S., May, M. S., and Berger, L. (1976) In vivo
methylation by Escherichia coli K-12 mec+ deoxyribonucleic acid-cytosine
72
- Chapter
-
methylase protects against in vitro cleavage by the RII restriction endonuclease
(R. Eco RII). J. Bacteriol. 126(2), 990-996.
(56) Takano, T., Watanabe, T., and Fukasawa, T. (1968) Mechanism ofhost-controlled
restriction of bacteriophage lambda by R factors in Escherichia coli K12.
Virology. 34(2), 290-302.
(57) Roberts, R. J., and Macelis, D. (2001) REBASE--restriction enzymes and
methylases. Nucleic Acids Res. 29(1), 268-269.
(58) Miki, T., Ebina, Y., Kishi, F., and Nakazawa, A. (1981) Organization of the lexA
gene of Escherichia coli and nucleotide sequence of the regulatory region. Nucleic
Acids Res. 9(3), 529-543.
(59) Ringquist, S., and Smith, C. L. (1992) The Escherichia coli chromosome contains
specific, unmethylated dam and dcm sites. Proc. Natl. Acad. Sci. U. S. A. 89(10),
4539-4543.
(60) Lindahl, T., and Nyberg, B. (1974) Heat-induced deamination of cytosine
residues in deoxyribonucleic acid. Biochemistry. 13(16), 3405-3410.
(61) Ehrlich, M., Norris, K. F., Wang, R. Y., Kuo, K. C., and Gehrke, C. W. (1986)
DNA cytosine methylation and heat-induced deamination. Biosci. Rep. 6(4), 387393.
(62) Shapiro, R., and Klein, R. S. (1966) The deamination of cytidine and cytosine by
acidic buffer solutions. Mutagenic implications. Biochemistry. 5(7), 2358-2362.
(63) Shapiro, R. (1981) Damage to DNA caused by hydrolysis. In Chromosome
Damage and Repair pp 3-12, Plenum Publishing Corp., New York.
(64) BROWN, D. M., and PHILLIPS, J. H. (1965) MECHANISM OF THE
MUTAGENIC ACTION OF HYDROXYLAMINE. J. Mol. Biol. 11:663-71.,
663-671.
(65) Chambers, R. W., and Kurkov, V. (1963) The Structure and Properties of some
Sulfur Analogs Acetoneuridine. J. Am. Chem. Soc. 85, 2160-2164.
(66) Johns, H., Le Blanc, J. C., and Freeman K.B. (1965) J. Mol. Biol. 13, 849.
(67) Lutsenko, E., and Bhagwat, A. S. (1999) Principal causes of hot spots for
cytosine to thymine mutations at sites of cytosine methylation in growing cells. A
model, its experimental support and implications. Mutat. Res. 437(1), 11-20.
(68) Lieb, M., and Rehmat, S. (1997) 5-Methylcytosine is not a mutation hot spot in
nondividing Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 94(3), 940-945.
73
- Chapter1 (69) Macintyre, G., Pitsikas, P., and Cupples, C. G. (1999) Growth phase-dependent
regulation of Vsr endonuclease may contribute to 5-methylcytosine mutational
hot spots in Escherichia coli. J. Bacteriol. 181(14), 4435-4436.
(70) Bell, D. C., and Cupples, C. G. (2001) Very-short-patch repair in Escherichia coli
requires the dam adenine methylase. J. Bacteriol. 183(12), 3631-3635.
(71) Dar, M. E., and Bhagwat, A. S. (1993) Mechanism of expression of DNA repair
gene vsr, an Escherichia coli gene that overlaps the DNA cytosine methylase
gene, dcm. Mol. Microbiol. 9(4), 823-833.
(72) Bhagwat, A. S., and Lieb, M. (2002) Cooperation and competition in mismatch
repair: very short-patch repair and methyl-directed mismatch repair in Escherichia
coli. Mol. Microbiol. 44(6), 1421-1428.
(73) Ehrlich, M., and Wang, R. Y.
Science. 212(4501), 1350-1357.
(1981) 5-Methylcytosine in eukaryotic DNA.
(74) Colot, V., and Rossignol, J. L. (1999) Eukaryotic DNA methylation as an
evolutionary device. Bioessays. 21(5), 402-411.
(75) Cooper, D. N., and Youssoufian, H. (1988) The CpG dinucleotide and human
genetic disease. Hum. Genet. 78(2), 151-155.
(76) Giannelli, F., Green, P. M., High, K. A., Lozier, J. N., Lillicrap, D. P., Ludwig,
M., Olek, K., Reitsma, P. H., Goossens, M., Yoshioka, A., and . (1990)
Haemophilia B: database of point mutations and short additions and deletions.
Nucleic Acids Res. 18(14), 4053-4059.
(77) Jones, P. A., Rideout, W. M., III, Shen, J. C., Spruck, C. H., and Tsai, Y. C.
(1992) Methylation, mutation and cancer. Bioessays. 14(1), 33-36.
(78)
Koeberl, D. D., Bottema, C. D., Ketterling, R. P., Bridge, P. J., Lillicrap, D. P.,
and Sommer, S. S. (1990) Mutations causing hemophilia B: direct estimate of the
underlying rates of spontaneous germ-line transitions, transversions, and deletions
in a human gene. Am. J. Hum. Genet. 47(2), 202-217.
(79) Hirschhorn, R., Tzall, S., and Ellenbogen, A. (1990) Hot spot mutations in
adenosine deaminase deficiency. Proc. Natl. Acad. Sci. U. S. A. 87(16), 61716175.
(80) Lieb, M., and Rehmat, S. (1995) Very short patch repair of T:G mismatches in
vivo: importance of context and accessory proteins. J. Bacteriol. 177(3), 660-666.
(81)
Glasner, W., Merkl, R., Schellenberger,
V., and Fritz, H. J.
(1995) Substrate
preferences of Vsr DNA mismatch endonuclease and their consequences for the
evolution of the Escherichia coli K-12 genome. J. Mol. Biol. 245(1), 1-7.
74
- Chapter1 (82)
Bhag'wat, A. S., and McClelland, M. (1992) DNA mismatch correction by Very
Short Patch repair may have altered the abundance of oligonucleotides in the E.
coli genome. Nucleic Acids Res. 20(7), 1663-1668.
(83) Merkl, R., Kroger, M., Rice, P., and Fritz, H. J. (1992) Statistical evaluation and
biological interpretation of non-random abundance in the E. coli K-12 genome of
tetra- and pentanucleotide sequences related to VSP DNA mismatch repair.
Nucleic Acids Res. 20(7), 1657-1662.
(84) Gabbara, S., Wyszynski, M., and Bhagwat, A. S. (1994) A DNA repair process in
Escherichia coli corrects U:G and T:G mismatches to C:G at sites of cytosine
methylation. Mol. Gen. Genet. 243(2), 244-248.
(85) Schaaper, R. M., and Radman, M. (1989) The extreme mutator effect of
Escherichia coli mutD5 results from saturation of mismatch repair by excessive
DNA replication errors. EMBOJ. 8(11), 3511-3516.
(86) Wu, J. C., and Santi, D. V. (1987) Kinetic and catalytic mechanism of HhaI
methyltransferase. J. Biol. Chem. 262(10), 4778-4786.
(87) Wyszynski, M., Gabbara, S., and Bhagwat, A. S. (1994) Cytosine deaminations
catalyzed by DNA cytosine methyltransferases are unlikely to be the major cause
of mutational hot spots at sites of cytosine methylation in Escherichia coli. Proc.
Natl. Acad. Sci. U. S. A. 91(4), 1574-1578.
(88)
Shen., J. C., Rideout, W. M., III, and Jones, P. A.
(1992) High frequency
mutagenesis by a DNA methyltransferase. Cell. 71(7), 1073-1080.
(89) Turner, D. P., and Connolly, B. A. (2000) Interaction of the E. coli DNA G:Tmismatch endonuclease (vsr protein) with oligonucleotides containing its target
sequence. J. Mol. Biol. 304(5), 765-778.
(90) Gonzalez-Nicieza, R., Turner, D. P., and Connolly, B. A. (2001) DNA binding
and cleavage selectivity of the Escherichia coli DNA G:T-mismatch endonuclease
(vsr protein). J. Mol. Biol. 310(3), 501-508.
(91)
Tsutakawa,
S. E., Jingami, H., and Morikawa, K.
(1999) Recognition
of a TG
mismatch: the crystal structure of very short patch repair endonuclease in
complex with a DNA duplex. Cell. 99(6), 615-623.
(92)
Bunting, K. A., Roe, S. M., Headley, A., Brown, T., Savva, R., and Pearl, L. H.
(2003) Crystal structure of the Escherichia coli dcm very-short-patch DNA repair
endonuclease bound to its reaction product-site in a DNA superhelix. Nucleic
Acids Res. 31(6), 1633-1639.
(93) Elliott, S. L., Brazier, J., Cosstick, R., and Connolly, B. A. (2005) Mechanism of
the Escherichia coli DNA T:G-mismatch endonuclease (Vsr protein) probed with
thiophosphate-containing oligodeoxynucleotides. J. Mol. Biol. 353(3), 692-703.
75
- Chapter1 (94) Monastiriakos, S. K., Doiron, K. M., Siponen, M. I., and Cupples, C. G. (2004)
Functional interactions between the MutL and Vsr proteins of Escherichia coli are
dependent on the N-terminus of Vsr. DNA Repair (Amst). 3(6), 639-647.
(95) Fehrst, A. Enzyme Structure and Mechanism (1984) W. H. Freeman and
Company, New York.
(96) Jen-Jacobson, L. (1995) Structural-perturbation approaches to thermodynamics
of site-specific protein-DNA interactions. Methods Enzymol. 259, 305-344.
(97) Jen-Jacobson, L. (1997) Protein-DNA recognition complexes: conservation of
structure and binding energy in the transition state. Biopolymers. 44(2), 153-180.
(98) Tsutakawa, S. E., Muto, T., Kawate, T., Jingami, H., Kunishima, N., Ariyoshi,
M., IKohda,D., Nakagawa, M., and Morikawa, K. (1999) Crystallographic and
functional studies of very short patch repair endonuclease. Mol. Cell. 3(5), 621628.
(99) Tsutakawa, S. E., and Morikawa, K. (2001) The structural basis of damaged
DNA recognition and endonucleolytic cleavage for very short patch repair
endonuclease. Nucleic Acids Res. 29(18), 3775-3783.
(100) Pingoud, A., and Jeltsch, A. (2001) Structure and function of type II restriction
endonucleases. Nucleic Acids Res. 29(18), 3705-3727.
(101) Ban, C., and Yang, W. (1998) Structural basis for MutH activation in E.coli
mismatch repair and relationship of MutH to restriction endonucleases. EMBO J.
17(5), 1526-1534.
(102) Kovall, R., and Matthews, B. W. (1997) Toroidal structure of lambdaexonuclease. Science. 277(5333), 1824-1827.
(103) Nishino, T., Komori, K., Tsuchiya, D., Ishino, Y., and Morikawa, K. (2001)
Crystal structure of the archaeal holliday junction resolvase Hjc and implications
for DNA recognition. Structure. 9(3), 197-204.
(104) Aggarwal, A. K. (1995) Structure and function of restriction endonucleases.
Curr. Opin. Struct. Biol. 5(1), 11-19.
(105) Pingoud, A., and Jeltsch, A. (1997) Recognition and cleavage of DNA by type-II
restriction endonucleases. Eur. J. Biochem. 246(1), 1-22.
(106) Jeltsch, A., Alves, J., Maass, G., and Pingoud, A. (1992) On the catalytic
mechanism of EcoRI and EcoRV. A detailed proposal based on biochemical
results, structural data and molecular modelling. FEBS Lett. 304(1), 4-8.
76
- ChapterI (107) Jeltsch, A., Alves, J., Wolfes, H., Maass, G., and Pingoud, A. (1993) Substrateassisted catalysis in the cleavage of DNA by the EcoRI and EcoRV restriction
enzymes. Proc. Natl. Acad. Sci. U. S. A. 90(18), 8499-8503.
(108) Horton, N. C., Newberry, K. J., and Perona, J. J. (1998) Metal ion-mediated
substrate-assisted catalysis in type II restriction endonucleases. Proc. Natl. Acad.
Sci. U. S. A. 95(23), 13489-13494.
(109) Lukacs, C. M., Kucera, R., Schildkraut, I., and Aggarwal, A. K. (2000)
Understanding the immutability of restriction enzymes: crystal structure of BglII
and its DNA substrate at 1.5 A resolution. Nat. Struct. Biol. 7(2), 134-140.
(110) Aishima, J., Gitti, R. K., Noah, J. E., Gan, H. H., Schlick, T., and Wolberger, C.
(2002) A Hoogsteen base pair embedded in undistorted B-DNA. Nucleic Acids
Res. 30(23), 5244-5252.
(111) Weston, S. A., Lahm, A., and Suck, D. (1992) X-ray structure of the DNase Id(GGTATACC)2 complex at 2.3 A resolution. J. Mol. Biol. 226(4), 1237-1256.
(112) Mol, C. D., Kuo, C. F., Thayer, M. M., Cunningham, R. P., and Tainer, J. A.
(1995) Structure and function of the multifunctional DNA-repair enzyme
exonuclease III. Nature. 374(6520), 381-386.
(113)
Gorman, M. A., Morera, S., Rothwell, D. G., de La, F. E., Mol, C. D., Tainer, J.
A., Hickson, I. D., and Freemont, P. S. (1997) The crystal structure of the human
DNA repair endonuclease HAP1 suggests the recognition of extra-helical
deoxyribose at DNA abasic sites. EMBO J. 16(21), 6548-6558.
(114) Pingoud, A., Fuxreiter, M., Pingoud, V., and Wende, W. (2005) Type II
restriction endonucleases: structure and mechanism. Cell Mol. Life Sci. 62(6),
685-707.
(115) Eckstein, F. (1983) Phosphorothioate analogues of nucleotides - tools for the
investigation of biochemical processes. Angew. Chem. Int. Ed. Engl. 22, 423-439.
(116) Frey, P. A. (1989) Chiral phosphorothioates: stereochemical analysis of
enzymatic substitution at phosphorus. Adv. Enzymol. Relat Areas Mol. Biol. 62,
119-201.
(117) Gerlt, J. (1993) Mechanistic principles of enzyme-catalyzed cleavage of
phosphodiester bonds. In Nucleases (Linn, S. M., Lloyd, R. S., and Roberts, S. J.,
Eds.) pp 1-34, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
(118) Claverys, J. P., and Lacks, S. A. (1986) Heteroduplex deoxyribonucleic acid base
mismatch repair in bacteria. Microbiol. Rev. 50(2), 133-165.
77
- Chapter
I(119) Lacks, S.
(1970) Mutants of Diplococcus pneumoniae that lack
deoxyribonucleases and other activities possibly pertinent to genetic
transformation. J. Bacteriol. 101(2), 373-383.
(120) Tiraby, J. G., and Fox, M. S. (1973) Marker discrimination in transformation and
mutation of pneumococcus. Proc. Natl. Acad. Sci. U. S. A. 70(12), 3541-3545.
(121) Guild, W. R., and Shoemaker, N. B.
(1976) Mismatch correction in
pneumococcal transformation: donor length and hex-dependent marker efficiency.
J. Bacteriol. 125(1), 125-135.
(122) Litman, R. M. (1961) Genetic and chemical alterations in the transforming DNA
of pneumococcus caused by ultraviolet light and nitrous acid. J. Chim. Phys. 58,
997-1003.
(123) Claverys, J. P., Roger, M., and Sicard, A. M. (1980) Excision and repair of
mismatched base pairs in transformation of Streptococcus pneumoniae. Mol. Gen.
Genet. 178(1), 191-201.
(124) Fox, M. S. (1978) Some features of genetic recombination in procaryotes. Annu.
Rev. Genet. 12, 47-68.
(125) Lacks, S. A., Dunn, J. J., and Greenberg, B. (1982) Identification of base
mismatches recognized by the heteroduplex-DNA-repair system of Streptococcus
pneumoniae. Cell. 31(2 Pt 1), 327-336.
(126) Kornberg, A., and Baker, T. A. DNA Replication (1991) W. H. Freeman and Co.,
New York.
(127)
Haber, L. T., Pang, P. P., Sobell, D. I., Mankovich,
J. A., and Walker, G. C.
(1988) Nucleotide sequence of the Salmonella typhimurium mutS gene required
for mismatch repair: homology of MutS and HexA of Streptococcus pneumoniae.
J. Bacteriol. 170(1), 197-202.
(128) Wagner, R., Jr., and Meselson, M. (1976) Repair tracts in mismatched DNA
heteroduplexes. Proc. Natl. Acad. Sci. U. S. A. 73(11), 4135-4139.
(129)
Radman, M., Wagner, R. E., Glickman, B. W., and Meselson, M. (1980) DNA
methylation, mismatch correction and genetic stability. In Progress in
Environmental Mutagenesis (Alacevic, M., Ed.) pp 121-130, Elsevier/NorthHolland Biomedical Press, Amsterdam.
(130) Lahue, R. S., Su, S. S., and Modrich, P. (1987) Requirement for d(GATC)
sequences in Escherichia coli mutHLS mismatch correction. Proc. Natl. Acad.
Sci. U. S. A. 84(6), 1482-1486.
(131)
Lahue, R. S., Au, K. G., and Modrich, P. (1989) DNA mismatch correction in a
defined system. Science. 245(4914), 160-164.
78
-Chapter
I(132) Au, K. G., Welsh, K., and Modrich, P. (1992) Initiation of methyl-directed
mismatch repair. J. Biol. Chem. 267(17), 12142-12148.
(133) Lyons, S. M., and Schendel, P. F. (1984) Kinetics of methylation in Escherichia
coli K-12. J. Bacteriol. 159(1), 421-423.
(134) Marinus, M. G. (1976) Adenine methylation of Okazaki fragments in Escherichia
coli. J. Bacteriol. 128(3), 853-854.
(135) Yamaguchi, M., Dao, V., and Modrich, P. (1998) MutS and MutL activate DNA
helicase II in a mismatch-dependent manner. J. Biol. Chem. 273(15), 9197-9201.
(136) Dao, V., and Modrich, P. (1998) Mismatch-, MutS-, MutL-, and helicase IIdependent unwinding from the single-strand break of an incised heteroduplex. J.
Biol. Chem. 273(15), 9202-9207.
(137) Grilley, M., Griffith, J., and Modrich, P. (1993) Bidirectional excision in methyldirected mismatch repair. J. Biol. Chem. 268(16), 11830-11837.
(138) Cooper, D. N. (1998) The nature and mechanisms of human gene mutation. In
The Genetic Basis of Human Cancer (Vogelstein, B., and Kinzler, K. W., Eds.) pp
65-94, McGraw-Hill, New York, N.Y.
(139) Viswanathan, M., and Lovett, S. T. (1999) Exonuclease X of Escherichia coli. A
novel 3'-5' DNase and Dnaq superfamily member involved in DNA repair. J. Biol.
Chem. 274(42), 30094-30100.
(140) Chase, J. W., and Richardson, C. C. (1974) Exonuclease VII of Escherichia coli.
Mechanism of action. J. Biol. Chem. 249(14), 4553-4561.
(141)
Burdett, V., Baitinger,
C., Viswanathan,
M., Lovett, S. T., and Modrich,
P.
(2001) In vivo requirement for RecJ, ExoVII, ExoI, and ExoX in methyl-directed
mismatch repair. Proc. Natl. Acad. Sci. U. S. A. 98(12), 6765-6770.
(142) Viswanathan, M., Burdett, V., Baitinger, C., Modrich, P., and Lovett, S. T.
(2001) Redundant exonuclease involvement in Escherichia coli methyl-directed
mismatch repair. J. Biol. Chem. 276(33), 31053-31058.
(143)
Cooper, D. L., Lahue, R. S., and Modrich, P. (1993) Methyl-directed
mismatch
repair is bidirectional. J. Biol. Chem. 268(16), 11823-11829.
(144) Lu, A. L. (1987) Influence of GATC sequences on Escherichia coli DNA
mismatch repair in vitro. J. Bacteriol. 169(3), 1254-1259.
(145) Modrich, P. (1989) Methyl-directed DNA mismatch correction. J. Biol. Chem.
264(12), 6597-6600.
79
- Chapter
-
(146) Meselson, M.
(1988) Methyl-directed repair of DNA mismatches. In
Recombination of the Genetic Material (Low, K. B., Ed.) pp 91-113, Academic
Press, Inc., San Diego, Calif.
(147) Blattner, F. R., Plunkett, G., III, Bloch, C. A., Perna, N. T., Burland, V., Riley,
M., Collado-Vides,
J., Glasner, J. D., Rode, C. K., Mayhew, G. F., Gregor, J.,
Davis, N. W., Kirkpatrick, H. A., Goeden, M. A., Rose, D. J., Mau, B., and Shao,
Y. (1997) The complete genome sequence of Escherichia coli K-12. Science.
277(5331), 1453-1474.
(148) Marinus, M. G., and Morris, N. R. (1973) Isolation of deoxyribonucleic acid
methylase mutants of Escherichia coli K-12. J. Bacteriol. 114(3), 1143-1150.
(149) Marinus, M. G., and Morris, N. R. (1975) Pleiotropic effects of a DNA adenine
methylation mutation (dam-3) in Escherichia coli K12. Mutat. Res. 28(1), 15-26.
(150) Kramer, B., Kramer, W., and Fritz, H. J. (1984) Different base/base mismatches
are corrected with different efficiencies by the methyl-directed DNA mismatchrepair system of E. coli. Cell. 38(3), 879-887.
(151) Lu, A. L., Clark, S., and Modrich, P. (1983) Methyl-directed repair of DNA basepair mismatches in vitro. Proc. Natl. Acad. Sci. U. S. A. 80(15), 4639-4643.
(152) Pukkila, P. J., Peterson, J., Herman, G., Modrich, P., and Meselson, M. (1983)
Effects of high levels of DNA adenine methylation on methyl-directed mismatch
repair in Escherichia coli. Genetics. 104(4), 571-582.
(153) Laengle-Rouault, F., Maenhaut-Michel, G., and Radman, M. (1986) GATC
sequence and mismatch repair in Escherichia coli. EMBO J. 5(8), 2009-2013.
(154) Herman, G. E., and Modrich, P. (1981) Escherichia coli K-12 clones that
overproduce dam methylase are hypermutable. J. Bacteriol. 145(1), 644-646.
(155) Marinus, M. G., Poteete, A., and Arraj, J. A. (1984) Correlation of DNA adenine
methylase activity with spontaneous mutability in Escherichia coli K-12. Gene.
28(1), 123-125.
(156) Szyf, M., vraham-Haetzni, K., Reifman, A., Shlomai, J., Kaplan, F., Oppenheim,
A., and Razin, A. (1984) DNA methylation pattern is determined by the
intracellular level of the methylase. Proc. Natl. Acad. Sci. U. S. A. 81(11), 32783282.
(157) Schlensog, V., and Bock, A. (1991) The Escherichia coli fdv gene probably
encodes mutS and is located at minute 58.8 adjacent to the hyc-hyp gene cluster.
J. Bacteriol. 173(23), 7414-7415.
80
-Chapter
I(158) Su, S. S., Lahue, R. S., Au, K. G., and Modrich, P. (1988) Mispair specificity of
methyl-directed DNA mismatch correction in vitro. J. Biol. Chem. 263(14), 68296835.
(159) Walker, J. E., Saraste, M., Runswick, M. J., and Gay, N. J. (1982) Distantly
related sequences in the alpha- and beta-subunits of ATP synthase, myosin,
kinases and other ATP-requiring enzymes and a common nucleotide binding fold.
EMBO J. 1(8), 945-951.
(160) Su, S. S., and Modrich, P. (1986) Escherichia coli mutS-encoded protein binds to
mismatched DNA base pairs. Proc. Natl. Acad. Sci. U. S. A. 83(14), 5057-5061.
(161) Grilley, M., Welsh, K. M., Su, S. S., and Modrich, P. (1989) Isolation and
characterization of the Escherichia coli mutL gene product. J. Biol. Chem. 264(2),
1000-1004.
(162) Welsh, K. M., Lu, A. L., Clark, S., and Modrich, P. (1987) Isolation and
characterization of the Escherichia coli mutH gene product. J. Biol. Chem.
262(32), 15624-15629.
(163) Haber, L. T., and Walker, G. C. (1991) Altering the conserved nucleotide binding
motif in the Salmonella typhimurium MutS mismatch repair protein affects both
its ATPase and mismatch binding activities. EMBO J. 10(9), 2707-2715.
(164) Wu, T. H., and Marinus, M. G. (1994) Dominant negative mutator mutations in
the mutS gene of Escherichia coli. J. Bacteriol. 176(17), 5393-5400.
(165) Fishel, R., Acharya, S., Berardini, M., Bocker, T., Charbonneau, N., Cranston, A.,
Gradia, S., Guerrette, S., Heinen, C. D., Mazurek, A., Snowden, T., Schmutte, C.,
Shim, K. S., Tombline, G., and Wilson, T. (2000) Signaling mismatch repair: the
mechanics of an adenosine-nucleotide molecular switch. Cold Spring Harb. Symp.
Quant. Biol. 65, 217-224.
(166) Gradia, S., Acharya, S., and Fishel, R. (2000) The role of mismatched
nucleotides in activating the hMSH2-hMSH6 molecular switch. J. Biol. Chem.
275(6), 3922-3930.
(167)
Schofield, M. J., Brownewell, F. E., Nayak, S., Du, C., Kool, E. T., and Hsieh, P.
(2001) The Phe-X-Glu DNA binding motif of MutS. The role of hydrogen
bonding in mismatch recognition. J. Biol. Chem. 276(49), 45505-45508.
(168) Jones, M., Wagner, R., and Radman, M. (1987) Repair of a mismatch is
influenced by the base composition of the surrounding nucleotide sequence.
Genetics. 115(4), 605-610.
(169) Obmolova, G., Ban, C., Hsieh, P., and Yang, W. (2000) Crystal structures of
mismatch repair protein MutS and its complex with a substrate DNA. Nature.
407(6805), 703-710.
81
- Chapter1 (170) Lamers, M. H., Winterwerp, H. H., and Sixma, T. K. (2003) The alternating
ATPase domains of MutS control DNA mismatch repair. EMBO J. 22(3), 746756.
(171) Lamers, M. H., Perrakis, A., Enzlin, J. H., Winterwerp, H. H., de, W. N., and
Sixma, T. K. (2000) The crystal structure of DNA mismatch repair protein MutS
binding to a G x T mismatch. Nature. 407(6805), 711-717.
(172) Junop, M. S., Obmolova, G., Rausch, K., Hsieh, P., and Yang, W. (2001)
Composite active site of an ABC ATPase: MutS uses ATP to verify mismatch
recognition and authorize DNA repair. Mol. Cell. 7(1), 1-12.
(173) Alani, E., Lee, J. Y., Schofield, M. J., Kijas, A. W., Hsieh, P., and Yang, W.
(2003) Crystal structure and biochemical analysis of the MutS.ADP.beryllium
fluoride complex suggests a conserved mechanism for ATP interactions in
mismatch repair. J. Biol. Chem. 278(18), 16088-16094.
(174)
Lamers, M. H., Georgijevic, D., Lebbink, J. H., Winterwerp, H. H., Agianian, B.,
de, W. N., and Sixma, T. K. (2004) ATP increases the affinity between MutS
ATPase domains. Implications for ATP hydrolysis and conformational changes. J.
Biol. Chem. 279(42), 43879-43885.
(175) Natrajan, G., Lamers, M. H., Enzlin, J. H., Winterwerp, H. H., Perrakis, A., and
Sixma, T. K. (2003) Structures of Escherichia coli DNA mismatch repair enzyme
MutS in complex with different mismatches: a common recognition mode for
diverse substrates. Nucleic Acids Res. 31(16), 4814-4821.
(176) Bjornson, K. P., Allen, D. J., and Modrich, P. (2000) Modulation of MutS ATP
hydrolysis by DNA cofactors. Biochemistry. 39(11), 3176-3183.
(177) Allen, D. J., Makhov, A., Grilley, M., Taylor, J., Thresher, R., Modrich, P., and
Griffith, J. D. (1997) MutS mediates heteroduplex loop formation by a
translocation mechanism. EMBO J. 16(14), 4467-4476.
(178) Sixma, T. K. (2001) DNA mismatch repair: MutS structures bound to
mismatches. Curr. Opin. Struct. Biol. 11(1), 47-52.
(179) Lopez de Saro, F. J., and O'Donnell, M. (2001) Interaction of the beta sliding
clamp with MutS, ligase, and DNA polymerase I. Proc. Natl. Acad. Sci. U. S. A.
98(15), 8376-8380.
(180)
Lu, A. L., Welsh, K., Clark, S., Su, S. S., and Modrich, P. (1984) Repair of DNA
base-pair mismatches in extracts of Escherichia coli. Cold Spring Harb. Symp.
Quant. Biol. 49, 589-596.
(181)
Guame, A., Ramon-Maiques,
S., Wolff, E. M., Ghirlando, R., Hu, X., Miller, J.
H., and Yang, W. (2004) Structure of the MutL C-terminal domain: a model of
intact MutL and its roles in mismatch repair. EMBO J. 23(21), 4134-4145.
82
-Chapter
I(182) Acharya, S., Foster, P. L., Brooks, P., and Fishel, R. (2003) The coordinated
functions of the E. coli MutS and MutL proteins in mismatch repair. Mol. Cell.
12(1), 233-246.
(183) Ban, C., and Yang, W. (1998) Crystal structure and ATPase activity of MutL:
implications for DNA repair and mutagenesis. Cell. 95(4), 541-552.
(184) Ban, C., Junop, M., and Yang, W. (1999) Transformation of MutL by ATP
binding and hydrolysis: a switch in DNA mismatch repair. Cell. 97(1), 85-97.
(185) Aronshtam, A., and Marinus, M. G. (1996) Dominant negative mutator mutations
in the mutL gene of Escherichia coli. Nucleic Acids Res. 24(13), 2498-2504.
(186) Spampinato, C., and Modrich, P. (2000) The MutL ATPase is required for
mismatch repair. J. Biol. Chem. 275(13), 9863-9869.
(187) Sancar, A., and Hearst, J. E. (1993) Molecular matchmakers. Science. 259(5100),
1415-1420.
(188) Hu, X., Machius, M., and Yang, W. (2003) Monovalent cation dependence and
preference of GHKL ATPases and kinases. FEBSLett. 544(1-3), 268-273.
(189) Lee, J. Y., Chang, J., Joseph, N., Ghirlando, R., Rao, D. N., and Yang, W. (2005)
MutH complexed with hemi- and unmethylated DNAs: coupling base recognition
and DNA cleavage. Mol. Cell. 20(1), 155-166.
(190) Glickman, B. W., and Radman, M. (1980) Escherichia coli mutator mutants
deficient in methylation-instructed DNA mismatch correction. Proc. Natl. Acad.
Sci. U. S. A. 77(2), 1063-1067.
(191) Grafstrom, R. H., Amsterdam, A., and Zachariasewycz, K. (1988) In vivo studies
of repair of 2-aminopurine in Escherichia coli. J. Bacteriol. 170(8), 3485-3492.
(192) Blackwell, L. J., Martik, D., Bjornson, K. P., Bjornson, E. S., and Modrich, P.
(1998) Nucleotide-promoted release of hMutSalpha from heteroduplex DNA is
consistent with an ATP-dependent translocation mechanism. J. Biol. Chem.
273(48), 32055-32062.
(193) Martik, D., Baitinger, C., and Modrich, P. (2004) Differential specificities and
simultaneous occupancy of human MutSalpha nucleotide binding sites. J. Biol.
CheR. 279(27), 28402-28410.
(194) Blackwell, L. J., Bjornson, K. P., Allen, D. J., and Modrich, P. (2001) Distinct
MutS DNA-binding modes that are differentially modulated by ATP binding and
hydrolysis. J. Biol. Chem. 276(36), 34339-34347.
83
- Chapter
-
(195) Blackwell, L. J., Wang, S., and Modrich, P. (2001) DNA chain length
dependence of formation and dynamics of hMutSalpha.hMutLalpha.heteroduplex
complexes. J. Biol. Chem. 276(35), 33233-33240.
(196) Gradia, S., Acharya, S., and Fishel, R. (1997) The human mismatch recognition
complex hMSH2-hMSH6 functions as a novel molecular switch. Cell. 91(7), 9951005.
(197) Wilson, T., Guerrette, S., and Fishel, R. (1999) Dissociation of mismatch
recognition and ATPase activity by hMSH2-hMSH3. J. Biol. Chem. 274(31),
21659-21664.
(198) Gradia, S., Subramanian, D., Wilson, T., Acharya, S., Makhov, A., Griffith, J.,
and Fishel, R. (1999) hMSH2-hMSH6 forms a hydrolysis-independent sliding
clamp on mismatched DNA. Mol. Cell. 3(2), 255-261.
(199) Heinen, C. D., Wilson, T., Mazurek, A., Berardini, M., Butz, C., and Fishel, R.
(2002) HNPCC mutations in hMSH2 result in reduced hMSH2-hMSH6 molecular
switch functions. Cancer Cell. 1(5), 469-478.
(200) Schofield, M. J., and Hsieh, P. (2003) DNA mismatch repair: molecular
mechanisms and biological function. Annu. Rev. Microbiol. 57, 579-608.
(201) Culligan, K. M., and Hays, J. B. (1997) DNA mismatch repair in plants. An
Arabidopsis thaliana gene that predicts a protein belonging to the MSH2
subfamily of eukaryotic MutS homologs. Plant Physiol. 115(2), 833-839.
(202) Larson, E. D., Nickens, D., and Drummond, J. T. (2002) Construction and
characterization of mismatch-containing circular DNA molecules competent for
assessment of nick-directed human mismatch repair in vitro. Nucleic Acids Res.
30(3), E14.
(203) Wang, H., and Hays, J. B. (2004) Signaling from DNA mispairs to mismatchrepair excision sites despite intervening blockades. EMBO J. 23(10), 2126-2133.
(204) Lee, S. D., and Alani, E. (2006) Analysis of interactions between mismatch
repair initiation factors and the replication processivity factor PCNA. J. Mol. Biol.
355(2), 175-184.
(205) Marti, T. M., Kunz, C., and Fleck, 0. (2002) DNA mismatch repair and mutation
avoidance pathways. J. Cell Physiol. 191(1), 28-41.
(206) Lieb, M., Rehmat, S., and Bhagwat, A. S. (2001) Interaction of MutS and Vsr:
some dominant-negative mutS mutations that disable methyladenine-directed
mismatch repair are active in very-short-patch repair. J. Bacteriol. 183(21), 64876490.
84
-Chapter
I(207) Doiron, K. M., Viau, S., Koutroumanis, M., and Cupples, C. G. (1996)
Overexpression of vsr in Escherichia coli is mutagenic. J. Bacteriol. 178(14),
4294-4296.
(208) Macintyre, G., Doiron, K. M., and Cupples, C. G. (1997) The Vsr endonuclease
of Escherichia coli: an efficient DNA repair enzyme and a potent mutagen. J.
Bacteriol. 179(19), 6048-6052.
(209) Drotschmann, K., Aronshtam, A., Fritz, H. J., and Marinus, M. G. (1998) The
Escherichia coli MutL protein stimulates binding of Vsr and MutS to
heteroduplex DNA. Nucleic Acids Res. 26(4), 948-953.
(210) Mansour, C. A., Doiron, K. M., and Cupples, C. G. (2001) Characterization of
functional interactions among the Escherichia coli mismatch repair proteins using
a bacterial two-hybrid assay. Mutat. Res. 485(4), 331-338.
(211) Morikawa, K., and Shirakawa, M. (2000) Three-dimensional structural views of
damaged-DNA recognition: T4 endonuclease V, E. coli Vsr protein, and human
nucleotide excision repair factor XPA. Mutat. Res. 460(3-4), 257-275.
(212) Feng, G., Tsui, H. C., and Winkler, M. E. (1996) Depletion of the cellular
amounts of the MutS and MutH methyl-directed mismatch repair proteins in
stationary-phase Escherichia coli K-12 cells. J. Bacteriol. 178(8), 2388-2396.
(213)
Harris, R. S., Feng, G., Ross, K. J., Sidhu, R., Thulin, C., Longerich, S., Szigety,
S. K., Winkler, M. E., and Rosenberg, S. M. (1997) Mismatch repair protein
MutL becomes limiting during stationary-phase mutation. Genes Dev. 11(18),
2426-2437.
(214) Harris, R. S., Feng, G., Ross, K. J., Sidhu, R., Thulin, C., Longerich, S., Szigety,
S. K., Hastings, P. J., Winkler, M. E., and Rosenberg, S. M. (1999) Mismatch
repair is diminished during stationary-phase mutation. Mutat. Res. 437(1), 51-60.
(215) Wenthzel, A. M., Stancek, M., and Isaksson, L. A. (1998) Growth phase
dependent stop codon readthrough and shift of translation reading frame in
Escherichia coli. FEBS Lett. 421(3), 237-242.
(216) Neddermann, P., and Jiricny, J. (1993) The purification of a mismatch-specific
thymine-DNA glycosylase from HeLa cells. J. Biol. Chem. 268(28), 2121821224.
(217) Neddermann, P., and Jiricny, J. (1994) Efficient removal of uracil from G.U
mispairs by the mismatch-specific thymine DNA glycosylase from HeLa cells.
Proc. Natl. Acad. Sci. U. S. A. 91(5), 1642-1646.
(218) Pearl, L. H. (2000) Structure and function in the uracil-DNA glycosylase
superfamily. Mutat. Res. 460(3-4), 165-181.
85
- Chapter1 (219) Gallinari, P., and Jiricny, J. (1996) A new class of uracil-DNA glycosylases
related to human thymine-DNA glycosylase. Nature. 383(6602), 735-738.
(220) Saparbaev, M., and Laval, J. (1998) 3,N4-ethenocytosine, a highly mutagenic
adduct, is a primary substrate for Escherichia coli double-stranded uracil-DNA
glycosylase and human mismatch-specific thymine-DNA glycosylase. Proc. Natl.
Acad Sci. U. S. A. 95(15), 8508-8513.
(221) Lutsenko, E., and Bhagwat, A. S. (1999) The role of the Escherichia coli mug
protein in the removal of uracil and 3,N(4)-ethenocytosine from DNA. J. Biol.
Chem. 274(43), 31034-31038.
(222) Sung, J. S., and Mosbaugh, D. W. (2000) Escherichia coli double-strand uracilDNA glycosylase: involvement in uracil-mediated DNA base excision repair and
stimulation of activity by endonuclease IV. Biochemistry. 39(33), 10224-10235.
(223) Hendrich, B., Hardeland, U., Ng, H. H., Jiricny, J., and Bird, A. (1999) The
thymine glycosylase MBD4 can bind to the product of deamination at methylated
CpG sites. Nature. 401(6750), 301-304.
(224) Friedberg, E. C., Walker, G. C., and Siede, W. DNA Repair and Mutagenesis
(1995) ASM Press, Washington D.C.
86
CHAPTER 2
Reconstitution of the E. coli Very Short Patch Repair
System with Purified Proteins
87
- Chapter2 2.1. ABSTRACT
The E. coli Very Short Patch Repair (VSPR) system repairs T:G mismatches that
arise from the deamination of 5-methylcytosines (5mC) created by Dcm. For reasons yet
to be understood, the Dcm protein methylates the second cytosine of 5'- CCAGG -3'
sequences, and to the detriment of the cell these 5mC residues can spontaneously
deaminate and give rise to T:G mismatches. To avoid the accumulation of mutations and
to maintain the sequence integrity of Dcm sites, VSPR removes the mismatched T and
restores the native DNA sequence. Repair is initiated by the Vsr endonuclease, which
produces a single strand nick 5' to the mismatched T. Studies done on cells lacking the
genes for either Pol I or DNA Ligase show diminished levels of VSPR and therefore
these proteins have been implicated in the repair process.
However, the steps
downstream of incision have not been studied biochemically. Herein, we report a novel
in vitro assay for monitoring the entire VSPR pathway. Using this assay, we demonstrate
repair of T:G mismatches contained within short oligodeoxynucleotides by the
combination of Vsr, Pol I, and DNA Ligase proteins. These studies represent the first
reconstitution of the VSPR pathway and lay the foundation for studying the mechanism
of repair more closely.
88
- Chapter2 2.2. INTRODUCTION
In E. coli, the second cytosine of each strand of the palindromic sequence 5'CCWGG -3' (where W is A or T) is methylated at the C5 position of cytosine by the
Dcm methylase to create a fully methylated duplex (Figure 2.1) (1;2). The precise
function of methylation is not understood, but it has been hypothesized to play a role in
defense against restriction modification systems of invading organisms (3).
detriment of the cell, 5-methylcytosine
To the
(5mC) residues created by Dcm can
spontaneously deaminate in a process analogous to that for cytosine (4-6). Deamination
of 5mC in these sequences gives rise to T:G mismatches in hemimethylated Dcm sites
that are known mutagenic hotspots (7;8). If removal of the mismatch does not precede
replication, a C -
T transition mutation, and loss of the Dcm methylation site, will
result. Mutation avoidance and maintenance of these sites is the responsibility of the
very short patch repair (VSPR) pathway (9).
VSPR of mismatched thymine in the sequence 5'- CTAGG -3' (where the T is
mismatched with G) is initiated by Vsr (10). This protein is a strand and sequence
specific endonuclease that creates a single strand nick on the 5' side of the mismatched T
(11). Recombination assays that measure VSPR in vivo have implicated the 5' to 3'
exonuclease activity of Pol I in removal of the mismatched T in addition to a few bases
downstream of the mismatch, and the polymerase activity of Pol I to conduct resynthesis
of the error containing strand (12).
Therefore, the model for VSPR hypothetically
involves nicking by Vsr, nick translation by Pol I, and ligation by the DNA Ligase.
However, no studies to date have explicitly tested the ability of these three enzymes to
carry out VSPR in vitro.
We describe here an assay to monitor the complete VSPR pathway in vitro.
Moreover, we report that the combination of purified Vsr, Pol I, and DNA Ligase protein
from E. coli are necessary and sufficient to repair T:G mismatches to C:G base pairs
contained within a short oligodeoxynucleotide (ODN). Consistent with the literature, we
observed that the nicking efficiency of Vsr is greater on hemimethylated duplexes as
89
- Chapter2 compared to their unmethylated counterparts (Figure 2.1) (13;14). Interestingly, this
preference of Vsr affected both the efficiency of G:T to G:C repair and the extent to
which the repair reactions went to completion. We also observed that the efficiency of
Vsr was increased in the presence of Pol I and DNA Ligase proteins.
Our results
therefore support a model for VSPR involving multiple protein-protein interactions for
maximal efficiency.
90
-Chapter2 2.3. EXPERIMENTAL PROCEDURES
Cloning,Expression, and Purification of Vsr. Preparation of E. coli genomic DNA for
vsr amplification was carried out essentially as described in Current Protocols (15).
Briefly, 1 mL of cells from an overnight culture was spun down and the resulting cell
pellet was resuspended in 570 !xLTE 8.0 (10 mM Tris, 1 mM EDTA). After the addition
of 30 L of 10% SDS and 1.8 units of Proteinase K (Roche), the solution was incubated
at 37 °C for 1 h. This solution was then supplemented
with 100 gL of 5 M NaCl, 80
L
of a 10% CTAC/0.7 M NaCl solution, and incubated at 65 C for 10 minutes. This
solution was extracted twice from 750 ,uLof a 25:24:1 ratio of phenol to chloroform to
isoamyl alcohol (Invitrogen). DNA was precipitated from the final aqueous layer with
420 pLLof Isopropanol, and the DNA pellet was washed with 250 tL of 70% Ethanol.
The resulting DNA was dried in vacuo for 5 minutes, resuspended in 100 1 iL TE, and
quantified using UV spectrometry.
The gene sequence for Vsr was amplified from GM30 genomic DNA by PCR
using the primers ODN 1 and ODN 2 (Table 2.1). PCR was carried out according to the
Opti-prime PCR Optimization Kit (Stratagene). The combination of Taq polymerase
(NEB), buffering conditions #6 or #7, and 2% DMSO as a reaction adjunct provided the
best results. The thermocycling parameters were as follows: 94 °C for 120 seconds, then
thirty rounds of 95 °C for 30 seconds, 54 °C for 45 seconds, and 72 °C for 60 seconds,
and then a final incubation at 72 °C for ten minutes. The 471 bp product was identified
by separating a fraction of the reaction products on a 1% agarose in TAE gel and purified
using the QIAquick PCR Purification kit (QIAGEN). Using the EcoRI and HindIII
restriction sites contained within the primers, the amplified segment was inserted into
complementary sites of pET28a (Novagen) using T4 DNA ligase (NEB). Following
transformation
into competent E. coli DH5a,
DNA from isolated clones were
characterized by restriction analyses and sequencing.
One successful recombinant
expression vector was then transformed into BL21-Gold(DE3)pLysS cells (Stratagene).
Expression of the expected 18 kDa protein containing Vsr (aa 1 to 156) fused to a N-
91
- Chapter2 terminal 6 x His tag was confirmed by Western blot analysis using anti-His tag antibodies
(Amersham).
To prepare cell pellets containing over-expressed Vsr, an overnight culture, grown
in LB containing 25 !xg/mL Chloroamphenicol (Cam) and 50 gg/mL Kanamycin (Kan),
was diluted (100 gtL to 10mL) in LB and grown (roller drum, 37 C) to mid log phase
(OD60 0-0.65).
Expression of Vsr was induced by adding IPTG (Roche) to a final
concentration of 0.75 mM and the cells were grown, as above, for an additional 3 h.
Cells were then harvested by centrifugation (6000 xg, 15 min) and stored at -20 °C for up
to one week.
When purified Vsr was desired, cell pellets were lysed using BugBuster® protein
extraction reagent (Novagen). Briefly, 500 gL of lysis buffer (containing 400 !ALof wash
buffer (25mM HEPES, 300mM NaCl, 10mM MgC12, 2mM ME, pH 8.0), 50 iL of 10x
BugBuster®, 50
L of SIGMA P8340 protease inhibitors cocktail, and 15 units
benzonase) was used to resuspend the pellet and transfer it to a 1.5 mL tube. Disruption
of the cell membrane proceeded by nutating the mixture for 15 min at RT.
The cell lysate was isolated from insoluble material by centrifugation (13,000 xg
for 15 min, at 4 C) and transferred to a QIAGEN Ni-NTA Spin Column pre-equilibrated
by loading the column with 600 }L wash buffer and centrifuging at 2,000 xg for 2 min at
4 C. The Vsr protein mixture was passed through the column by centrifugation as
above, washed five times with 600 FL wash buffer, and purified Vsr was eluted from the
column by passing two fractions of elution buffer (wash buffer plus 250 mM imidazole,
100 tL each) through the column. The Vsr eluates (-0.15 mg/mL) were combined and
kept on ice until subsequent reactions.
Nicking assays. To prepare substrates for nicking assays, 1 nmole of 38-mer ODN 3 was
5'-end labeled with 16.5 pmole [y-3 2 P]ATP (6000 Ci/mmol)(NEN brand, Perken Elmer)
using 10 units of T4 polynucleotide kinase (T4PNK)(NEB) in the supplier's reaction
buffer. The 20 L reaction proceeded for 30 min at 37 C and was then heated to 65 °C
92
- Chapter2 for 30 min to inactivate the reaction. Fractions (2 RL, 100 pmoles) were annealed in 50
mM NaCi with equimolar amounts of complementary strands ODN 4, ODN 5, and ODN
6 to construct duplexes containing unmethylated T:G substrate, hemimethylated T:G
substrate, and unmethylated T:A control duplex respectively (Figure 2.1).
These
duplexes (1 AM) were stored at -20 °C for up to 2 weeks.
Nicking assays were performed by combining 250 fmoles substrate, 1.25 jiL
salmon sperm competitor DNA (Stratagene), 10 RL Vsr protein (-1.5 jig), and reaction
buffer (25 mM HEPES, 100 mM NaCl, 10mM MgC12, 2mM ME, pH 7.5) to a final
volume of 20
L. Reactions were incubated at 37 C (for 0, 15, 30, 60, 120, and 180
min) and stopped by adding an equal volume of formamide plus 10mM EDTA loading
buffer and putting them at -20 C. DNA fragments were separated via 20% denaturing
PAGE, visualized by PhosphorImagery (Storm 840, Molecular Dynamics), and
quantified using ImageQuant 5.2 software (Molecular Dynamics).
Reconstitution of VSPR with Purified E. coli Proteins.
To prepare substrates for
reconstitution reactions, 500 pmoles of 28-mer ODN 7 was 5' phosphorylated using 6.6
pmoles [y-3 2 P]ATP (6000 Ci/mmol)(NEN brand, Perken Elmer) and 15 units of T4PNK
for 10 min in a total reaction volume of 20 [tL, buffered according to the supplier. After
incubation at 37 C for 30 min, the reaction was supplemented with excess (300 nmoles)
non-radiolabeled ATP, incubated at 37 C for an additional 2.5 h, and then heated to 65
°C for 30 min to inactivate the reaction. The completely 5' phosphorylated 28-mer ODN
7 was then ligated to an equimolar amount of 17-mer ODN 8 (500 pmoles), using the
scaffold ODN 9 (500 pmoles), and 1200 units of T4 DNA Ligase (NEB). The 60
L
reaction was buffered according to the enzyme supplier and proceeded at 16 C over
night. A fraction (6 L, 50 pmoles) of the resulting 45-mer ODN, containing an internal
radiolabel, was then mixed with an equimolar amount of 45-mer ODN 10, and an
equimolar amount of 45-mer ODN 11, in separate tubes. The two DNA mixtures (1000
,tL in Vsr reaction buffer) were heated to 75 C and cooled to room temperature slowly,
allowing the unmethylated T:G duplex and hemimethylated T:G duplex to anneal.
93
-Chapter
2Reconstitution reactions were performed by combining 50 fmoles of substrate, 1
AiLsalmon sperm competitor DNA (Stratagene), 15 ptL Vsr (2.25 ig), 5 units E. coli
DNA Polymerase I (NEB), and 5 units E. coli DNA Ligase (NEB) in reconstitution
buffer (25 mM HEPES, 100 mM NaCl, 1OmM MgCl2, 2mM J3ME, pH 7.5). Each
reaction was also supplemented to contain 2 mM ATP (Roche), 100 jiM P-NAD (free
acid, grade I, Roche), 100 M each dNTP (NEB), 50 jig Creatine Kinase (Roche), and 50
mM Creatine Phosphate (Roche). Reactions (50 tL total) were incubated at 37 °C for 1 h
and were stopped by adding an equal volume of formamide plus 10mM EDTA loading
buffer. In addition to performing VSPR reaction in triplicate for each duplex, parallel
control reactions were performed in the presence of all reactants except Pol I and DNA
Ligase proteins to assess Vsr nicking.
Analysis of Reconstition. After reconstitution reaction, the substrate product mix
was separated from the other reaction components via PAGE (20% in TBE, 7 M Urea).
The 45-mer substrate product mix was identified by its position on the gel and its location
relative to the other bands. The appropriate cuts were made to excise the 45-mer and the
gel slice was put into a 1.5 mL screw top tube, crushed thoroughly with 2 wood
applicators, resuspended in 750 jL ddH2O, vortexed for 20 seconds, and put on a rotator
overnight at room temperature for the DNA to elute from the gel bits.
The next day, the DNA was isolated from the gel bits as follows. The top of the
tube was removed and the center of the bottom was punctured using a 21 guage needle.
This tube was then put on top of a 1.5 mL flip top tube which had its top cut off, its
bottom punctured with a 26 guage needle, and contained 0.2 g of sand. This tube was, in
turn, put on top of a collection tube consisting of a 1.5 ml flip top tube with its top cut
off. This three tiered set up was put into a 15 mL centrifugation tube for stability and
centrifuged at 8500 xg for 1.5 hours.
The DNA eluate was transferred from the
collection tube into a sterile 1.5 mL tube by pipet. The middle tube containing the sand
and the trapped gel bits was disposed of appropriately as a radioactive material.
94
-Chapter2 The resulting DNA solution was concentrated to dryness in vacuo and
resuspended in 40
L ddH 2O. The final volume of each sample was brought to 100 [gL
with ddH20. Desalting was accomplished via two G-25 Sephadex Quick Spin Columns
(Roche). The substrate product mixes were then suitable for downstream enzymatic
steps.
Digests were conducted on the 60 gL scale by adding 5 p.L 10X NEB buffer #2, 1
gL BsmI (10 units), 1 jiL BstYI (10 units), and 3 gL ddH 2 0 to each 50 jl of DNA. The
reactions were allowed to proceed at 60°C for 2 h. Then, 2 FL of phosphatase (Roche, 1
units/L)
was added to the digests and incubated at 370 C for an additional 2 h. The
digest/phosphatase reactions were terminated by heating to 80°C for 15 minutes.
The resulting DNA fragments in each mixture were radiolabeled as follows. Each
60 gtL mixture was supplemented with 2
L of a mix containing 0.5 jiM [y-32 P]-ATP
(NEN brand, Perkin Elmer), 0.75 M ATP (Roche), 2.5 units/pzLT4PNK (NEB), and 1 X
T4PNK buffer (supplied by NEB). The kinase reactions were incubated at 37 °C for 45
minutes to ensure that every DNA fragment 5' end was phosphorylated.
The mixtures containing radiolabeled DNA fragments were separated via PAGE
(20%, 7 M Urea). The target 10-mer ODN was located via PhosphorImagery, excised
from the gel, and isolated from the gel slice as described above. Once completely dried,
the DNA was resuspended in a total of 50 pjL of 20 mM sodium acetate pH 5.3, and
desalted using Micro Bio-Spin Chromatography Columns (BioRad) according to
protocol. Before applying the sample to the columns, however, the resin was equilibrated
with 500 ,uLof 20 mM sodium acetate pH 5.3 two times. The resulting DNA sample was
therefore buffered in sodium acetate and ready for digestion into mononucleotides.
To each DNA sample, 2
L of nuclease P1 (United States Biological,
resuspended with ddH2O to 0.3 units/jL) was added. Digestion was allowed to take
place at 65C for 2 h. Afterwards, the mixtures of mononucleotides were concentrated to
-10
L in vacuo and ready for TLC separation.
95
- Chapter2 -
TLC was conducted using cellulose 20 cm x 20 cm Baker-flex PEI-F sheets (J.T.
Baker, Phillipsburg NJ), which were previously washed in ddH2O and allowed to dry
completely.
Approximately 5
L of mixture was spotted and separation of the
mononucleotides proceeded for 12 h using saturated ammonium phosphate pH 5.8 as the
mobile phase. The TLC plate then dried on the bench for -1 h., was protected with Saran
Wrap, and visualized by PhosphorImagery (Storm 840, Molecular Dynamics).
The
relative amounts of thymine and cytosine mononucleotides were calculated using
ImageQuant 5.2 software (Molecular Dynamics).
To account for any contamination that survived the Vsr protein preparation and
contributed to VSPR, the 45-mer DNA from control reconstitution reactions lacking Pol I
and DNA Ligase was isolated and treated in parallel. Given that these reactions were not
supplemented with Pol I and DNA Ligase, isolated 45-mer DNA was theoretically 100%
substrate and representative of 100% T at the interrogation site. The %C found for these
"control reactions" could therefore account for the total background in the assay and was
subtracted from the sample values.
96
- Chapter2 2.4. RESULTS
Analysis of nicking activity. The ability of Vsr to incise substrates that contained T:G
mismatches in the canonical sequence 5'- CTAGG -3' (where the T is mismatched with
G) was measured using a nicking assay.
Incubation of purified Vsr with duplex
substrates that contained a mismatch afforded incision 5' to the mismatched T and
converted the radiolabeled 38-mer T containing strand into a radiolabeled 17-mer and an
unlabeled 21-mer.
Triplicates of identical reactions were stopped over a range of
incubation times and the substrate to product ratio was determined by separating the
reaction mixture by PAGE and visualizing the distribution of 38 and 17-mers by
PhosphorImagery.
A control duplex, in which the radiolabeled T was correctly paired
with A in duplex DNA, was analyzed in parallel.
In total, three DNA duplexes were analyzed by the nicking assay: an
unmethylated duplex, a hemimethylated duplex, and a control duplex (Figure 2.1). Gel
analysis of the replicates (Figure 2.2) shows that there was a good degree of
reproducibility of nicking extent for each substrate.
fragmentation over time.
The control duplex showed no
Additionally, gel analysis of the control duplex reactions
revealed that the oligo experienced essentially no degradation by exonucleases.
The
observation that the substrates and products are well defined bands reflects the propensity
of phosphorothioate bonds in deterring exonuclease degradation, which manifests in a
laddering effect on the gel.
In the presence of normal phosphodiester bonds,
oligonucleotide substrates were readily degraded, primarily from their 3' ends (data not
shown).
Both the unmethylated substrate and the hemimethylated substrates showed
significant fragmentation of the 38-mer substrate to the 17-mer product over time (Figure
2.2). Quantification of the amount of nicked substrate was carried out using ImageQuant
5.2 software and revealed that Vsr was more active on the hemimethylated substrate as
compared to the unmethylated substrate.
Depending on the time of incubation, the
preference of Vsr for hemimethylated substrate was between 2- and 2.5-fold (Figure 2.3).
97
- Chapter2 -
The ability of Vsr to cleave a hemimethylated substrate more efficiently than an
unmethylated substrate was also observed when the duplexes were incubated with Vsr
under reaction conditions that were supplemented for VSPR reconstitution (Figure 2.4).
Again the replicates for each duplex showed good correlation with each other, and the
preference of Vsr for hemimethylated substrate was approximately 3-fold. After the time
allotted, 19 + 5% of unmethylated substrate was nicked and 63 + 2% of hemimethylated
substrate was nicked (Figure 2.5).
There was no signal from nicked product in the VSPR reconstitution reactions
that were supplemented with Vsr, Pol I, and DNA Ligase (Figure 2.4). These results are
consistent with the notion that Pol I and DNA Ligase were capable of the repair events
downstream of Vsr nicking. According to the model for the mechanism of VSPR, action
by Pol I would lead to the removal and replacement of the radiolabeled T mononucleotide
and a small number of additional nucleotides downstream.
Upon ligation of the
remaining nick, a 45-mer product would be formed that lacked a radiolabel and was the
same size as the 45-mer substrate.
The absence of nicked product and the loss of
radioactivity signal from the 45-mer are both consistent with this model.
The signal loss in the reactions containing the hemimethylated substrate (72 +
3%) was more than those for the unmethylated substrate (55 + 4%) (Figure 2.5), and was
consistent with the notion that nicking and subsequent repair was more efficient on the
hemimethylated duplex. It is interesting to note that the amount of unmethylated and
hemimethylated substrate converted to the nicked species was less in the control
reconstitution reactions lacking Pol I and DNA Ligase (19
5% and 63
compared to the reactions that contained these enzymes (55 + 4% and 72
+
2%) as
3%) (Figure
2.5).
After reconstitution of VSPR, 45-mer DNA was separated from the reaction
mixture by denaturing PAGE, isolated from the gel, and purified (Figure 2.6). The 45mer VSPR substrate/product mixture was digested into fragments that were then
98
- Chapter2 radiolabeled.
The 10-mer DNA fragment that contained the radiolabel at the
interrogation site was separated from the reaction mixture by denaturing PAGE and
isolated from the gel. The target oligonucleotides, containing a radiolabeled T or C at the
interrogation site, were digested to mononucleotides, and separated by TLC.
The
resulting TLC plate was analyzed by PhosphorImagery (Figure 2.7) to quantify
percentage cytosine. Some enzymatic degradation of the oligo mixture was evident as
PhosphorImagery revealed DNA fragmentation. Therefore, alongside the target 10-mer
oligo containing a T or C at the interrogation site, a small amount of undesired 10-mer
was isolated.
The resulting values for %C at the interrogation site were 21, 27, and 23 for the
unmethylated substrate. The values for the hemimethylated substrate were 54, 50, and 4.
This third value was analyzed and rejected using the Q test. The Qcritvalue for three
observations at 90% confidence is 0.9 and the Qcalc for the data point was 0.92.
Therefore, the average and standard deviation of three observations is used for the
unmethylated substrate and two observations for the hemimethylated substrate. The % C
at the interrogation site for the unmethylated and hemimethylated substrate was 24
and 52
i
3
3 respectively.
For the reconstitution reactions containing all the enzymes, the number of repair
events initiated that went to completion can be calculated using the %C at the
interrogation site and the radiosignal from the 45-mer DNA after reaction. In the case of
unmethylated duplex, total 45-mer DNA (radiolabeled and non-radiolabeled) isolated
from the gel contained 0.45
+
0.04 parts radiolabeled T from the unreacted substrate and
some number of additional parts non-radiolabeled C from VSPR. The number of parts of
repaired C was calculated to be 0.14 ± 0.02, as this value over the total (0.45 qt 0.04 +
0.14 + 0.02) is equal to the portion determined by TLC analysis (0.24 t 0.03) Therefore,
14 + 2% of total unmethylated substrate was repaired. This value corresponds to 25
4% of the repair events that were initiated (55
4%)(Figure 2.8). The value for the
hemimethylated duplex repaired was derived, as above, to be 30
99
+
i
4%. This value
- Chapter2 corresponds to 42 + 6% of the repair events that were initiated (72 I 3%)(Figure 2.8).
These data are organized in Table 2.2.
100
- Chapter2 2.5. DISCUSSION
Repair of T:G mismatches that arise in the sequence 5'- CTAGG -3' from the
methylation and deamination of cytosine are repaired in E. coli by the very short patch
repair (VSPR) pathway (16). Inititaion of repair is carried out by the Vsr endonuclease
which nicks the T containing strand 5' to the mismatch.
Studies have previously
established that the activity of Vsr for hemimethylated substrates (14) is approximately
2.5-fold better than that for unmethylated substrates (13). Presence of the C5-methyl
group on cytosine in the hemimethylated substrate has been reported to correlate both a
decrease in the dissociation constant (KD) and an increase in the single turnover rate
constant (kst) (17). Analysis of available crystallographic data for Vsr bound to duplex
DNA (18;19) suggests that the C5-methyl group packs against an aliphatic surface of the
protein (Phe-77, Lys-78, Val-79), and assists binding at the protein DNA interface (20).
Consistent with these observations, our purified Vsr protein showed a nicking
efficiency of approximately 2- to 3-fold higher for the hemimethylated substrate as
compared to the unmethylated one. This preference was observed in both the nicking
assay conditions (Figure 2.3) and in the control reconstitution reactions lacking Pol I and
DNA Ligase protein (Figure 2.5). Gel separation of the reaction products from the
nicking assays also shows that, in line with previous literature, strand incision by Vsr is
carried out on the T containing strand and is specific to a T:G mismatch (Figure 2.2).
Subsequent to nicking at the mismatch site, the mismatch must be removed and
replaced to complete the repair event. The model for the steps subsequent to Vsr nicking
was elucidated by data gathered from in vivo VSPR assays in cells of a variety of genetic
backgrounds.
Studies done by Dzidic and Radman established that the polA gene
fragments associated with the 5' to 3' exonuclease activity of Pol I and the polymerase
activity of Pol I were both necessary for efficient VSPR (12).
These results were
consistent with the notion that Pol I removes the mismatched T and conducts strand
resynthesis. This same body of work investigated the level of VSPR in strains lacking
xth, nth, and nfo genes and revealed that these nucleases did not affect VSPR. Lieb
101
- Chapter2 established that strains which lack the functions of the MutS and MutL recognition
proteins show reduced, but not eliminated, levels of VSPR (21). Therefore, these
proteins are thought to assist Vsr in mismatch recognition but are not necessary for the
endonuclease activity. In line with this observation, the decrease in VSPR in mutS or
mutL- strains can be counteracted by complementing the cell with additional Vsr from a
plasmid (10).
Additional evidence that Vsr does not require the MutS and MutL proteins has
come from biochemical experiments. While it has been hypothesized that MutL assists
Vsr by orienting its N-terminus to a conformation that stabilizes DNA binding (22),
standard investigation of the incision event by Vsr is conducted without any additional
enzymes (23). We therefore felt comfortable in the ability of Vsr to initiate VSPR, and
did not include the MutS and MutL proteins in the reconstitution assay mixtures.
Reconstitution reactions in the presence of Vsr, Pol I, and DNA Ligase created
products that were consistent with reconstitution of the VSPR pathway (Figure 2.4). The
presence of one radioactive band, as opposed to the two bands seen when Pol I and DNA
Ligase were not included, suggested that the product of Vsr incision was processed
further by the Pol I and DNA Ligase enzymes. Removal of the mismatched T by Pol I
releases the mismatched mononucleotide and thereby removes the radiosignal from the
nicked product.
Subsequent to removing the mismatched T, Pol I carries out strand
resynthesis and replaces a short tract of the DNA typical for the enzyme (24). The
resulting non-radiolabeled product co-migrated with the substrate in the gel, facilitating
the isolation of the substrate/product mixture to be analyzed for the distribution of T and
C at the interrogation site.
The method by which the distribution of T and C at the interrogation site was
analyzed is outlined in Figure 2.6. The assay involves cleavage of the duplexes into
smaller fragments using restriction enzymes.
interrogation site for 5' radiolabeling.
This step also serves to expose the
The DNA fragments were all of different size
allowing the target oligo containing the radiolabeled interrogation site to be separated and
102
-Chapter2 identified by PAGE. After isolation, the target oligodeoxynucleotide mixture containing
a radiolabeled T or C was digested to mononucleotides and separated by TLC.
Phoshporimager analysis was then used to quantify the %C at the interrogation site. This
parameter was used (as described in the experimental procedures section) in conjunction
with the radiosignal from the 45-mer after the reaction to calculate the extent of repair.
Both substrates were indeed repaired by the combination of Vsr, Pol I, and DNA
Ligase proteins, though with different efficiencies and extent of completion (Table 2.2).
The hemimethylated substrate was repaired (30
4%) about twice as well as the
unmethylated substrate (14 ± 2%). In light that the hemimethylated substrate was also
cleaved more efficiently, it appears as though the nicking step was at least partially rate
limiting in the reaction conditions used. Interestingly, the percent completion was also
higher for the hemimethylated duplex (42 + 6%) as compared to the unmethylated duplex
(25
4%).
These results may indicate that increased affinity of Vsr for the
hemimethylated substrate results in protection of the incised DNA until Pol I and DNA
Ligase can properly process the nick. As a result, a greater percentage of the events
initiated by Vsr on the hemimethylated substrate are reserved for downstream events.
Additionally, it could be the case that Pol I identifies nicked sites more efficiency by the
presence of Vsr protein bound to the DNA.
Vsr is known to form a stable complex with its DNA product (25). Indeed, Vsr
bound to its product has been crystallized (18;19) and observed by band-shift
experiments (25). It is therefore very likely that Vsr stays bound to and protects its
incised DNA product in vivo until Pol I arrives. In turn, Pol I may protect its product
until DNA Ligase can seal the juxtaposing DNA ends. Accordingly, the efficiency of
VSPR would depend on the availability of all proteins involved, and would benefit from
a coordinated process. Figure 2.8 shows that only a fraction of VSPR events initiated
went to completion. Therefore, it is a possibility that additional components are critical
for maximal VSPR efficiency. Studies to investigate this question are underway.
103
-Chapter2 It is also noteworthy that the amount of each duplex incised by Vsr was less in the
control reconstitution reactions as compared to the reactions containing Pol I and DNA
Ligase protein (Figure 2.5). Perhaps, these proteins play a role in functionally assisting
the activity of Vsr and increase its incision efficiency.
There is a growing body of
evidence that supports the notion of multi-component repair complexes. Association of
the MutS, Pol I, and DNA Ligase proteins with the P-clamp has recently been reported
(26). These findings suggest that certain types of repair may have mismatch recognition
and repair that is coordinated. Slight variations of the assay described here may be able
to address these possibilities in the context of VSPR.
104
-Chapter2 Figure 2.1. Unmethylated and hemimethylated Vsr substrates
CH 3
5' - CCAGG -3'
Dcm
3'- GGTCC -5'
I
5'- CCAGG -3'
3'- GGTCC -5'
I
CH 3
Spontaneous
Deamination
of 5mC
5'- CTAGG -3'
3'- GGTCC -5'
5'- CTAGG -3'
3'- GGTCC -5'
I
CH 3
Unmethylated
Vsr Substrate
Hemimethylated
Vsr Substrate
105
- Chapter2 Figure 2.2. PhosphorImager results from Vsr nicking assays on unmethylated,
hemimethylated, and control substrates
Replicate
Replicate 2
1
Replicate 3
Unmethylated T/G Substrate
*5'-
(N)
CTAGG(N)
-3'
3'- (N) GGTCC(N) -5'
'Time
-- --Time
--
--ime
Hemimethylated T/G Substrate
*5' - (N) CTAGG(N) 17-3'
3'-(N) 6GGTCC(N) -5'
I
CH
-ime
wti-ue
Time
>
Time
- >
W
Time
-m
T
eMPm
Time
Control T/A Substrate
*5'- (N) CTAGG(N) -3'
3'-(N)
106
GATCC(N)
17
5'
-Chapter
2Figure 2.3. Graphical representation of unmethylated and hemimethylated substrate
nicking by V7sr
--- unmethylated
110
hemimethylated
100
90
01
e
r
80
70
E
60
50
Uv
.0
Um
o~A
40
30
20
10
0
0
10
20
30
50
40
60
Min at 37 C
% substrate
Unmeth.
Hemimeth.
0 min
100 t 12
100 + 15
60 min
15 min
30 min
85 ± 2
71 + 12
57 + 4
62 + 13
43 + 11
19 + 7
107
- Chapter2 Figure 2.4. PhosphorImagery results from incubating T:G containing duplexes with
VSPR proteins. Reaction products were separated by 20% denaturing PAGE and
visualized by PhosphorImagery. The first three lanes of each gel are replicates of control
reconstitution reactions containing Vsr but lacking Pol I and DNA Ligase. The second
set of reactions are replicates of reconstitution reactions containing Vsr, Pol I, and DNA
Ligase enzymes.
Unmethylated T/G duplex
45 mer
>
28 mer
nicking replicates
VSPR replicates
Hemimethylated T/G duplex
45 mer
28 mer
nicking replicates
108
VSPR replicates
- Chapter 2Figure 2.5. Graphical representation of Vsr nicking results from reconstitution reaction
conditions. Signal loss is represented for control reconstitution reactions lacking Pol I
and DNA Ligase (gray columns) and reconstitution reactions containing those proteins
(black columns).
100
90
72
80
E
e«
LLZ
enD
en
o Q)
:t
3
70
60
55
:t
4
50
-l+-'
ro
Lro+-'
40
C)..Q
30
CJ)CJ)
20
-
C
._
~0
en
::J
10
0
Unmethylated
Substrate
109
Hemimethylated
Substrate
- Chapter 2Figure 2.6. Methodology for assessing VSPR efficiency in vitro
Substrate/Product
Interrogation Site
Mixture
Bsm I
BstY I
5'-TACGGTAGTGGAATG~GGCTTCG~CATTGCTTGAACAT-3'
3'-ATGCCATCACCTTA~GTCCGAAGCTCTAqpTAACGAACTTGTA-5'
x -
T, C
1
dgest
2. Pt"nsJtlatase
3. Racfda~ ( * ) 5' erds
1. Ssml, ~Y1
*S'-TACGGTAGTGGAATGCC-3' *S'-XAGGCTTCGA-3' *S'-GATCCATTGCTTGAACAT-3'
3'-ATGCCATCACCTTAC-S'* 3'-GGGTCCGAAGCTCTAG-S'* 3'-GTAACGAACTTGTA-S'
I
1. 10 mers isolated
and digested
2. mononucleotides
separated by TLC
C
)
T
t
f
•t t t
*5'-pTAGGCTTCGA-3'
Standards
+
Rxn 1 Rxn 2
*5'-pCAGGCTTCGA-3'
% C
110
=
100
*
C I (C + T)
- Chapter2 Figure 2.7.
Representative PhosphorImagery results from TLC separation of
mononucleotides at interrogation site. Separation of cytosine (C) and thymine (T) was
accomplished with TLC and visualized by PhosphorImagery. Shown are representative
results from C and T standards and VSPR reactions on (1) unmethylated substrate, and
(2) hemimethylated substrate.
it
C
T
a
A&
w
W
_
STANDARDS
1
111
2
- Chapter 2Figure 2.8. Percentage of VSPR events initiated that went to completion
100
_~~
_
Repair Initiated
Repair Completed
90
72 :f: 3
80
JY
m
L..
1;)
70
60
..c
50
CJ)
40
::J
~0
55 :f: 4
30
20
10
0
Unmethylated
Substrate
Hemimethylated
Substrate
112
- Chapter2 Table 2.1. Oligodeoxynucleotides (ODNs) used for cloning Vsr into pET28a and
construction of nicking assay and reconstitution assay substrates.
ODN
SEQUENCE
1
5'-TT
2
5'-GAC AAG CTT TCA AGC GAG TAA ATG AAT CCC-3'
3
5'-C*A*C*GGTAGATGGA*C*G*CCTAGGTCTTCAGTATGCTT*G*A*T-3
4
5'-A*T*C*A*A*GCATACTGAAGACCTGGGCGTCCATCTA*C*C*G*T*G-3'
5
5'-A*T*C*A*A*GCATACTGAAGAC(5mC)TGGGCGTCCATCTA*C*C*G*T*G-3'
6
5'-A*T*C*A*A*GCATACTGAAGACCTAGGCGTCCATCTA*C*C*G*T*G-3'
7
5'-TAGGCTTCGAGATCCATTGCTTG*A*A*C*A*T-3'
8
5' -T*A*C*G*G*TAGTGGAATGCC-3'
9
5'-GAAGCCTAGGCATTCC-3'
10
5'-A*T*G*T*T*CAAGCAATGGATCTCGAAGCCTGGGCATTCCACTA*C*C*G*T*A-3'
11
5'-A*T*G*T*T*CAAGCAATGGATCTCGAAGC(5mC)TGGGCATTCCACTA*C*C*G*T*A-3'
5mC
- 5-methylcytosine
GAA TTC ATG ATG GCC GAC GTT CAC GAT-3'
- phosphorothioate linkage
113
-Chapter2 Table 2.2. Quantitative results from reconstitution assays containing Vsr, Pol I, and
DNA Ligase protein
Unmethylated
Substrate
% SubstrateRemaining
45
Hemimethylated
Substrate
4
28 ± 3
55 + 4
72 ± 3
%C at interrogation site
24 ± 3
52 ± 3
% of startingmaterial
14 + 2
30 ± 4
25 ± 4
42
i
(T at interrogation site)
% SubstrateLoss
(Repair Initiated)
repaired
% of repair initiatedthat
went to completion
114
i
6
- Chapter2 2.6. REFERENCES
(1) May, M. S., and Hattaman, S.
(1975) Deoxyribonucleic acid-cytosine
methylation by host- and plasmid-controlled enzymes. J. Bacteriol. 122(1), 129138.
(2) Marinus, M. G. (1984) Methylation of prokaryotic DNA. In DNA Methylation
(Razin, A., Cedar, H., and Riggs, A. D., Eds.) pp 81-109, Springer-Verlag, New
York.
(3) Takahashi, N., Naito, Y., Handa, N., and Kobayashi, I. (2002) A DNA
methyltransferase can protect the genome from postdisturbance attack by a
restriction-modification gene complex. J. Bacteriol. 184(22), 6100-6108.
(4) Lindahl, T., and Nyberg, B. (1974) Heat-induced deamination of cytosine
residues in deoxyribonucleic acid. Biochemistry. 13(16), 3405-3410.
(5) Ehrlich, M., Norris, K. F., Wang, R. Y., Kuo, K. C., and Gehrke, C. W. (1986)
DNA cytosine methylation and heat-induced deamination. Biosci. Rep. 6(4), 387393.
(6) Shapiro, R., and Klein, R. S. (1966) The deamination of cytidine and cytosine by
acidic buffer solutions. Mutagenic implications. Biochemistry. 5(7), 2358-2362.
(7) Coulondre, C., Miller, J. H., Farabaugh, P. J., and Gilbert, W. (1978) Molecular
basis of base substitution hotspots in Escherichia coli. Nature. 274(5673), 775780.
(8) Duncan, B. K., and Miller, J. H. (1980) Mutagenic deamination of cytosine
residues in DNA. Nature. 287(5782), 560-561.
(9) Lieb, M. (1983) Specific mismatch correction in bacteriophage lambda crosses
by very short patch repair. Mol. Gen. Genet. 191(1), 118-125.
(10) Sohail, A., Lieb, M., Dar, M., and Bhagwat, A. S. (1990) A gene required for
very short patch repair in Escherichia coli is adjacent to the DNA cytosine
methylase gene. J. Bacteriol. 172(8), 4214-4221.
(11) Hennecke, F., Kolmar, H., Brundl, K., and Fritz, H. J. (1991) The vsr gene
product of E. coli K-12 is a strand- and sequence-specific DNA mismatch
endonuclease. Nature. 353(6346), 776-778.
(12) Dzidic, S., and Radman, M.
(1989) Genetic requirements for hyperrecombination by very short patch mismatch repair: involvement of Escherichia
coli DNA polymerase I. Mol. Gen. Genet. 217(2-3), 254-256.
115
-Chapter2 (13)
Monastiriakos,
S. K., Doiron, K. M., Siponen, M. I., and Cupples, C. G. (2004)
Functional interactions between the MutL and Vsr proteins of Escherichia coli are
dependent on the N-terminus of Vsr. DNA Repair (Amst). 3(6), 639-647.
(14) Elliott, S. L., Brazier, J., Cosstick, R., and Connolly, B. A. (2005) Mechanism of
the Escherichia coli DNA T:G-mismatch endonuclease (Vsr protein) probed with
thiophosphate-containing oligodeoxynucleotides. J. Mol. Biol. 353(3), 692-703.
(15) Current Protocols in Molecular Biology(2001) John Wiley & Sons, Inc.,
Hoboken, N.J.
(16) Lieb, M., and Bhagwat, A. S. (1996) Very short patch repair: reducing the cost of
cytosine methylation. Mol. Microbiol. 20(3), 467-473.
(17) Turner, D. P., and Connolly, B. A. (2000) Interaction of the E. coli DNA G:Tmismatch endonuclease (vsr protein) with oligonucleotides containing its target
sequence. J. Mol. Biol. 304(5), 765-778.
(18) Tsutakawa, S. E., Jingami, H., and Morikawa, K. (1999) Recognition of a TG
mismatch: the crystal structure of very short patch repair endonuclease in
complex with a DNA duplex. Cell. 99(6), 615-623.
(19)
Bunting, K. A., Roe, S. M., Headley, A., Brown, T., Savva, R., and Pearl, L. H.
(2003) Crystal structure of the Escherichia coli dcm very-short-patch DNA repair
endonuclease bound to its reaction product-site in a DNA superhelix. Nucleic
Acids Res. 31(6), 1633-1639.
(20) Tsutakawa, S. E., and Morikawa, K. (2001) The structural basis of damaged
DNA recognition and endonucleolytic cleavage for very short patch repair
endonuclease. Nucleic Acids Res. 29(18), 3775-3783.
(21) Bell, D. C., and Cupples, C. G. (2001) Very-short-patch repair in Escherichia coli
requires the dam adenine methylase. J. Bacteriol. 183(12), 3631-3635.
(22) Mansour, C. A., Doiron, K. M., and Cupples, C. G. (2001) Characterization of
functional interactions among the Escherichia coli mismatch repair proteins using
a bacterial two-hybrid assay. Mutat. Res. 485(4), 331-338.
(23) Cupples, C. G., and Macintyre, G. (2000) Methods in Molecular Biology. In
DNA Repair PRotocols: Prokaryotic systems (Vaughan, P., Ed.) pp 65-75,
Humana Press Inc., Totwa, N.J.
(24) Bambara, R. A., Uyemura, D., and Choi, T. (1978) On the processive mechanism
of Escherichia coli DNA polymerase I. Quantitative assessment of processivity. J.
Biol. Chem. 253(2), 413-423.
116
- Chapter2 (25) Drotschmann, K., Aronshtam, A., Fritz, H. J., and Marinus, M. G. (1998) The
Escherichia coli MutL protein stimulates binding of Vsr and MutS to
heteroduplex DNA. Nucleic Acids Res. 26(4), 948-953.
(26) Lopez de Saro, F. J., and O'Donnell, M. (2001) Interaction of the beta sliding
clamp with MutS, ligase, and DNA polymerase I. Proc. Natl. Acad. Sci. U. S. A.
98(15), 8376-8380.
117
CHAPTER
3
The Size of Repair Tracts Created by E. coli Very Short
Patch Repair System
118
- Chapter3 3.1 ABSTRACT
The E. coli Very Short Patch Repair (VSPR) pathway is responsible for repairing
T:G mismatches that arise through the Dcm-mediated methylation and subsequent
deamination of the underlined cytosine residue in the sequence 5'-CCAGG -3'. While
the purpose of Dcm-mediated methylation is not well understood, the importance of Dcm
site maintenance is evident, based on the overlap between the dcm and vsr genes. The
Vsr protein initiates repair of T:G mismatches that arise in Dcm sites by producing a
single strand nick 5' to the mismatched T. Pol I then removes and replaces a short tract
of nucleotides downstream of the incision and DNA Ligase seals the nick to complete the
repair event. The small size of repair tracts excised is reflected in the repair system's
name. Bacteriophage crosses revealed that a significant number of VSPR events do not
include co-repair of markers as close as five bases away from the T:G mismatch.
However, closer examination of the repair patch size and distribution was convoluted by
monitoring repair of T:G in close proximity to another marker mismatch. To circumvent
this issue, we have devised a novel in vitro assay to measure the distribution of VSPR
patch sizes in whole cell extracts. The assay involves monitoring the loss of radiosignal
from a series of substrates that contain the signal at prescribed distances downstream of
the T:G mismatch.
We found that the repair patch is distributed about 2 to 4
deoxynucleotides in length. Interestingly, under certain reaction conditions, the addition
of DNA Ligase actually improved the efficiency of repair initiation, suggesting that
VSPR may operate best in the context of a multi-protein complex.
119
- Chapter3 3.2 INTRODUCTION
In E. coli, the second cytosine of 5'- CCWGG -3' sequences (where W is A or T)
is subject to methylation at the C5 position of the pyrimidine ring by the Dcm methylase
(1;2). These 5-methylcytosine (5mC) residues are known to deaminate spontaneously,
giving rise to T:G mismatches (3;4). If correction of the mismatch does not precede
replication, a C
-*
T mutation will result and the Dcm site will be lost. Therefore, a
system must exist to avoid mutations and maintain these sites. Maintenance of the 5mC
sites is the responsibility of the Very Short Patch Repair (VSPR) system (5).
The VSPR system in E. coli is so named because of the short repair tracts it
creates (6). Studies done in the early 1980s by Lieb estimated that the repair tract size
rarely exceeds 10 deoxynucleotides in vivo (7). VSPR of the mismatched thymine is
initiated by Vsr. This protein is a strand and sequence specific endonuclease that creates
a single strand nick 5' to the mismatched T (8). The activity of Vsr is increased by
association
with
the mismatch recognition
proteins MutS
and MutL (9-11).
Recombination assays used to measure VSPR in vivo have implicated the 5' to 3'
exonuclease activity of Pol I in removal of the mismatched T, and the polymerase activity
of Pol I in resynthesis of the error containing strand, because cells that are deficient in
either fragment of the polA gene are deficient in VSPR (12). DNA Ligase is reasonably
assumed to seal the nick and complete the repair event. While the Vsr protein has been
studied biochemically (9;13-16), in vitro investigation of the VSPR events downstream
of incision is lacking (17).
An advantage to studying systems biochemically is that the concentrations of
substrate(s) and protein(s) can be varied incrementally. The ability to easily modulate
individual components facilitates observing their relative contributions to the overall
event. Additionally, these types of studies can yield insight in to how the proteins
cooperate. As a result, biochemical studies can provide a more complete understanding
of the system.
120
- Chapter3 Herein, we describe studies investigating the VSPR system in whole cell extracts
bearing over-expressed Vsr. We have developed a novel assay for monitoring the repair
patch size and distribution created by VSPR, as well as the effect of DNA Ligase on the
repair event. The assay involves a series of oligodeoxynucleotides (ODNs) containing
radioactive signals in the DNA backbone at prescribed distances downstream of the T:G
mismatch. VSPR is monitored through the detection of radiosignal loss from substrate
DNA as the
32 P-group
is removed as part of the repair patch. Our results indicate that
VSPR removes and replaces 2 to 4 deoxynucleotides and may act as a multi-protein
complex for optimal efficiency.
121
- Chapter3 3.3 EXPERIMENTAL PROCEDURES
Cloning and Expression of Vsr. Preparation of E. coli genomic DNA for vsr
amplification was carried out essentially as described in Current Protocols (18). Briefly,
1 mL of GM30 cells from an overnight culture was spun down and the resulting cell
pellet was resuspended in 570 gL TE 8.0 (10 mM Tris, 1 mM EDTA). After the addition
of 30 gL of 10% SDS and 1.8 units of Proteinase K (Roche), the solution was incubated
at 37 °C for 1 h. This solution was then supplemented with 100 jtL of 5 M NaCl, 80 jiL
of a 10% CTAC/0.7 M NaCl solution, and incubated at 65 C for 10 minutes. This
solution
was
extracted
twice
from
750
jtL
of
a
25:24:1
ratio
of
phenol:chloroform:isoamyl alcohol (Invitrogen). DNA was precipitated from the final
aqueous layer with 420 ,tL of isopropanol, and the DNA pellet was washed with 250 jtL
of 70% Ethanol. The resulting DNA was dried in vacuo for 5 minutes, resuspended in
100 AtLTE, and quantified using UV spectrometry.
The gene sequence for Vsr was amplified from GM30 genomic DNA by PCR
using the primers 5'- TTT GAA TTC ATG ATG GCC GAC GTT CAC GAT -3' and 5'GAC AAG CTT TCA AGC GAG TAA ATG AAT CCC -3'.
PCR was carried out
according to the Opti-prime PCR Optimization Kit (Stratagene). The combination of Taq
polymerase (NEB), buffering conditions #6 or #7, and 2% DMSO as a reaction adjunct
provided the best results. The thermocycling parameters were as follows: 94 °C for 120
seconds, then thirty rounds of 95 °C for 30 seconds, 54 °C for 45 seconds, and 72 °C for
60 seconds, and then a final incubation at 72 °C for ten minutes. The -500 bp product
was identified by separating a fraction of the reaction products on a 1% agarose in TAE
gel and purified using the QIAquick PCR Purification kit (QIAGEN). Using the EcoR I
and Hind III restriction sites contained within the primers, the amplified segment was
inserted into complementary sites of pET28a (Novagen) using T4 DNA ligase (NEB).
Following transformation into competent E. coli DHSac, DNA from isolated clones was
characterized by restriction analysis and sequencing.
One successful recombinant
expression vector was then transformed into BL21-Gold(DE3)pLysS cells (Stratagene).
Expression of the expected 18 kDa protein containing Vsr (aa 1 to 156) fused to a N-
122
- Chapter3 terminal 6 x His tag was confirmed by Western blot analysis using an anti His tag
antibody (Amersham).
To prepare cell pellets containing over-expressed Vsr, an overnight culture grown
in LB containing 25 g/mL Cam and 50 ttg/mL Kan, was diluted (100 /IL to lOmL) in
LB and grown on roller drum at 37 °C to mid log phase (OD600-0.65). Expression of Vsr
was induced by adding IPTG (Roche) to a final concentration of 0.75 mM following
which the culture was grown, as above, for an additional 3 h. Cells were then harvested
by centrifugation (6000 xg, 15 min, 4 °C) and stored at -20 °C for up to one week.
When protein extract bearing over-expressed Vsr was desired, cell pellets were
lysed using BugBuster® protein extraction reagent (Novagen). Briefly, 300-500 p.L of
lysis buffer (containing, by volume, 8/10ths reaction buffer (25mM HEPES, 100mM
NaCl, 10mM MgCl 2, 2mM
ME, pH 8.0), 1/10th 10x BugBuster®, and 1/10th SIGMA
P8340 protease inhibitors cocktail) were used to resuspend the cell pellet and transfer it to
a 1.5 mL tube. Disruption of the cell membrane proceeded by nutating the mixture for 15
min at RT. If the solution remained extremely viscous, 0.5 units of benzonase (Novagen)
was added and the tube was nutated for an additional 10-20 minutes. Protein extract was
isolated from insoluble material by centrifugation (13,000 xg, 15 min, 4 °C), transferred
to a sterile tube, and kept on ice until use. Depending on the volume of lysis buffer used,
the total protein concentration ranged from 3.3 to 5.6 mg/mL.
Radioactivity
Incorporation Assay
To prepare
substrate for the radioactivity
incorporation assay, the ODN 5' - C*A*C *G*G*T AGA TGG ACG CCT AGG TCT
TCA GTA TGC T*T*G *A*A*ddC - 3' (where * denotes a phosphorothioate linkage and
ddC represents dideoxy-cytosine) was annealed to 5' - T*T*C *A*A*G CAT ACT GAA
GAC CTG GGC GTC CAT CTA C*C*G *T*G*ddC - 3' in a mixture (250 !xL)
containing 5 nmoles of each oligo, 10 mM Tris, and 20 mM NaCI. The resulting duplex
(20 RIM)was stored at -20 °C.
123
- Chapter3 Radioactivity incorporation assays were performed by combining 20 pmoles
duplex substrate and 70 g protein extract in reconstitution buffer (25 mM HEPES, 100
mM NaCl, 30mM MgC12, 2mM
ME, pH 7.5). Each reaction (Vt = 50
supplemented to contain 2 mM ATP (Roche), 100
L) was also
M f3-NAD (free acid, grade I,
Roche), 50 jig Creatine Kinase (Roche), 50 mM Creatine Phosphate (Roche), 1 nmole of
each dNTP, and 3.3 pmole of [a32P]-dCTP (6000 Ci/mmol, NEN brand, Perkin Elmer).
Reactions were incubated at 37 °C (for 0, 15, 30, 60, and 120 min), stopped by adding an
equal volume of formamide plus 10 mM EDTA loading buffer, and stored at -20 °C until
electrophoretic separation. Reaction mixtures were separated via PAGE (20% in TBE, 7
M Urea), and the radioactive signals were visualized with PhosphorImagery (Storm 840,
Molecular Dynamics) and quantified using ImageQuant 5.2 software (Molecular
Dynamics).
Radioactivity Depletion Assays
To prepare substrates for the radioactivity depletion
assays, 500 pmoles of ODNs 2, 4, 6, 8, 10 and 12 (Table 3.1) were each incubated at 37
°C with kinase (T4 PNK, 10 units, NEB) and 16.5 pmoles of [y3 2 P]-ATP(6000 Ci/mmol,
NEN brand, Perkin Elmer) for 30 min under conditions optimal for the enzyme in a
volume of 20 jiL. After supplementation with of 200 nmoles of cold ATP (Roche), the
reactions proceeded for an additional 60 min. The resulting mixtures, containing fully 5'
phosphorylated strands, were then heat inactivated by incubation at 65 °C for 25 min and
added to mixtures containing 500 pmoles of ODNs 1, 3, 5, 7, 9 and 11 respectively, 500
pmoles of scaffold 5'- GAA GAC CTA GGC GTC C -3', and 3 gL of 10X Ligase buffer
(NEB). Scaffolding ODN 5'- GAA GAC CTA GGC GTC C -3' was utilized for ODNs
1, 3, 5, 7, and 9, while ODN 5'- ATA CTG AAG ACC TAG G -3' was utilized for ODN
10. The resulting solutions (50 tL) were heated to 80 C and cooled to RT slowly to
allow the DNA fragments to anneal. A 5 IL aliquot of each sample (50 pmoles) was
removed to analyze accurately the radioactivity signal from a known amount of DNA.
To ligate the strands held together by the scaffold, each tube was supplemented with
1200 units of T4 DNA Ligase (NEB) and incubated overnight at 16 °C. PAGE analysis
(20% in TBE, 7 M Urea) of a fraction of the pre- and post-ligation mixtures was
124
- Chapter3 conducted to verify the presence of DNA fragments of the correct size. The radiolabeled
38-mers (Int, 2nt, 3nt, 4nt, 5nt, and 10nt Table 3.2) were purified from all reactants by
sequentially centrifuging the ligation mixture through two ProbeQuant G-50 Micro
Columns (Amersham) according to the supplied protocol. A fraction of each purified 38mer was separated by PAGE (20% in TBE, 7 M Urea) next to a fraction of the preligation control. The radioactivity signals were visualized with PhosphorImagery (Storm
840, Molecular Dynamics) and quantified using ImageQuant 5.2 software (Molecular
Dynamics). The stock concentration of each 38-mer was calculated by comparing the
signal intensity of gel-separated product to that for the control aliquot of known
concentration.
Substrate mixtures were made by annealing, as described above, 50
pmoles of radiolabeled DNA to 50 pmoles of the ODN 5'- T*T*C *A*A*G CAT ACT
GAA GAC CTG GGC GTC CAT CTA *C*C*G *T*G -3' in a total of 500 FL 50 mM
NaCl. Additionally, a control duplex was prepared by annealing 50 pmoles of lnt with
50 pmoles of 5' - T*T*C *A*A*G CAT ACT GAA GAC CTA GGC GTC CAT CTA
*C*C*G *T*G - 3', as above, to produce a radiolabeled duplex containing a perfectly
matched T:A pair at the mismatch site.
Radioactivity depletion assays were typically performed by combining 100
fmoles duplex substrate and -55
g protein extract in reconstitution buffer (25 mM
HEPES, 100 mM NaCl, 30mM MgC12, 2mM pME, pH 7.5). Each reaction (Vt = 50 LL)
was also supplemented to contain 2 mM ATP (Roche), 100 gM f3-NAD (free acid, grade
I, Roche), 50 jug Creatine Kinase (Roche), 50 mM Creatine Phosphate (Roche), and 500
pmoles of each dNTP. Reactions were incubated at 37 C for 120 min, stopped by adding
an equal volume of formamide plus 10 mM EDTA loading buffer, and stored at -20 °C
until electrophoretic separation.
Zero time points were analyzed for each substrate in
each reaction condition to verify integrity of substrate and normalize for error. Reaction
mixtures were separated via PAGE (20% in TBE, 7 M Urea), and the radioactive signals
were visualized with PhosphorImagery (Storm 840, Molecular Dynamics) and quantified
using ImageQuant 5.2 software (Molecular Dynamics).
125
- Chapter3 In the reactions where the effect of DNA Ligase was studied, serial dilutions of E.
coli DNA Ligase (10 units/gL, NEB) were made using 1X buffer, and 5-10 gL of the
total 50 p.L reaction volume were added to the reactions just before incubation. Under
normal conditions, each reaction contained 100 fmoles of substrate and 35 }xgprotein
extract. For assay conditions with excess substrate, 1000 fmoles of duplex and 45 }g
protein extract was used in each reaction. All other buffering conditions, supplements,
and handling procedures were the same as above.
Following quantification of radioactive signals, the intensity for each substrate at t
= 0 was designated as 100% and used as a reference for the percent loss of signal from
that substrate after reaction. Additionally, as a redundant control, the relative intensities
of the zero time points were compared to those for the control aliquots to account for
minute deviations from equal amounts of substrates in every reaction. Contribution of
exonuclease degradation from the DNA ends was defined by signal loss from the control
duplex containing no mismatch and subtracted from the respective data values. These
values were then normalized so that the relative signal loss from position 1 was 100.
126
- Chapter3 3.4 RESULTS
The ability of whole cell extracts bearing over-expressed Vsr to carry our VSPR
was first monitored using a simple assay involving incorporation of radiolabeled [a 32P]dCTP into substrate DNA (Figure 3.1). It was hypothesized that if VSPR could execute
repair of T:G mismatches in short duplexes, the repair event would incorporate dCTP
opposite G and provide a radiosignal by which to measure the extent to which VSPR
occurred over time. Gel separation of the reaction products confirmed this hypothesis
and also fortuitously revealed the presence of a small amount of VSPR reaction
intermediate (Figure 3.2). The incorporation of [a32P]-dCTP increased over time in two
species that detail the abundance of resynthesis product pre- (repair intermediate) and
post- (repair product) ligation (Figure 3.3).
The level of the signal in the repair
intermediate remains relatively constant while the signal for the repair product increases
steadily for the first 30 minutes and then levels off after approximately 1 h.
The
incorporation of [a32P]-dCTP into substrate DNA was dependent on Vsr activity and too
low to be detected in wild type strains (data not shown).
Upon the observation that whole cell extracts bearing over-expressed Vsr can
carry out VSPR on T:G mismatches, we modified our assay to address the relative
frequencies of VSPR patch sizes created in whole cell extracts. In this second assay, we
started with mismatch containing duplexes that possessed a radiolabel in the phosphate
backbone at a prescribed distance downstream from the mismatch. In these reactions, the
ability to monitor VSPR depended on detecting radiosignal loss from substrate as the 3 2 p_
group was removed as part of the repair patch (Figure 3.4). Signal loss from the duplexes
containing the signal adjacent to the mismatched T was representative of the total of
repair on all substrates as replacement of mismatched T is a qualifier for repair. The
relative signal loss for this position was therefore normalized to 100. For the substrates
containing a radio signal downstream of the mismatch, the degree of signal loss from
those substrates related the frequency that the resynthesis patch continued through the
position of that signal. In quantifying the signal losses from a series of substrates
127
-Chapter3 containing radiosignals at prescribed distances downstream from the mismatch, we could
therefore address the frequency of specific repair patch sizes.
The relative signal losses for position one, two, and three are very high and
roughly equivalent (Figure 3.5). In contrast, the signal loss from position 4, 5, and 10 are
significantly lower than that for the first three, indicating that a smaller fraction of VSPR
events monitored included repair tracts of these sizes. For eight individual reaction sets
analyzed (data not shown), the signal loss from position 10 was 98
7% of that for
position 5.
To investigate the effect of surplus DNA Ligase on the relative signal losses and
address the step at which exonucleases access the incised DNA, we conducted reactions
containing zero, 0.005 units, and 0.5 units of DNA Ligase. Signal losses from all the
reactions showed a trend similar to what we observed previously (Figure 3.6). The
relative losses for positions 1, 2, and 3 were similarly high and the signal losses for
position 4 and 5 were significantly less. The relative signal loss values for positions 4
and 5 ranged from low fifties to low sixties, and showed no general trend that depended
on the concentration of DNA Ligase. These data suggested that the observed noise did
not result from a discontinuity in the VSPR pathway in between nick translation by Pol I
and Ligation. Rather, the noise in these values was likely due to exonuclease removal of
the radiosignal before nick translation could occur.
To further investigate the effect of DNA Ligase, we conducted assays in which an
overwhelming amount of substrate was used. We hypothesized that, in the absence of
sufficient DNA Ligase, we would observe equal loss of signal from all positions because
DNA ends would persist and exonuclease activity would remove the radiosignal
independent of how far downstream it was located. Our data are consistent with this
hypothesis. The relative signal loss in the reactions containing no supplemental DNA
Ligase are similar and do not follow a trend of decreasing values (Figure 3.7, white
columns). The absence of decreased signal losses at positions 4 and 5 are consistent with
128
- Chapter3 the notion that exonuclease activity is degrading the DNA from the nick site and
removing the radiolabel.
Additional roles for DNA Ligase are suggested by the data from the reactions
containing excess substrate and supplemented protein (Figure 3.7). Not only do the
reactions supplemented with additional enzyme show the characteristic decrease in
relative signal loss around position 3, but they reveal much greater signal losses at
positions 1, 2, and 3 as compared to the reactions containing no DNA Ligase. This
observation was surprising because the protein responsible for terminating the repair
event had a positive influence on the number of events initiated.
The relative signal losses for the reaction containing excess DNA Ligase showed
a gradual decrease as the distance from the mismatch to the radiolabel increased (Figure
3.7, black columns). These results showed a slightly broader distribution of relative
signal losses than previously observed, and indicate that repair tract sizes in these
reaction conditions were approximately 2 to 4 deoxynucleotides in length. The relative
signal loss from position 5 was similar to the relative signal losses from this position in
previous reactions. This result is consistent with the notion that the majority of noise in
this assay is due to exonuclease degradation from the Vsr incision site before Pol I and
DNA Ligase can restore the DNA to its native form.
129
- Chapter3 3.5 DISCUSSION
The Very Short Patch Repair (VSPR) pathway was discovered upon the
observation that the recombination frequencies of certain amber mutations in the (cl)
gene of bacteriophage k were higher than predicted by the physical distance separating
them (19). Further study of the am6 amber mutation revealed that these events arose
from C
T transition mutations in a 5'- CAG -3' glutamine codon located within the
Dcm recognition sequence 5'- CCAGG -3' (6). The recombinants observed included corepair of markers within 10 nucleotides of the mismatch but not in areas further away (6).
Moreover, recombinants were found to contain the C:G genotype and were not affected
by the adenine methylation activity of Dam (5). These observations distinguished VSPR
from methyl-directed mismatch repair, and spawned many studies to understand its
mechanism.
Lieb and coworkers launched a thorough investigation into the length of the repair
tracts created by VSPR (7). By crossing all available repair-prone amber mutations in
gene clwith
available clmarkers up to 68 bp away, their data afforded a comprehensive
picture of VSPR tract lengths. They observed that the frequency of co-repair for markers
10 base pairs and further away was the same. Therefore, VSPR repair tracts rarely
exceed 10 bases in vivo. Closer examination revealed that co-repair was only observed
for markers downstream of the amber mutation, and markers as close as 5 base pairs
away were not always co-repaired. The very short nature of VSPR repair tracts was
becoming clear. Using this assay to investigate the retention of markers within a few
base pairs of the amber mutation, however, was complicated.
The authors noted that one drawback of measuring the repair tract length using
their recombination assay was that VSPR needed to act on T:G mismatches which were
located close to another mismatch, the marker. It was hypothesized that the base pairing
at one mispair could hinder the base pairing at another, and thereby affect repair of the
T:G mismatch.
Thus, interpreting the frequency to which repair tracts went short
130
- Chapter3 distances was limited. One primary goal of ours was to design an assay free from such
limitations.
Our in vitro assay utilizes a series of duplex substrates that all have the same
sequence and bear a centrally located T:G mismatch contained in the sequence 5'CTAGG -3'.
The substrates differ by the location of a radioactive phosphate in their
backbone that is adjacent to the mismatched thymine (position 1) or another base
downstream of the mismatch (position 2, 3, 4, 5, or 10). Repair of the mismatch proceeds
by the removal of the mismatched T and some number of nucleotides downstream.
Therefore, the number of nucleotides that are removed as part of the repair patch may be
probed by monitoring the signal loss from each of the substrates as VSPR occurs (Figure
3.3).
Before conducting these assays, however, we first verified that extracts bearing
over-expressed Vsr would carry out complete VSPR on short substrate duplexes. We
constructed a reaction using a non-radioactive substrate and extracts supplemented with a
small amount of [a32P]-dCTP (Figure 3.1). In doing so, we were able to monitor the
repair event by restoration of a correct C:G base pairing in the DNA. Results in Figure
3.2 show that we were able to resolve the Pol I resynthesis product pre- (repair
intermediate) and post- (repair product) ligation.
The relative abundance (Figure 3.3) of the two species revealed that ligation of
the resynthesis product was partially rate limiting.
The ability to see the repair
intermediate before ligation revealed that the polymerase was providing substrate for the
DNA Ligase faster than the DNA Ligase could act on that substrate. Therefore, the
repair intermediate was kinetically competent and was observed as part of the reaction
mixture.
The relatively small increase in the signal for the repair intermediate, as
compared to that for the repair product, demonstrated that while the ligation step was
partially rate limiting, the rate was not dramatically slower than those of the polymerase
activities. Therefore, our reaction conditions were competent for VSPR.
131
- Chapter3 We next conducted our assay on duplexes containing the radiosignal at positions
adjacent to and downstream of the T:G mismatch. Our in vitro results (Figure 3.5) are in
good agreement with what has previously been estimated in vivo (7), and provide
additional detail as to the distribution of VSPR patch lengths. The frequencies of repair
tracts 1, 2, and 3 bases in length were similarly high. The frequencies of repair tracts 4,
5, and 10 bases in length were also similar to each other but lower than the first three
positions. Therefore, the distribution of VSPR tracts we observed was quite narrow and
consistent with a length of 3 deoxynucleotides.
Our duplexes experienced a significant amount of nonspecific radiosignal loss.
VSPR repair tracts rarely exceed 10 nucleotides in length (7). Therefore, the radiosignal
loss we observed from this position was a result of interfering activity. We believe the
signal losses for positions 4, 5, and 10 are predominantly from nonspecific degradation of
nicked DNA by nucleases.
Each strand of our duplexes contained phosphorothioate
bonds to deter exonucleolytic degradation from the ends.
Additionally, loss of
radiosignal from exonuclease degradation through the substrate ends was accounted for
using a control duplex that did not contain a mismatch. We therefore surmised that the
loss of radiosignal was due to 5' -, 3' exonuclease degradation from the DNA ends
produced by Vsr incision.
We rationalized that the degradation could occur (1) after Vsr incision but before
Pol I nick translation, or (2) after Pol I nick translation but before Ligation of the
juxtaposed DNA ends. If the latter case was the primary contribution to noise in the
assay, it could be overcome by supplementing the reactions with purified E. coli DNA
Ligase.
Addition of surplus DNA Ligase had no significant effect of reducing the
relative signal loss at positions 4 and 5 (Figure 3.6). Therefore, exonuclease degradation
of duplex DNA takes place just after incision by Vsr.
Given this observation, the
substrate duplexes we used may not have been optimal.
Vsr binds and cleaves DNA duplexes composed of 5'- CTAGG -3' and 5'C5mCTGG -3' (where 5mC is 5-methylcytosine) strands more efficiently than those
132
- Chapter3 composed of 5'- CTAGG -3' and 5'-CCTGG -3' strands (15). This preference has also
been rationalized using structural data (20,21), and is justified as the hemimethylated
substrate is the form encountered in vivo (1).
Our duplex substrates were not
hemimethylated and therefore could have experienced relatively destabilized binding to
Vsr. Perhaps, the use of hemimethylated substrates in our assay would increase Vsr
binding and decrease exposure of DNA ends susceptible to degradation.
Conducting a repair reaction under standard conditions plus supplemental DNA
Ligase also allowed us to study the effects of DNA Ligase on VSPR. We rationalized
that if the size of the repair tract created by Pol I was affected by the concentration of
DNA Ligase present, we would be able to observe that affect with our assay. We first
constructed assay conditions that allowed us to observe the contribution of surplus DNA
Ligase to reactions (Figure 3.6). The size and distribution of the repair patch was not
affected by the addition of DNA Ligase, providing good evidence that surplus DNA
Ligase does not influence Pol I to creating short repair tracts.
To test the contribution of DNA Ligase to reactions that did not have sufficient
DNA Ligase activity to keep up with substrate repair, we used reaction conditions which
contained excess substrate and a range of DNA Ligase supplementation.
In these
reactions (Figure 3.7), the added protein did benefit the repair and assist in VSPR
completion.
Without the addition of DNA Ligase the relative signal loss from the
substrates was not typical for a 3 nucleotide repair tract. Instead, the signal losses were
relatively equivalent and did not decrease significantly between positions 1 and 5. In
contrast to reactions containing no supplement, those containing additional DNA Ligase
did show signal losses characteristic for repair tracts of 2 to 4 nucleotides in length.
These results suggested that DNA Ligase does not have a significant effect on the length
of VSPR tract lengths.
One of the most interesting observations was that the relative level of VSPR was
greater in reactions containing sufficient amounts of DNA Ligase (Figure 3.7, black
columns).
We did not expect that the concentration of DNA Ligase, responsible for
133
- Chapter3 terminating the repair event, would have an influence on the efficiency of repair
initiation. This finding suggests that VSPR might be coordinated so that the proteins are
more efficient at starting repair events that can be successfully completed because
sufficient downstream repair pathway proteins are present.
VSPR is possibly most
efficient when accomplished by multiprotein repair complexes.
It has been observed that the MutS, DNA Ligase, and Pol I proteins all interact
with the
sliding clamp protein (22). This protein was originally studied for its role in
tethering DNA polymerase III holoenzyme to DNA by forming a ring around the DNA
and thereby assuring replication processivity.
More recently, however, its ability to
coordinate multiple proteins on DNA has received attention in the context of DNA repair.
One hypothesis (22) posits that the clamps loaded on the lagging strand during DNA
synthesis can assist MutS damage recognition by localizing MutS on newly synthesized
DNA. If the presence of DNA Ligase was somehow beneficial for this interaction, the
result may be more efficient damage recognition and repair initiation.
substrates used in this study are likely too small for
Though the
-clamp sliding to affect the
efficiency of repair initiation, the possibility remains that the P-clamp can organize a
complete assembly of machinery for optimal repair.
134
- Chapter 3Figure 3.1. Method for detecting VSPR by monitoring the incorporation of radioactivity
into substrate DNA
5/-CACGGTAGATGGACGCCTAGGTCTTCAGTATGCTTGAA-3'
3/-GTGCCATCTACCTGCGGGTCCAGAAGTCATACGAACTT-5'
MutS recognizes
and binds to
the mismatch
Vsr creates an incision
5' to the mismatched T
5'-CACGGTAGATGG~AGGTCTTCAGTATGCTTGAA-3'
3'-GTGCCATCTACC~TCCAGAAGTCATACGAACTT-5'
ATP
ADP + Pi
MutSI Mutl complex
presents the
mismatch in a loop
-dGTP
dATP
dTTP
DNA polymerase I
removes and replaces
a short patch of DNA
DNA ligase
seals the nick
5/-CACGGTAGATGGACGCOiAGGTCTTCAGTATGCTTGAA-3'
3/-GTGCCATCTACCTGCGGGTCCAGAAGTCATACGAACTT-5'
135
- Chapter3 Figure 3.2. Extracts from cells over-expressing recombinant Vsr incorporate [a32P]dCTP into short duplex substrates via VSPR
1
2
3 4
5
VSPR
Product
Repair
Intermediate
4'. :-
. Ill
Time in minutes
1.
0
2.
3.
15
30
4.
60
5. 120
20% denaturing PAGE
136
- Chapter3 Figure 3.3. Graphical representation of results from measuring VSPR via radioactivity
incorporation
10) 9) 801)-
/
z 70.
60)-
oO
c-
Repair Intermediate
V
-/-
VSPR Product
40-
'
302() -
V
~-
V
y-V
10 11) O0
'
l/,',,....
I
20
0
I
40
I
I
80
60
I
I
100
120
I
Incubation Time (min)
Intermediate
Product
0 min
1
0
15 min
9
51
137
30 min
60 min
120 min
10
74
16
97
18
100
- Chapter 3Figure 3.4. Method for detecting VSPR by measuring
signal from substrate DNA
the depletion of radioactivity
5'-CACGGTAGATGGACGC~GGTCTTCAGTATGCTTGAA-3'
3'-GTGCCATCTACCTGCGGGTCCAGAAGTCATACGAACTT-5'
MutSrecognizes
and binds to
the mismatch
Vsr creates an
incision 5' to the
mismatched T
5'-CACGGTAGATGG~
GGTCTTCAGTATGCTTGAA-3'
3'-GTGCCATCTACC~CCAGAAGTCATACGAACTT-5'
AlP
ADP + Pi
MutSI Mutl complex
presents the
mismatch in a loop
dNTPS
DNA polymerase I
emoves and replaces
a short patch of DNA
DNA ligase
seals the nick
5'-CACGGTAGATGGACGCCCAGGTCTTCAGTATGCTTGAA-3'
3'-GTGCCATCTACCTGCGGGTCCAGAAGTCATACGAACTT-5'
138
Chapter3 Figure 3.5. Distribution of VSPR patch lengths created in vitro
110100-
100
-[
4
Cla , P
4
91
7
9080-
62
70.0
._>
10
64
+6
± 11
63
1
I
50-
0o rc
I
50-
A. CD
40I1
2
4
3
5
-. II
II
# of Oligodeoxynucleotides Downstream of Incision
139
-l
10
- Chapter 3-
Figure 3.6. Addition of surplus DNA Ligase to VSPR reactions
150
I
_
_
140
130
I No Ligase added
0.005 units Ligase
0.5 units Ligase
120
If)
If)
a 110
--l
co
c: 100
U) 90
0')
(f2.
Q.)
80
>
"Z:;
co
70
Q.)
a::
60
50
40
1
2
3
# of Oligodeoxynucleotides
Position
Position
Position
Position
Position
1
2
3
4
5
No Ligase added
100
106
99
55
57
4
Downstream of Incision
0.005 units Ligase
107
109
100
61
60
140
5
0.5 units Ligase
110
112
91
52
63
- Chapter 3Figure 3.7. Addition of DNA Ligase to VSPR reactions containing excess substrate
150
I
140
I No
_
_
130
Ligase added
0.2 units Ligase
20 units Ligase
120
(j)
(j)
0
-.J
110
ctl
c 100
.2'
(/)
?f2.
Q)
>
'zj
ctl
90
80
70
Q)
~
60
50
40
2
1
3
# of Oligodeoxynucleotides
No Ligase added
Position
Position
Position
Position
Position
1
2
3
4
5
4
Downstream of Incision
0.2 units Ligase
108
111
100
89
73
100
111
89
86
78
141
5
20 units Ligase
132
130
111
81
46
- Chapter3 Table 3.1. Oligodeoxynucleotides (ODNs) used to construct duplexes containing
internal radiolabels at prescribed distances away from the mismatched T
1
2
5'- C*A*C*G*G*TAGATGGACGCC
-3'
5'- TAGGTCTTCAGTATGC*T*T*G*A*A -3'
4
3
5'- C*A*C*G*G*TAGATGGACGCCT -3'
5'- AGGTCTTCAGTATGC*T*T*G*A*A -3'
6
5
5'- C*A*C*G*G*TAGATGGACGCCTA
-3'
5' - GGTCTTCAGTATGC*T*T*G*A*A
-3'
7
5'- C*A*C*G*G*TAGATGGACGCCTAG -3'
8
5'- GTCTTCAGTATGC*T*T*G*A*A -3'
9
10
5' - C*A*C*G*G*TAGATGGACGCCTAGG
-3'
5' - TCTTCAGTATGC*T*T*G*A*A
-3'
11
12
5' - C*A*C*G*G*TAGATGGACGCCTAGGTCTTC - 3'
5' - AGTATGC*T*T*G*A*A- 3'
complement:5'- T*T*C*A*A*GCATACTGAAGACCTGGGCGTCCATCTA**C*G*T*G3'
* = phosphorothioate linkage
142
- Chapter3 T
Table 3.2. DNA strands containing radiolabels at prescribed distances away from the
lnt
;TATGC*T*T*G*A*A- 3'
5'- C*A*C*G*G*TAGATGGACG
2nt
5'- C*A*C*G*G*TAGATGGACG(
2[naD~lIQOiTt[1[1G
rATGC*T*T*G*A*A- 3'
3nt
rATGC*T*T*G*A*A- 3'
5' - C*A*C*G*G*TAGATGGACG(
4nt
5' - C*A*C*G*G*TAGATGGACG(
EUI1el [1l
5'- C*A*C*G*G*TAGATGGACG(
C
5' - C*A*C*G*G*TAGATGGACG(
Cme~Il[1[~
TATGC*T*T*G*A*A- 3'
5nt
alKRllEIrATGC*T*T*G*A*A10 nt
3'
TATGC*T*T*G*A*A- 3'
0 denotes the position of the radiolabel
* = thosphorothioate linkage
,
,
I
.
-
143
- Chapter3 3.6. REFERENCES
(1) May, M. S., and Hattaman, S.
(1975) Deoxyribonucleic acid-cytosine
methylation by host- and plasmid-controlled enzymes. J. Bacteriol. 122(1), 129138.
(2) Marinus, M. G. (1984) Methylation of prokaryotic DNA. In DNA Methylation
(Razin, A., Cedar, H., and Riggs, A. D., Eds.) pp 81-109, Springer-Verlag, New
York.
(3) Coulondre, C., Miller, J. H., Farabaugh, P. J., and Gilbert, W. (1978) Molecular
basis of base substitution hotspots in Escherichia coli. Nature. 274(5673), 775780.
(4) Duncan, B. K., and Miller, J. H. (1980) Mutagenic deamination of cytosine
residues in DNA. Nature. 287(5782), 560-561.
(5) Lieb, M. (1985) Recombination in the lambda repressor gene: evidence that very
short patch (VSP) mismatch correction restores a specific sequence. Mol. Gen.
Genet. 199(3), 465-470.
(6) Lieb, M. (1983) Specific mismatch correction in bacteriophage lambda crosses
by very short patch repair. Mol. Gen. Genet. 191(1), 118-125.
(7) Lieb, M., Allen, E., and Read, D. (1986) Very short patch mismatch repair in
phage lambda: repair sites and length of repair tracts. Genetics. 114(4), 10411060.
(8) Hennecke, F., Kolmar, H., Brundl, K., and Fritz, H. J. (1991) The vsr gene
product of E. coli K-12 is a strand- and sequence-specific DNA mismatch
endonuclease. Nature. 353(6346), 776-778.
(9) Monastiriakos,
S. K., Doiron, K. M., Siponen, M. I., and Cupples, C. G. (2004)
Functional interactions between the MutL and Vsr proteins of Escherichia coli are
dependent on the N-terminus of Vsr. DNA Repair (Amst). 3(6), 639-647.
(10)
Drotschmann,
K., Aronshtam,
A., Fritz, H. J., and Marinus, M. G.
(1998) The
Escherichia coli MutL protein stimulates binding of Vsr and MutS to
heteroduplex DNA. Nucleic Acids Res. 26(4), 948-953.
(11) Lieb, M. (1987) Bacterial genes mutL, mutS, and dcm participate in repair of
mismatches at 5-methylcytosine sites. J. Bacteriol. 169(11), 5241-5246.
(12) Dzidic, S., and Radman, M.
(1989) Genetic requirements for hyperrecombination by very short patch mismatch repair: involvement of Escherichia
coli DNA polymerase I. Mol. Gen. Genet. 217(2-3), 254-256.
144
- Chapter3 (13) Elliott, S. L., Brazier, J., Cosstick, R., and Connolly, B. A. (2005) Mechanism of
the Escherichia coli DNA T:G-mismatch endonuclease (Vsr protein) probed with
thiophosphate-containing oligodeoxynucleotides. J. Mol. Biol. 353(3), 692-703.
(14) Glasner, W., Merkl, R., Schellenberger, V., and Fritz, H. J. (1995) Substrate
preferences of Vsr DNA mismatch endonuclease and their consequences for the
evolution of the Escherichia coli K-12 genome. J. Mol. Biol. 245(1), 1-7.
(15) Turner, D. P., and Connolly, B. A. (2000) Interaction of the E. coli DNA G:Tmismatch endonuclease (vsr protein) with oligonucleotides containing its target
sequence. J. Mol. Biol. 304(5), 765-778.
(16) Gonzalez-Nicieza, R., Turner, D. P., and Connolly, B. A. (2001) DNA binding
and cleavage selectivity of the Escherichia coli DNA G:T-mismatch endonuclease
(vsr protein). J. Mol. Biol. 310(3), 501-508.
(17) Bhagwat, A. S., and Lieb, M. (2002) Cooperation and competition in mismatch
repair: very short-patch repair and methyl-directed mismatch repair in Escherichia
coli. Mol. Microbiol. 44(6), 1421-1428.
(18) Current Protocols in Molecular Biolog2001)
John Wiley & Sons, Inc.,
Hoboken, N.J.
(19) Lieb, M. (1981) A fine structure map of spontaneous and induced mutations in
the lambda repressor gene, including insertions of IS elements. Mol. Gen. Genet.
184(3), 364-371.
(20) Tsutakawa, S. E., Jingami, H., and Morikawa, K. (1999) Recognition of a TG
mismatch: the crystal structure of very short patch repair endonuclease in
complex with a DNA duplex. Cell. 99(6), 615-623.
(21) Bunting, K. A., Roe, S. M., Headley, A., Brown, T., Savva, R., and Pearl, L. H.
(2003) Crystal structure of the Escherichia coli dcm very-short-patch DNA repair
endonuclease bound to its reaction product-site in a DNA superhelix. Nucleic
Acids Res. 31(6), 1633-1639.
(22) Lopez de Saro, F. J., and O'Donnell, M. (2001) Interaction of the beta sliding
clamp with MutS, ligase, and DNA polymerase I. Proc. Natl. Acad. Sci. U. S. A.
98(15), 8376-8380.
145
CHAPTER
4
06-Methylguanine - Formation, Consequences, and Repair
146
- Chapter4 -
4.1. 06-METHYLGUANINE - A BIOLOGICALLY IMPORTANT ADDUCT
In the early 1960s, alkylating agents were thought to cause mutations primarily
through modification of the N7 position of guanine, resulting in atypical base pairing
(luring replication (1,2). Acid hydrolysates of DNA that had been treated with a variety
of agents demonstrated the major product to be 7-alkyl-guanine (3;4). However, the
observation that the ability of an agent to produce N7-guanine adducts did not correlate
with its mutagenicity suggested that some less abundant DNA modification could be
responsible for the mutagenic effects of alkylating agents (3;5).
In 1963, Friedman reported that treating deoxyguanosine with diazomethane
produced an alkylation product,
6-methylguanine (O6mG), that was converted to
guanine upon perchloric acid hydrolysis (6;7).
This finding revealed that certain
alkylation products did not survive typical DNA acid hydrolysis workups and could be
the hidden source of the mutagenic properties of certain agents. The identification of
such products generated within DNA would require modifications to the typical
hydrolysis procedure.
In the late 1960s, Loveless isolated the O6-alkylG product by allowing
deoxyguanosine to react with N-methylnitrosourea (MNU), N-ethylnitrosourea (ENU), or
ethyl methanlesulphonate (EMS) (5).
This lesion, could therefore be produced by
classical alkylating agents and contribute to their mutagenic effects in viv.
The
hydrogen bonding properties of the 06 and N1 atoms in G are altered in 06mG and
change its base pairing properties as a consequence.
In response to these findings,
Loveless commented,
"Therelationshipbetween nucleic acid alkylation and the carcinogenicity
of nitrosaminesand nitroamides compared with "classical" alkylatingagents is
also unsatisfactory.
The extent of nucleic acid alkylation, with which a
correlationwith carcinogenicityhas been vainly sought, has been estimatedafter
acid hydrolysis. It is particularly interesting, in this context, that Kruger et al
147
- Chapter4 found no ethylation of tissueRNAs followingNEU administration. Thissubstance
is a powerful carcinogenand of all the reagents discussedhere affords thelargest
yield of O-alkyl-deoxyguanosine.It seems therefore that it mayyet be possible to
relate carcinogenicitywith nucleic acid reaction but only when the extent of 0alkylationcan be measured "
These observations paved the way for a better understanding of the mechanisms
underlying the mutagenicity and cytotoxicity of alkylating agents. It is now generally
accepted that although 06mG is one of the less abundant lesions created by alkylating
agents, it is a primary adduct contributing to mutagenesis and carcinogenesis (5;8;9).
The following sections detail what has been learned about the lesion and how it is
repaired in cells.
4.2. 06-METHYLGUANINEIS MUTAGENIC
DNA contains many sites susceptible towards alkylation (Figure 4.1), and the
relative proportions of the different alkylation products is dependent on the reaction
mechanism of the agent and the secondary structure of the DNA target. Alkylating
agents can be categorized into two groups that differ with respect to the transition state of
reaction with the DNA bases.
Agents such as methyl methanesulphonate (MMS),
dimethylsulphate (DMS) and methyl iodide (MeI) react through a SN2 (bimolecular)
transition state because the nitrogen groups on DNA are sufficiently nucleophilic for
reaction before the leaving group has left. On the other hand, the activated forms of
agents such as N-methyl-N'-nitro-N-nitrosoguanidine
(MNNG)(Figure 4.2) and MNU
are less sensitive to the groups on DNA with which they react and through a SN1
(unimolecular) transition state alkylate the oxygen atoms in addition to the nitrogen
groups in DNA.
The difference in reactivity between SN1 and SN2 agents is roughly correlated
with the Swain-Scott "selectivity" constant (S) that ranges from 0 to 1 (10). Agents with
148
- Chapter4 low S values tend to react more extensively with the less nucleophilic oxygen centers on
DNA, while agents with high S values react with the more nucleophilic nitrogen centers
(11). For example, the SN1 agent MNU has a S value of 0.42 (12) and produces about
20-fold more O6mG than the SN2 alklyator MMS (13). A summary of the alkylation
product spectra for a typical SN2 agent, MMS, and a typical SN1 agent, MNU, is
presented in Table 4.1.
In duplex DNA, a number of the reactive sites are involved in base pairing and
therefore shielded from alkylation. Accordingly, the most abundant adducts created in
double strand DNA are 7-methylguanine (7mG), 3-methyladenine (3mA), and O6mG. In
single stranded DNA, the 1-methyladenine (mA) and 3-methylcytosine (3mC) lesions
are efficiently produced, as well (14). In addition to the DNA bases, alkylation of the
phosphate backbone can occur to create a mixture of R and S diastreoisomers (Figure
4.3) (15).
In addition to the exogenous agents discussed above, there are a number of
endogenous sources that can damage DNA and cause mutations.
A number of
biomethylation processes utilize S-adenosylmethionine (SAM) as a methyl source, and
this molecule has been observed to weakly methylate DNA in vitro (16). However,
varying the cellular levels of SAM over a 100-fold range had no effect on the
spontaneous mutagenesis in E. coli (17). Bacterially catalyzed nitrosation has been
shown to result in DNA alkylation, likely through the activation of amino acids and
peptides (18;19). Moreover, mutants deficient in nitrite reductase activity was less
susceptible to spontaneous mutagenesis suggesting that nitrosation may be a biologically
relevant source of mutations (18;20;21).
In humans, potent mutagens such as
nitrosamines and related N-nitroso compounds are produced as a result of metabolism
(22).
There are many factors that influence whether a particular DNA alkylation
product will be harmful to the cell. These factors include if the lesion is mutagenic, a
replication block, and/or efficiently repaired. The 7mG lesion is not mutagenic (13) and
149
- Chapter4 does not interfere with transcription (23) or replication (24). However, methylation at
this position increases the rate at which this base depurinates (25). The resulting apurinic
(AP) site can result in a mutation during DNA replication (26). Alternatively, 7mG
residues have been shown to undergo an imidazole ring opening that produces a
formamidopyrimidine residue which is a block to DNA synthesis in vitro (27).
Therefore, one cannot base the biological importance of lesions merely upon their
abundance.
By the 1970s, it was well understood that in addition to reacting with DNA in
vitro the SN1 alkylating agent MNNG was mutagenic in vivo (28-30). Using a forward
mutational assay, Coulondre and Miller studied the base changes necessary to generate
nonsense codons and determined that the predominant class of mutations caused by
MNNG was G:C --- A:T transition mutations (31;32). Years earlier, Loveless had
hypothesized T to pair with O6mG during replication (5), and these results were
consistent with that notion. Subsequently, this hypothesis was directly demonstrated in
vivo using adducts that were site specifically incorporated into DNA (8;33;34). The
preference of O6mG to base pair with T during replication was also confirmed in vitro
using a variety of purified polymerases (35).
Initially, the mutagenicity of O6mGwas thought to arise from the stability of the
resulting 06mG:T base pair, however this idea proved too simplistic as more data on
O6mG pairings became available (36). The stability of duplexes containing O6mG
opposite the four possible bases demonstrated that the O6mG:Cbase pair was actually the
most stable (37). Moreover, 2D-NMR studies (38;39) revealed a O6mG:T pair in
solution with only one strong hydrogen bond between the exocyclic amine group of
O6mGand the 2 of T (Figure 4.4). The 06 methyl group was observed to be syn to the
N1 of O6mG and likely overlapping with the 04 of T sterically. This clashing would
weaken hydrogen bonding between the N1 of O6mG and the N3 of T, providing a weaker
association overall.
In comparison, O6mG:C pairs were observed to pair via two
hydrogen bonds in a wobble arrangement (Figure 4.4). These data suggested that the
preference for O6mG to pair with T is controlled by factors apart from thermodynamics.
150
- Chapter4 -
The original base pairing model had the methyl group of 06mG in the anti
conformation (5). Subsequently, this conformer was crystallized and studied using X-ray
diffraction analysis (40). This Watson-Crick pairing is stabilized by two hydrogen bonds
between 06mG and T (Figure 4.4), and the mutagenic pattern of 06mG suggests that this
anti form of the methoxyl group exists in addition to the syn form in vivo (41).
In vitro replication studies on DNA substrates containing d0mG also suggest that
the lesion can exist in two conformers, only one of which can be bypassed during
replication (42). When 06mG was paired opposite T or C in DNA, about one half of the
substrate could be replicated with the Klenow fragment of DNA Pol I. Substrates where
the 06mG had been dealkylated showed no replication blockage, suggesting that only one
half of 0 6mG is easily bypassed by polymerases.
It has been hypothesized that the 06mG:T pairs with the 06 methyl group anti is
the replicated form on account of its unperturbed geometry (39;43). Swann suggested the
structure of T opposite 06mG, as opposed to C opposite 06mG, provides a more suitable
geometry for forming the phosphodiester bond of T to the growing strand (36).
Additionally, T opposite the lesion may evade exonucleolytic proofreading better than C
(36). This hypothesis was supported by the observations that the rate limiting step for
nucleotide incorporation opposite 06mG by Klenow was the formation of the
phosphodiester bond, and bond formation was faster for T than C (44).
The frequency with which 06mG elicits G:C
-
A:T transition mutations as a
function of the sequence context has been studied.
Dosanjh and coworkers first
conducted in vitro kinetic assays to study the extension of 06mG paired with T or C when
the adduct was flanked by cytosines or thymines (45). They reported that the adduct was
replicated more readily when flanked by cytosines and increased G:C -- A:T transitions
mutations. They also observed that there was a 2-3 fold preference for incorporation of T
opposite the lesion as compared to C, regardless of the context. However, extensive
investigation of the mutation frequency of 06mG in vivo does not support these findings.
151
- Chapter4 -
Delaney and Essigmann (46,47) studied the mutation frequency of
56mGin the
16 nearest neighbor sequence contexts and observed that the lesion pairs with T nearly
100% of the time, independent of the surrounding bases. The latter assay represented the
outcome for replication past O6mG by the DNA polymerase holoenzyme in vivo, as
opposed to the Klenow fragment used in in vitro, and provides the most accurate picture
of O6mGmutagenesis in E. coli.
4.3. O6-METHYLGUANINE
IS CYTOTOXIC
In addition to its mutagenic properties, O6mG is cytotoxic in E. coli and in higher
organisms. Studies done by Karran and Marinus in the early 1980s identified a number
of E. coli mutants that lacked the Dam adenine methylase (dam) and were sensitive to
agents that produced O6mG such as MNNG (48). The Dam protein (see Chapter 1) was
known to methylate d(GATC) sequences and provide a signal by which the parental
DNA strand could be discriminated from the newly synthesized strand for mismatch
repair (MMR) (49-51). Therefore, it was hypothesized that the sensitivity of dam- strains
was dependent on the inability to distinguish the appropriate strand for MMR. Consistent
with this idea, the sensitivity of dam- strains was abolished when the cells additionally
lacked the mismatch recognition protein, MutS (52). It was later shown that O6mG:Tand
C base pairs were recognized by MutS and substrates for MMR (52;53). Furthermore,
subsequent mutagenesis studies revealed that MMR preferentially acted on O6mGpaired
with T rather than C (54), compounding the deleterious effects of O6mG.
There are a number of theories of how MMR introduces sensitivity to O6mG:T
base pairs. One theory posits that replication past the lesion creates O6mG:Tmismatches
that are regenerated when acted on by MMR machinery (55). As a consequence,
reiterative attempts by MMR to repair O6mG:Tpairs could create persistent strand breaks
that may arrest DNA replication (54). It has also been hypothesized that two MMR
events that are close in proximity could produce overlapping repair tracts on opposite
152
- Chapter4 strands and give rise to a toxic double strand break (56). This theory, however, received
criticism on the grounds that mismatches would not occur frequently enough to produce
excision tracts that would overlap (57). Years later, the hypothesis that a single mismatch
could give rise to a double strand break was vindicated when it was observed that in the
absence of d(GATC) methylation, MutH can cleave both DNA strands (58). In higher
organisms, MMR intermediates have been observed to give rise to double strand breaks
that can trigger the apoptotic pathway (59;60) and in some cases block polymerase
replication (61). However, the roles of these events in cellular death are not completely
understood.
Therefore, while the toxicity of agents which produce O6mG has been
established to involve MMR and double strand breaks, the precise mechanism is still
under investigation.
4.4. THE ADAPTIVE RESPONSE TO ALKYLATING AGENTS
It was discovered in 1970 that the O6mGlesions created by MNNG were removed
from DNA (62), however the mechanism by which the damage was removed was not
understood until discovery of the adaptive response.
In the mid 1970s, Cairns and
Samson were attempting to devise an experimental system to segregate mutated from
nonmutated DNA in bacterial cells that were exposed to low doses of alkylating agents
for a short time. While these efforts never came to fruition, they realized that their
system could be used to examine the mutation rates invoked by low levels of these
agents. Subsequent experiments measured the mutation rates affiliated with exposure of
cells to low doses of MNNG, and demonstrated that, under certain conditions, mutations
would accumulate for the first hour but not afterwards (63).
Additional experiments confirmed this phenomenon and revealed that when E.
coli cells were exposed to low levels of MNNG before challenging the cells with a much
higher dose, the cells had a significant resistance to the lethal and mutagenic effects of
the agent (63-65). This resistance was also discovered to be dependent on protein
synthesis, suggesting that this response involves the synthesis of new proteins to protect
153
- Chapter4 the cells from alkylation damage (63). Demonstration that the induction of proteins
relates an increase in removal of O6mG lesions from DNA was consistent with the ideas
that this lesion accounted for most MNNG induced mutagenesis and was targeted by the
adaptive response for increased repair (66). Indeed, quantitative assessment of the
removal of O6mGpre and post induction revealed that the removal activity increased by
approximately 1000-fold (67). This phenomenon is now referred to as the adaptive
response to alkylation damage (68;69).
By this time, measuring DNA damage and repair was readily accomplished by
monitoring the creation and retention of radiolabeled DNA damage.
Typically, cells
were treated with a radiolabeled alkylating agent and the high molecular weight DNA
was isolated and analyzed for radiosignal. Therefore, the observation that there was less
radiosignal from DNA containing O6mG lesions (where the signal was on the methyl
group) from cells that were adapted, as compared to those that were not, was expected
(66). It was also expected that released DNA adduct could be recovered in the acid
soluble fraction containing excised DNA products, however, this was not the case
(67;70). Further investigation revealed that the radioactive methyl groups, released from
DNA, were tightly associated with protein (71) that had methyltransferase (MTase)
activity (72). Subsequently, the Ada protein was isolated and its ability to demethylate
O6mGwas directly demonstrated (73). Additionally, Ada was shown to be the positive
regulator of the adaptive response as cells that lacked functional Ada were significantly
more sensitive to MNNG treatment (74;75).
In the past fifteen years, our understanding of the adaptive response has
developed substantially. In addition to ada, a number of genes involved in the adaptive
response were mapped (76) and identified by their contribution to decreasing the
mutagenecity or cytotoxicity of alkylating agents.
It is now understood that, by a
mechanism described below, the Ada protein regulates increased expression of four
genes, ada, alkB, alkA, and aidB, upon exposure of E. coli to sub-lethal doses of
alkylating agents (Figure 4.5) (77). In total, these genes, which map at 45, 47 and 95 min
154
- Chapter4 on the E. coli genetic map (78-80), relate protection against the mutagenecity of
methylation, ethylation, propylation, and bulkier adducts, as well (79;81).
In efforts to understand how each repair protein contributes to the protective
effects of the adaptive response, mutant strains were studied. The ada- mutants were
found to be sensitive to both the mutagenecity and cytotoxicity of alkylating agents (74)
owing to, as discussed above, the mutagenic and cytotoxic effects of O6mG (5;8;9). The
number of Ada molecules increases from one or two molecules per cell in an uninduced
state (82) to approximately 1000 upon induction (67), in a process that takes about one
hour (83). This Ada protein, and how it regulates the adaptive response, is discussed in
detail below.
The AidB protein is the product of the aidB gene, is so named because of its
alkylation inducibility. The 60 kDa protein shows homology with isovaleryl-coenzyme
A dehydrogenases (84). Indeed, crude extract from cells bearing over-expressed AidB
protein show isovaleryl-coenzymeA dehydrogenase activity (84). Over-expression of the
protein leads to reduced mutagenesis after exposure to alkylating agents, and thus the role
of AidB has been hypothesized to involve repair of unidentified DNA lesions or the
detoxification of the alkylating agents or their reaction intermediates (84-86). Recently,
AidB was reported to use flavin adenine dinucleotide (FAD) and bind to DNA directly
(86). In light of these observations, another hypothesis posits that AidB is a methylated
base dehydrogenase.
Strains deficient in alkA were found to be sensitive to killing, but not mutagenesis
(87). The AlkA protein was discovered to be a glycosylase that has the ability to excise a
wide variety of damaged bases including 3mA, 7mG, 7-methyladenine (7mA), 3methylguanine (3mG), d-methylthymine
(OmT),
and &-methylcytosine (mC)
(88;89). In contrast to the other alkylation lesions, 3mA was found to be biological
significance as its ability to block DNA synthesis produces cytotoxicity
(90;91).
Therefore, upon a 10 to 20-fold induction as part of the adaptive response (77;87), the
155
- Chapter4 primary importance of this glycosylase is to remove the 3mA methylation product. For
this reason, AlkA is oftentimes referred to as 3-methyladenine DNA glycosylase.
Strains that are deficient in alkB were isolated as early as 1983 (80) however the
function of the protein remained unknown for a long time afterwards. While it was
established that the bacterial alkB gene conferred resistance to MMS (SN2) damage both
in bacterial and human cells (92), it was not until repair studies were conducted with
single stranded phage that substrates were proposed.
The observation that AlkB
proficient and deficient strains processed methylated single stranded DNA differently,
suggested that the mA and 3mC lesions were AlkB substrates (93). These lesions are
primarily created in single stranded DNA because the 1 position of A and the 3 position
of C are not exposed in duplex structures (Figure 4.1). The discovery that AlkB shares a
protein fold typical for dioxygenases inspired the idea that the protein could be a aketoglutarate-Fe(II)-dependent dioxygenase (94). This notion was subsequently verified
biochemically, when in the presence of Fe(II) and a-ketoglutarate, AlkB exhibited repair
of
mA and 3mC substrates by an oxidative demethylation mechanism (95-99).
Subsequently, AlkB has been shown to also repair 3-ethyldeoxycytidine (100), 1methyldeoxyguanosine (100), 3-methylthymine (101), I,PA-ethenoadenine (102), and
3,NM-ethenocytosine (102).
4.5. ADA - IN VIVO AND IN VITROPROPERTIES
Ada is the product of the ada gene located at 47 min on the E. coli genetic map
(80;103), and is so named because of its central role in the adaptive response (104).
Upon induction, levels of Ada increase from approximately one molecule per cell (82) to
approximately 1000 molecules per cell (67). It has also been reported that uninduced
cells experience a 20-fold increase in Ada upon entry into stationary phase (105). The
protein
displays MTase
activity
(Figure
methylphosphtriesters (15;88;107), and
4.6) on
O6mG (71), O4mT
(106)
larger alkyl groups, as well (79;108;109).
However, the activity against alkyl groups generally decreases as the size of the groups
156
- Chapter4 increases. For example, the rate of transferase activity for ethyl groups is about one tenth
that for methyl groups in vivo (79). Ada additionally appears to repair o6-chloroethyl
groups from guanine. These groups are formed by agents such as 1,3-bis(2-chloroethyl)1-nitrosourea (BCNU) and N-(2-chloroethyl)-NP-cyclohexyl-N-nitrosourea (CCNU) and
are toxic on account of their ability to form interstrand cross links in DNA (110). The
presence of Ada provides protection against the lethality of these agents, suggesting that
Ada repairs chloroethyl monoadducts before they can cross link (111;112).
Despite the fact that the O6-methyl bond is stable at neutral pH (77) Ada carries
out direct reversal of the methylation in about 1 second at 37 °C (113), or at a rate of k >
10-8 M-1 sec
'
(114).
Additionally, Ada shows no requirement for Mg2+ or other
cofactors (77). Therefore, this protein appears to be especially tuned to overcome
significant energy barriers in its ability to remove alkyl groups from DNA.
Under limiting conditions, Ada was found to be more efficient at repairing O6mG
as compared to O4mT (88). This preference was also observed in in vitro mutagenesis
studies (115), as the mutation frequency for 06mG (0 - 1.7%) was lower than that for
O4 mT (12%). The alkyl groups on the 06 position of G and the O4position of T are both
exposed from the major groove of DNA. However, Ada does not repair any other forms
of alkylation damage that project into the major groove. Additionally, Ada shows no
MTase activity for N methylated residues in DNA (113). Therefore, the specificity of
MTase activity on damaged DNA bases is apparently tuned for alkyl groups that project
into the major groove of DNA and are oxygen-linked (88).
Ada repairs O6mG in single stranded DNA, albeit at a dramatically reduced rate
(113). This finding is especially relevant since significant single stranded DNA regions
exist in replication forks. The inefficiency of OmG repair in these regions may be one
of the factors that contributes to the known property of MNNG to generate mutations in
clusters at replication forks (116-118). Additionally, MTase activity assays on substrates
of varying lengths revealed that the rate of repair increases as the length of the substrate
increases (114). MTase repair of the free O6mG base is 10-7-fold slower than &OmGin
157
- Chapter4 duplex DNA (114). Therefore, the overall structure of DNA must play a role in the
recognition and repair process.
Ada activity was originally purified to physical homogeneity from an E. coli
mutant that constitutively expresses the adaptive response (119;120). The molecular
weight of the Ada protein is 39 kDa (354 aa), however, the protein is readily degraded in
crude cell extracts to an 19 kDa fragment (72;110,'113;121;122) that retains the ability to
repair 06mG (122) and O4mT (106). For this reason, the Ada protein was originally
isolated as the C-terminal domain (120;123), and was not purified as the entire Ada
protein until it was expressed recombinantly (15;124).
Endogenous fragmentation of Ada was shown to occur at two sites adjacent to
Lys-178 and Lys-129 (125). Interestingly, the two sites show considerable sequence
homology to each other (126) and to a sequence in UvrB (126;127). The similarity
between the primary structure of the Ada and UvrB sites suggested that a single protease
could be responsible for the cleavage at their sites.
Subsequently, purified OmpT
protease was observed to cut the purified Ada and UvrB proteins into their expected
fragment sizes (77,128). Ada was cleaved primarily adjacent to Lys-178 (129). A
possible role for this proteolytic cleavage is discussed below.
Treatment of Ada with trypsin, chymotrypsin, subtilisin, and V8 protease
results in cleavage within a stretch of 10 amino acids (aa 171 to 181) near the center of
the protein (125), consistent with the notion that Ada has two functional domains of
similar size connected by a short hinge region. MTase assays demonstrated that the
separate C-terminal and N-terminal fragments of Ada retained distinct transferase
activities. More specifically, the C-terminal fragment of Ada repaired 06mG and O4 mT,
while the N-terminal fragment repaired methylphosphotriesters (125). In accordance
with the presence of two distinct MTase domains, when purified Ada was incubated with
excess 06mG and phosphotriester substrates, both substrates were repaired and the
protein contained approximately two methyl groups per Ada protein (15).
158
- Chapter4 The transfer of an alkyl group from DNA to Ada results in regeneration of the
native DNA structure (73) and a methylated cysteine residue (72). Methylated cysteine
residues are stable and therefore, in light that no detectable Ada demethylase activity has
been detected (77;120), MTase repair results in a dead end protein product (113). This so
called "suicide in activation" has been demonstrated numerous times both in vitro
(72;113;122;123) and in vivo (67).
The cysteine residue that participates in MTase repair of 06mG and O4mT
adducts is Cys-321 (123). The Cys residue is found in the Pro-Cys-His-Arg active site
sequence found in all known 06mG MTases (130). Structural data of the C-terminal
domain of Ada revealed that this residue is surprisingly buried (130). Therefore, it was
hypothesized that must undergo a conformational change to expose the cysteine to act as
a nucleophile (130). An alternative model suggested that the helix-turn-helix (HTH)
motif in Ada binds to the DNA to invoke the alkylated base to flip out of the DNA (131).
More recent structural data, for active and alkylated human Ada protein, implicate a HTH
motif in DNA binding and the use of an "arginine finger" to extrude the alkylated base
from the stacked DNA duplex (132).
The known eukaryotic MTases are homologous to the 19 kDa C-terminal domain
of E. coli Ada (77;133) and have a high degree of structural similarity (134). Therefore,
structure activity relationships for one homolog can assist understanding for the entire
family of proteins. Additionally, a nucleotide flipping mechanism has great precedent
and is observed in a number of DNA MTases and glycosylases (135). The model that
includes repair of an extra-helical base is therefore more mechanistically established.
MTase repair of methylphosphotriesters is carried out in the N-terminal domain of
Ada (Figure 4.6) (125). This chemical group can exist in one of two isomeric forms, R
and S, and the observation that only one half of methylphosphotriesters are repaired
suggested that Ada can only repair one of these forms (15). Quantitative studies
subsequently revealed that only the S form is repaired (15;136). Docking models suggest
that the ability of Ada to repair the S isomer relies on the accessibility of the methyl
159
- Chapter4 group.
The S isomer projects the methyl group into solution whereas the R isomer
projects into the major groove of DNA and is not within reach of the active site of Ada
(Figure 4.3) (137).
Structural data (138) revealed that the N-terminal domain of Ada is composed of
a central B-sheet sandwiched between two ct-helices. A tightly bound Zn2+ (139) is
coordinated by four cysteines (Cys-38, Cys-42, Cys-69, and Cys-72), from two adjacent
loops, in a tetrahedral geometry.
In addition to the role of Zn2+ in activating the
nucleophilicity of the acceptor cysteine (140), the metal acts to coordinate the structure of
Ada (138). Repair of alkyl groups on the DNA backbone by the N-terminus of Ada
converts the protein to a strong transcriptional activator of several genes (77) by eliciting
a protein surface that enhances the affinity of the protein for the promoters of the adaalkB operon and the alkA and aidB genes (Figure 4.5) (137;141-143). Cys-69 was
previously assigned as the residue that accepted the alkyl group from the DNA backbone
(125), however recent X-ray and NMR structural data (144) indicate that Cys-38 is the
methyl acceptor. It has been suggested that methyl transfer to this ligand could cause a
reorganization of the Zn2 + coordination and subsequent conformational changes in the
protein structure (145). However, extended x-ray absorption fine structure and x-ray
absorption near edge structure data indicate that the integrity of the thiolate-zinc center is
preserved after methytransfer (137). Additionally, solution structures revealed that the
transferred methyl group can not make contacts with promoter DNA, suggesting that
methyltransfer to Ada causes conformational changes that creates a high affinity DNA
binding face (137). However, the precise mechanism by which alkylated Ada converts
the protein into a transcriptional activator, however, is unknown (146).
Upon the repair of a methylphosphotriester in DNA by Ada, the N-terminus of the
protein binds to a transcriptional regulatory element called the "Ada box" (77;114). The
minimal element, identified by deletion studies and site directed mutagenesis, consists of
the octanucleotide sequence 5'- AAAGCGCA -3' just upstream of ada-alkB operon
(147). The extended sequence 5'- AAANNAAAGCGCA -3' is found upstream of the
alkA gene. These Ada boxes are just upstream of the putative RNA polymerase binding
160
- Chapter4 site (143;148).
Therefore, it appears as though alkylated Ada serves to initiate the
adaptive response by facilitating the binding of RNA polymerase (77;104;114). The Cterminal domain of Ada has been implicated in the binding of the RNA polymerase
(114).
As a consequence to the inability of MTases to act enzymatically on their
substrates, the mutagenesis of alkylating agents accelerates after the available MTase
molecules have conducted repair and are thus incapacitated.
For this reason it is
important that the adaptive response has a sensitive mechanism for signaling the cell to
cope with excess alkylation damage. The level of methylphosphotriesters produced by
alkylating agents is more than that for 06mG and O4mT (13), and they therefore do serve
as an acute trigger for the adaptive response.
It is not completely understood how the adaptive response switches off upon the
withdraw of alkylation pressure. One simple possibility for returning the concentration
of repair proteins to their steady state levels is through the diluting effects of cell division
(77;114). It has also been hypothesized that proteolysis of Ada could serve to terminate
its ability to bind to promoters. As discussed above, the Ada protein is readily cleaved by
the endogenous OmpT protease in vitro (77;128).
The N-terminal product was
subsequently shown to bind to promoter sites but have no effect on gene expression
(77;114). It follows then that Ada degradation could be responsible for down-regulation
of the adaptive response in vivo (149). Studies conducted in ompT mutant strains,
however, showed normal levels of Ada and kinetics of induction (129). Direct evidence
for OmpT regulation of the adaptive response therefore is still lacking. Lastly, high
concentrations of unmethylated Ada protein have been related to the inhibition of ada
gene activation by the methylated form (150). This observation provides yet another
possibility for how cells return to constitutive levels of repair proteins.
4.6. OGT - IN VIVO AND IN VITRO PROPERTIES
161
- Chapter4 In the late 1980s, the isolation of E. coli mutants that lacked the function of Ada
led to the discovery of a second protein with alkyltransferase activity (149;151). The
cellular level of Ogt was found to be independent of the adaptive response (152), as cells
that lacked ada showed a low threshold for MNNG tolerance that did not increase upon
low dose exposure. Approximately 30 molecules of Ogt are present in the cell at all
times (153), and therefore provides a constant level of protection and can hold off a low
level of damage while the cell adapts for high level repair.
Ogt is the protein product of the ogt gene which maps at 29.4 min on the E. coli
chromosome (154). The protein contains 171 amino acids and a molecular weight of 19
kDa (153). The fact that Ogt is so similar in size and function to the C-terminal domain
of Ada was likely to have been the source of come confusion (82). Therefore, it wasn't
until the use of mutant strains and recombinant expression techniques that trustworthy
studies were possible (104;153;155).
Ogt has been shown to remove methyl groups from O6mG and O4mT, but has no
detectable activity for methylphosphotriesters (149;151;153). Additionally, experiments
conducted in E. coli cell extracts demonstrated that the concentrations of O6mG and
O4 mT substrates necessary to inactivate 50% of the Ada or Ogt methyltransferase activity
were quite different (156). The Ogt protein showed a high affinity for the O6mG
substrate (8.1 nM) and an even higher affinity for the O4mT substrate (3 nM). Ada, on
the other hand showed good affinity for O4 mT (27.5 nM) but very high affinity for O6mG
(1.25 nM).
These preferences may be evidence that Ada and Ogt divide the
responsibility of repairing alkylation damage.
The ogt gene has a nucleotide sequence that is distinct from that of ada, however
the amino acid sequence of Ogt shows extensive regions of homology with the C-teminal
domain of Ada (104;123;153;157). The methyl accepting cysteine in Ogt (Cys-139)
corresponds with Cys-321 in Ada, and is part of a region that is highly conserved in
MTases from a number of species (153). To date, no structural data for Ogt has been
reported.
162
- Chapter4 -
4.7. PROTECTION AGAINST ALKYLATION INDUCED TOXICITY BY ADA AND OGT
The cytotoxic effects of MNNG have been studied in E. coli strains with a variety
of repair proficiencies to define the relative contributions to alkylation protection. In
cells that were not pre-exposed to alkylating agent, strains of (wild type) wt and ogf
backgrounds were similarly resistant whereas the adaexperienced a high level of killing
strain was sensitive and
(152;154). These results suggested that even in
conditions that are not optimal for inducing the adaptive response (66;74) Ada provides
protection from alkylation damage. While both the ada- and the ogt ada- double mutant
were sensitive to MNNG, Ogt over-expression was sufficient to confer resistance in these
strains to wt level (154). These results indicate that O6mG is the major alkylation product
involved in cell death.
The overlapping of O5mG repair by Ada and Ogt was also seen in MTase double
mutants. Cells that lacked Ada and Ogt were more sensitive to alkylation treatment than
cells lacking just Ogt (154). The ada- ogf double mutant was reported to be more
sensitive than the ogt mutant in one case (152) but not another (154). Interestingly, adacells showed a low threshold to mutation induction owing to the presence of Ogt. On the
other hand, mutation induction in the ada- ogf double mutant was graded and linear
(152). Clearly, the two proteins tag-team MTase repair of O6mG to evade the lethal
effects of the lesion.
As described above, toxicity data indicate that O6mG achieves killing via MMR.
Studies done by Karran and Marinus in the 1980s established that cells that are defective
in the Dam methylase are sensitive to agents that produce O6mG such as MNNG (48).
This effect is due to the O6mG lesions which evade MTase repair and are subsequently
recognized by the mismatch repair proteins and inaccurately processed. Accordingly,
while dam- mutants are sensitive to alkylation pressure, dam- ada- double mutants are
163
- Chapter4 more sensitive (52). Additionally, cells that lack Ada in addition to Dam and MutS
experience more lethality from MNNG treatment (52).
In contrast with the view that O-alkyl repair by Ada provides direct protection
against the cytotoxic effects of MNNG, some reports have posited that Ada provides
indirect resistance to alkylation damage by inducing AlkA which then initiates BER on
toxic lesions other than 06mG (77;87;158;159). However, Ada has been shown to have
a direct effect of protecting cells against MNNG induced toxicity.
Comparison of alkA- and alkA- ada- strains showed that the double mutant was
more sensitive to MNNG (152). This result would not be expected if the sole protection
effect of Ada was through the adaptive response. Therefore, while it is possible Ada may
provide some indirect protection to cells by AlkA induction, Ada also protects cells
directly.
4.8. REPAIR OF 06-METHYLGUANINE BY NUCLEOTIDE EXCISION REPAIR
In addition to the ability of MMR and the Ada and Ogt MTases to act on 06mG
and O4 mT, nucleotide excision repair (NER) has also been shown to process these lesions
(83;160-164). Removal of these lesions by NER was originally monitored in a range of
host strains using monoclonal antibodies (83), and results suggested that NER provided a
low level of protection against alkylation induced mutation. It was subsequently shown
that O6mG is repaired by NER much less efficiently than other types of DNA damage
(165), and therefore MTase repair is likely the primary repair mechanism for this lesion.
On the other hand, repair of O4 mT by NER appeared to be quite efficient and actually
blocked by the presence of MTases (164;166). Therefore, repair of the 06mG and O4mT
lesions are independently balanced by a number of distinct repair systems.
164
- Chapter4 Figure 4.1. Sites of methylation on the DNA bases
06 G
P
N 7G
qH2
CN
q
.......
... .
NO
DNA
......
DNA
H2 N
t
A
N3 C
N 3G
T
0c
N 7A
04 T
H2
-....
DNA
DNA
N T
N3 T
t|
N1 A
N 3A
02 T
165
- Chapter4 Figure 4.2. Conversion of MNNG to the active methanediazonium ion by cysteine.
Adapted from reference (146).
HOOC
Cysteine
H 2N
SH
+
NH 2
H3C.N-N
NO 2
MNNG
N-.
I
HOOC
HOOC
NH3
NH 2
H2N
#11'
S--
N
NO 2
S
S NH
N-NO 2
+
H3 C,
N
II
N OH
H
-
H+
-
H3 C-N-N
H20
methanediazonium
166
- Chapter4 Figure 4.3. The R and S sterioisomers of methylphosphotriesters
Position of
r
for R
Position of r
for S i!
167
- Chapter4 Figure 4.4. Base pairing models for 06mG:T and 06mG:C. Shown are the models for
(A) &0mG in the anti conformation bound with T, (B) 06mG in the syn conformation
bound with T, and (C) 06mG in the syn conformation bound with C in a wobble
arrangement.
R
6
FN
rN
N- -
........ F
t N,
Nzz
A
NH2
NN
0
I.........
0-R
x;N<N
N=<
B
N
X/
NH 2
rN
N
-R
H2 N
N=<
C
NH 2
0
168
tN/
- Chapter4 Figure 4.5. Genetic structure and regulation of the adaptive response in E. coli
-aH
Q
r _|f
3
~
~\i,./
/
N terminal
of Ada
unmethylated
Ada with
methylated
N-terminus
scriptiona i Activation
TrankB
-
M
<
ada
.
MTase
alkB
,
.
. .~~~~
alkA
.
"..
I
Oxidative
-
I
Glycosylase
demethylase
aidB
I
I-
I
Dehydrogenase
. . .
169
- Chapter4 Figure 4.6. Direct reversal of base damage by Ada
Ada
S-H
Cys-321
I
Cys-38
4
X
¾\-
H3C\ o
I
Li
'-Z hi
,
-IN
aN
S-methyl
phosphotriester
)YmG
phosphodiester
G
CH -S
S-CH
I
170
3
NH2
- Chapter4 Table 4.1. Relative amounts of DNA alkylated products following reaction of double
stranded DNA with MMS (SN2 ) and MNU (SN1) alkylating agents. Adapted from Singer
and Grunberger (13).
MMS (SN 2
in vitro
)
in vivo
MNU(SN1)
Adenine
N1
N3
N7
3.8
10.4
1.8
8.7
4.2
2.4
9.0
1.7
Guanine
N3
N7
0.6
83.0
1.3
0.8
0.5
83.0
67.0
69.0
cP
0.3
0.3
6.3
5.1
ND
0.08
ND
0.11
0.3
5.1
ND
0.1
in vitro
in vivo
1.3
3.1
Thymine
N3
0.4
Cytosine
N3
<1
Phosphotriester
0.8
<1
171
0.6
0.4
16
9.0
- Chapter4 4.9. REFERENCES
(1) Lawley, P. D., and BROOKES, P.
(1961) Acidic dissociation of 7:9dialkylguanines and its possible relation to mutagenic properties of alkylating
agents. Nature. 192:1081-2., 1081-1082.
(2) NAGATA, C., IMAMURA, A., FUKUI, K., and SAITO, H. (1963) QUANTITY
OF CHARGE TRANSFER AND CARCINOGENIC ACTIVITY OF 4NITROQUINOLINE
1-OXIDES
AND
OF
SOME
AROMATIC
HYDROCARBONS. Gann. 54:401-14., 401-414.
(3) BROOKES, P., and Lawley, P. D. (1963) EFFECTS OF ALKYLATING
AGENTS ON T2 AND T4 BACTERIOPHAGES. Biochem. J. 89:138-44., 138144.
(4) Lawley, P. D. (1966) Effects of some chemical mutagens and carcinogens on
nucleic acids. Prog. Nucleic Acid Res. Mol. Biol. 5:89-131., 89-131.
(5) Loveless, A. (1969) Possible relevance of 0-6 alkylation of deoxyguanosine to
the mutagenicity and carcinogenicity of nitrosamines and nitrosamides. Nature.
223(202), 206-207.
(6) Friedman, O. M., Mahapatra, G. N., and Stevenson, R. (1963) The methylation of
deoxyribonucleosides by diazomethane. Biochim. Biophys. Acta. 68:144-6., 144146.
(7) Friedman, O. M., Mahapatra, G. N., Dash, B., and Stevenson, R. (1965) Studies
on the action of diazomethane on deoxyribonucleic acid. The action of
diazomethane on deoxyribonucleosides. Biochim. Biophys. Acta. 103(2), 286-297.
(8) Loechler, E. L., Green, C. L., and Essigmann, J. M. (1984) In vivo mutagenesis
by 06-methylguanine built into a unique site in a viral genome. Proc. Natl. Acad.
Sci. U. S. A. 81(20), 6271-6275.
(9) Pegg, A. E.
(1990) Mammalian 06-alkylguanine-DNA alkyltransferase:
regulation and importance in response to alkylating carcinogenic and therapeutic
agents. Cancer Res. 50(19), 6119-6129.
(10) Swain, C. G. and Scott, C. B. (1953) Quantitative correlation of relative rates.
Comparison of hydroxide ion with other nucleophilic reagents toward alkyl
halides, esters, epoxides and acyl halides. pp 141-147,
(11) Roberts, J. J. The repair of DNA modified by cytotoxic , mutagenic, and
carcinogenic chemicals. pp 211-435,
(12) Veleminsky, J., Osterman-Golkar, S., and Ehrenberg, L. (1970) Reaction rates
and biological action of N-methyl- and N-ethyl-N-nitrosourea. Mutat. Res. 10(3),
169-174.
172
- Chapter4 (13) Singer, B., and Grunberger D Molecular Biology of Mutagens and Carcinogens:
Reactions of Directly Acting Agents with Nucleic Acids (1983) Plenum Press,
New York.
(14) Bodell, W. J., and Singer, B. (1979) Influence of hydrogen bonding in DNA and
polynucleotides on reaction of nitrogens and oxygens toward ethylnitrosourea.
Biochemistry. 18(13), 2860-2863.
(15) McCarthy, T. V., and Lindahl, T. (1985) Methyl phosphotriesters in alkylated
DNA are repaired by the Ada regulatory protein of E. coli. Nucleic Acids Res.
13(8), 2683-2698.
(16) Rydberg, B., and Lindahl, T. (1982) Nonenzymatic methylation of DNA by the
intracellular methyl group donor S-adenosyl-L-methionine is a potentially
mutagenic reaction. EMBO J. 1(2), 211-216.
(17) Posnick, L. M., and Samson, L. D. (1999) Influence of S-adenosylmethionine
pool size on spontaneous mutation, dam methylation, and cell growth of
Escherichia coli. J. Bacteriol. 181(21), 6756-6762.
(18) Taverna, P., and Sedgwick, B. (1996) Generation of an endogenous DNAmethylating agent by nitrosation in Escherichia coli. J. Bacteriol. 178(17), 51055111.
(19) Garcia-Santos, M. P., Calle, E., and Casado, J. (2001) Amino acid nitrosation
products as alkylating agents. J. Am. Chem. Soc. 123(31), 7506-7510.
(20) Harrison, K. L., Jukes, R., Cooper, D. P., and Shuker, D. E. (1999) Detection of
concomitant formation of 06-carboxymethyl- and 06-methyl-2'-deoxyguanosine
in DNA exposed to nitrosated glycine derivatives using a combined
immunoaffinity/HPLC method. Chem. Res. Toxicol. 12(1), 106-111.
(21) Sedgwick, B.
mutagens
in
Carcinogenesis.
(1997) Nitrosated peptides and polyamines as endogenous
06-alkylguanine-DNA
alkyltransferase
deficient
cells.
18(8), 1561-1567.
(22) Marnett, L. J., and Burcham, P. C. (1993) Endogenous DNA adducts: potential
and paradox. Chem. Res. Toxicol. 6(6), 771-785.
(23) Ludlum, D. B. (1970) The properties of 7-methylguanine-containing templates
for ribonucleic acid polymerase. J. Biol. Chem.(245), 477-482.
(24) Prakash, L., and Strauss, B. (1970) Repair of alkylation damage: stability of
methyl groups in Bacillus subtilis treated with methyl methanesulfonate. J.
Bacteriol. 102(3), 760-766.
173
- Chapter4 (25) Chetsanga, C. J., and Lindahl, T. (1979) Release of 7-methylguanine residues
whose imidazole rings have been opened from damaged DNA by a DNA
glycosylase from Escherichia coli. Nucleic Acids Res. 6(11), 3673-3684.
(26) Michaels, M. L., Pham, L., Cruz, C., and Miller, J. H. (1991) MutM, a protein
that prevents G.C----T.A transversions, is formamidopyrimidine-DNA
glycosylase. Nucleic Acids Res. 19(13), 3629-3632.
(27) O'Connor, T. R., Boiteux, S., and Laval, J. (1988) Ring-opened 7-methylguanine
residues in DNA are a block to in vitro DNA synthesis. Nucleic Acids Res.
16(13), 5879-5894.
(28) Neale, S. (1972) Effect of pH and temperature on nitrosamide-induced mutation
in Escherichia coli. Mutat. Res. 14(2), 155-164.
(29) Neale, S. (1976) Mutagenicity of nitrosamides and nitrosamidines in microorganisms and plants. Mutat. Res. 32(3-4), 229-266.
(30) Jimenez-Sanchez, A., and Cerda-Olmedo, E. (1975) Mutation and DNA
replication in Escherichia coli treated with low concentrations of N-methyl-N'nitro-N-nitrosoguanidine. Mutat. Res. 28(3), 337-345.
(31) Coulondre, C., and Miller, J. H. (1977) Genetic studies of the lac repressor. III.
Additional correlation of mutational sites with specific amino acid residues. J.
Mol. Biol. 117(3), 525-567.
(32) Coulondre, C., and Miller, J. H. (1977) Genetic studies of the lac repressor. IV.
Mutagenic specificity in the lacI gene of Escherichia coli. J. Mol. Biol. 117(3),
577-606.
(33) Hill-Perkins, M., Jones, M. D., and Karran, P. (1986) Site-specific mutagenesis
in vivo by single methylated or deaminated purine bases. Mutat. Res. 162(2), 153163.
(34) Bhanot, O. S., and Ray, A. (1986) The in vivo mutagenic frequency and
specificity of 06-methylguanine in phi X174 replicative form DNA. Proc. Natl.
Acad. Sci. U. S. A. 83(19), 7348-7352.
(35) Singer, B., and Dosanjh, M. K. (1990) Site-directed mutagenesis for quantitation
of base-base interactions at defined sites. Mutat. Res. 233(1-2), 45-51.
(36) Swann, P. F. (1990) Why do 06-alkylguanine and 04-alkylthymine miscode?
The relationship between the structure of DNA containing 06-alkylguanine and
04-alkylthymine and the mutagenic properties of these bases. Mutat. Res. 233(12), 81-94.
174
- Chapter4 (37) Gaffney, B. L., and Jones, R. A. (1989) Thermodynamic comparison of the base
pairs formed by the carcinogenic lesion 06-methylguanine with reference both to
Watson-Crick pairs and to mismatched pairs. Biochemistry. 28(14), 5881-5889.
(38)
Patel, D. J., Shapiro, L., Kozlowski,
S. A., Gaffney, B. L., and Jones, R. A.
(1986) Structural studies of the 06meG.T interaction in the d(C-G-T-G-A-A-T-TC-O6meG-C-G) duplex. Biochemistry. 25(5), 1036-1042.
(39) Patel, D. J., Shapiro, L., Kozlowski, S. A., Gaffney, B. L., and Jones, R. A.
(1986) Covalent carcinogenic 06-methylguanosine lesions in DNA. Structural
studies of the 06 meG X A and 06meG X G interactions in dodecanucleotide
duplexes. J. Mol. Biol. 188(4), 677-692.
(40) Leonard, G. A., Thomson, J., Watson, W. P., and Brown, T. (1990) Highresolution structure of a mutagenic lesion in DNA. Proc. Natl. Acad. Sci. U. S. A.
87(24), 9573-9576.
(41) Seo, K. Y., Jelinsky, S. A., and Loechler, E. L. (2000) Factors that influence the
mutagenic patterns of DNA adducts from chemical carcinogens. Mutat. Res.
463(3), 215-246.
(42) Dosanjh, M. K., Loechler, E. L., and Singer, B. (1993) Evidence from in vitro
replication that 06-methylguanine can adopt multiple conformations. Proc. Natl.
Acad. Sci. U. S. A. 90(9), 3983-3987.
(43) Kalnik, M. W., Li, B. F., Swann, P. F., and Patel, D. J. (1989) 06-ethylguanine
carcinogenic lesions in DNA: an NMR study of 06etG.T pairing in
dodecanucleotide duplexes. Biochemistry. 28(15), 6170-6181.
(44) Tan, H. B., Swann, P. F., and Chance, E. M. (1994) Kinetic analysis of the
coding properties of 06-methylguanine in DNA: the crucial role of the
conformation of the phosphodiester bond. Biochemistry. 33(17), 5335-5346.
(45) Dosanjh, M. K., Galeros, G., Goodman, M. F., and Singer, B. (1991) Kinetics of
extension of 06-methylguanine paired with cytosine or thymine in defined
oligonucleotide sequences. Biochemistry. 30(49), 11595-11599.
(46) Delaney, J. C., and Essigmann, J. M. (1999) Context-dependent mutagenesis by
DNA lesions. Chem. Biol. 6(10), 743-753.
(47) Delaney, J. C., and Essigmann, J. M. (2001) Effect of sequence context on 0(6)methylguanine repair and replication in vivo. Biochemistry. 40(49), 14968-14975.
(48) Karran, P., and Marinus, M. G. (1982) Mismatch correction at 06-methylguanine
residues in E. coli DNA. Nature. 296(5860), 868-869.
175
- Chapter4 (49) Marinus, M. G., and Morris, N. R. (1974) Biological function for 6methyladenine residues in the DNA of Escherichia coli K12. J. Mol. Biol. 85(2),
309-322.
(50) Marinus, M. G., and Morris, N. R. (1975) Pleiotropic effects of a DNA adenine
methylation mutation (dam-3) in Escherichia coli K12. Mutat. Res. 28(1), 15-26.
(51) Marinus, M. G. (1976) Adenine methylation of Okazaki fragments in Escherichia
coli. J. Bacteriol. 128(3), 853-854.
(52) Calmann, M. A., Evans, J. E., and Marinus, M. G. (2005) MutS inhibits RecAmediated strand transfer with methylated DNA substrates. Nucleic Acids Res.
33(11), 3591-3597.
(53) Rasmussen, L. J., and Samson, L. (1996) The Escherichia coli MutS DNA
mismatch binding protein specifically binds 0(6)-methylguanine DNA lesions.
Carcinogenesis. 17(9), 2085-2088.
(54) Pauly, G. T., Hughes, S. H., and Moschel, R. C. (1994) Response of repaircompetent and repair-deficient Escherichia coli to three 06-substituted guanines
and involvement of methyl-directed mismatch repair in the processing of 06methylguanine residues. Biochemistry. 33(31), 9169-9177.
(55) Karran, P., and Bignami, M. (1994) DNA damage tolerance, mismatch repair and
genome instability. Bioessays. 16(11), 833-839.
(56) Glickman, B. W., and Radman, M. (1980) Escherichia coli mutator mutants
deficient in methylation-instructed DNA mismatch correction. Proc. Natl. Acad.
Sci. U. S. A. 77(2), 1063-1067.
(57) Grafstrom, R. H., Amsterdam, A., and Zachariasewycz, K. (1988) In vivo studies
of repair of 2-aminopurine in Escherichia coli. J. Bacteriol. 170(8), 3485-3492.
(58) Au, K. G., Welsh, K., and Modrich, P. (1992) Initiation of methyl-directed
mismatch repair. J. Biol. Chem. 267(17), 12142-12148.
(59)
Meikrantz,
W., Bergom,
M. A., Memisoglu,
A., and Samson, L.
(1998) 06-
alkylguanine DNA lesions trigger apoptosis. Carcinogenesis. 19(2), 369-372.
(60) Kaina, B., Ziouta, A., Ochs, K., and Coquerelle, T. (1997) Chromosomal
instability, reproductive cell death and apoptosis induced by 06-methylguanine in
Mex-, Mex+ and methylation-tolerant mismatch repair compromised cells: facts
and models. Mutat. Res. 381(2), 227-241.
(61)
Voigt, J. M., and Topal, M. D.
(1995) 06-methylguanine-induced
blocks. Carcinogenesis. 16(8), 1775-1782.
176
replication
Chapter4 (62) Lawley, P. D., and Orr, D. J. (1970) Specific excision of methylation products
from DNA of Escherichia coli treated with N-methyl-N'-nitro-Nnitrosoguanidine. Chem. Biol. Interact. 2(2), 154-157.
(63) Samson, L., and Cairns, J. (1977) A new pathway for DNA repair in Escherichia
coli. Nature. 267(5608), 281-283.
(64) Jeggo, P., Defais, T. M., Samson, L., and Schendel, P. (1977) An adaptive
response of E. coli to low levels of alkylating agent: comparison with previously
characterised DNA repair pathways. Mol. Gen. Genet. 157(1), 1-9.
(65) Jeggo, P., Defais, M., Samson, L., and Schendel, P. (1978) An adaptive response
of E. coli to low levels of alkylating agent. In DNA Synthesis, Present and Future
(Molineux, I., and Kohiyama, M., Eds.) pp 1011-1024, Plenum Publishing Corp.,
New York.
(66) Schendel, P. F., and Robins, P. E. (1978) Repair of 06-methylguanine in adapted
Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 75(12), 6017-6020.
(67) Robins, P., and Cairns, J. (1979) Quantitation of the adaptive response to
alkylating agents. Nature. 280(5717), 74-76.
(68) Kleibl, K. (2002) Molecular mechanisms of adaptive response to alkylating
agents in Escherichia coli and some remarks on 0(6)-methylguanine DNAmethyltransferase in other organisms. Mutat. Res. 512(1), 67-84.
(69) Sedgwick, B., and Lindahl, T. (2002) Recent progress on the Ada response for
inducible repair of DNA alkylation damage. Oncogene. 21(58), 8886-8894.
(70)
Schendel, P. F., Edington, B. V., McCarthy, J. G., and Todd, M. L. (1983) Repair
of alkylation damage in E. coli. In Cellular Responses to DNA Damage
(Friedberg, E. C., and Bridges, B. A., Eds.) pp 227-240, Alan R. Liss, Inc., New
York.
(71) Karran, P., Lindahl, T., and Griffin, B. (1979) Adaptive response to alkylating
agents involves alteration in situ of 06-methylguanine residues in DNA. Nature.
280(5717), 76-77.
(72) Olsson, M., and Lindahl, T. (1980) Repair of alkylated DNA in Escherichia coli.
Methyl group transfer from 06-methylguanine to a protein cysteine residue. J.
Biol. Chem. 255(22), 10569-10571.
(73) Foote, R. S., Mitra, S., and Pal, B. C. (1980) Demethylation of 06methylguanine in a synthetic DNA polymer by an inducible activity in
Escherichia coli. Biochem. Biophys. Res. Commun. 97(2), 654-659.
177
- Chapter4 (74) Jeggo, P. (1979) Isolation and characterization of Escherichia coli K-12 mutants
unable to induce the adaptive response to simple alkylating agents. J. Bacteriol.
139(3), 783-791.
(75)
Mitra,
S., Pal, B. C., and Foote,
R. S.
(1982)
06-methylguanine-DNA
methyltransferase in wild-type and ada mutants of Escherichia coli. J. Bacteriol.
152(1), 534-537.
(76) Sedgwick, B., and Robins, P. (1980) Isolation of mutants of Escherichia coli with
increased resistance to alkylating agents: mutants deficient in thiols and mutants
constitutive for the adaptive response. Mol. Gen. Genet. 180(1), 85-90.
(77) Lindahl, T., Sedgwick, B., Sekiguchi, M., and Nakabeppu, Y. (1988) Regulation
and expression of the adaptive response to alkylating agents. Annu. Rev. Biochem.
57:133-57., 133-157.
(78) Volkert, M. R., Nguyen, D. C., and Beard, K. C. (1986) Escherichia coli gene
induction by alkylation treatment. Genetics. 112(1), 11-26.
(79) Sedgwick, B., and Lindahl, T. (1982) A common mechanism for repair of 06methylguanine and 06-ethylguanine in DNA. J. Mol. Biol. 154(1), 169-175.
(80) Kataoka, H., Yamamoto, Y., and Sekiguchi, M. (1983) A new gene (alkB) of
Escherichia coli that controls sensitivity to methyl methane sulfonate. J.
Bacteriol. 153(3), 1301-1307.
(81) Todd, M. L., and Schendel, P. F. (1983) Repair and mutagenesis in Escherichia
coli K-12 after exposure to various alkyl-nitrosoguanidines. J. Bacteriol. 156(1),
6-12.
(82)
Rebeck,
G. W.,
Smith,
C. M.,
Goad,
D. L., and Samson,
L.
(1989)
Characterization of the major DNA repair methyltransferase activity in unadapted
Escherichia coli and identification of a similar activity in Salmonella
typhimurium. J. Bacteriol. 171(9), 4563-4568.
(83) Samson, L., Thomale, J., and Rajewsky, M. F. (1988) Alternative pathways for
the in vivo repair of 06-alkylguanine and 04-alkylthymine in Escherichia coli:
the adaptive response and nucleotide excision repair. EMBO J. 7(7), 2261-2267.
(84) Landini, P., Hajec, L. I., and Volkert, M. R. (1994) Structure and transcriptional
regulation of the Escherichia coli adaptive response gene aidB. J. Bacteriol.
176(21), 6583-6589.
(85) Landini, P., and Volkert, M. R. (1995) Transcriptional activation of the
Escherichia coli adaptive response gene aidB is mediated by binding of
methylated Ada protein. Evidence for a new consensus sequence for Ada-binding
sites. J. Biol. Chem. 270(14), 8285-8289.
178
- Chapter4 (86)
Rohankhedkar,
M. S., Mulrooney,
S. B., Wedemeyer, W. J., and Hausinger, R. P.
(2006) The AidB component of the Escherichia coli adaptive response to
alkylating agents is a flavin-containing, DNA-binding protein. J. Bacteriol.
188(1), 223-230.
(87) Evensen, G., and Seeberg, E. (1982) Adaptation to alkylation resistance involves
the induction of a DNA glycosylase. Nature. 296(5859), 773-775.
(88) McCarthy, T. V., Karran, P., and Lindahl, T. (1984) Inducible repair of Oalkylated DNA pyrimidines in Escherichia coli. EMBO J. 3(3), 545-550.
(89) Hollis, T., Lau, A., and Ellenberger, T. (2001) Crystallizing thoughts about DNA
base excision repair. Prog. Nucleic Acid Res. Mol. Biol. 68:305-14., 305-314.
(90) Yamamoto, Y., Katsuki, M., Sekiguchi, M., and Otsuji, N. (1978) Escherichia
coli gene that controls sensitivity to alkylating agents. J. Bacteriol. 135(1), 144152.
(91) Lindahl, T. (1993) Instability and decay of the primary structure of DNA. Nature.
362(6422), 709-715.
(92) Chen, B. J., Carroll, P., and Samson, L. (1994) The Escherichia coli AlkB protein
protects human cells against alkylation-induced toxicity. J. Bacteriol. 176(20),
6255-6261.
(93) Dinglay, S., Trewick, S. C., Lindahl, T., and Sedgwick, B. (2000) Defective
processing of methylated single-stranded DNA by E. coli AlkB mutants. Genes
Dev. 14(16), 2097-2105.
(94) Aravind, L., and Koonin, E. V. (2001) The DNA-repair protein AlkB, EGL-9,
and leprecan define new families of 2-oxoglutarate- and iron-dependent
dioxygenases. Genome Biol. 2(3), RESEARCH0007.
(95)
Falnes, P. O., Johansen, R. F., and Seeberg, E. (2002) AlkB-mediated
oxidative
demethylation reverses DNA damage in Escherichia coli. Nature. 419(6903), 178182.
(96) Trewick, S. C., Henshaw, T. F., Hausinger, R. P., Lindahl, T., and Sedgwick, B.
(2002) Oxidative demethylation by Escherichia coli AlkB directly reverts DNA
base damage. Nature. 419(6903), 174-178.
(97) Aas, P. A., Otterlei, M., Falnes, P. O., Vagbo, C. B., Skorpen, F., Akbari, M.,
Sundheim, O., Bjoras, M., Slupphaug, G., Seeberg, E., and Krokan, H. E. (2003)
Human and bacterial oxidative demethylases repair alkylation damage in both
RNA and DNA. Nature. 421(6925), 859-863.
179
- Chapter4 (98) Begley, T. J., and Samson, L. D. (2003) AlkB mystery solved: oxidative
demethylation of NI-methyladenine and N3-methylcytosine adducts by a direct
reversal mechanism. Trends Biochem. Sci. 28(1), 2-5.
(99) Jiricny, J. (2002) DNA repair: bioinformatics helps reverse methylation damage.
Curr. Biol. 12(24), R846-R848.
(100) Delaney, J. C., and Essigmann, J. M. (2004) Mutagenesis, genotoxicity, and
repair of 1-methyladenine, 3-alkylcytosines, 1-methylguanine, and 3methylthymine in alkB Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 101(39),
14051-14056.
(101) Koivisto, P., Robins, P., Lindahl, T., and Sedgwick, B. (2004) Demethylation of
3-methylthymine in DNA by bacterial and human DNA dioxygenases. J. Biol.
Chem. 279(39), 40470-40474.
(102)
Delaney, J. C., Smeester, L., Wong, C., Frick, L. E., Taghizadeh, K., Wishnok, J.
S., Drennan, C. L., Samson, L. D., and Essigmann, J. M. (2005) AlkB reverses
etheno DNA lesions caused by lipid oxidation in vitro and in vivo. Nat. Struct.
Mol. Biol. 12(10), 855-860.
(103) Sedgwick, B. (1982) Genetic mapping of ada and adc mutations affecting the
adaptive response of Escherichia coli to alkylating agents. J. Bacteriol. 150(2),
984-988.
(104) Samson, L. (1992) The suicidal DNA repair methyltransferases of microbes. Mol.
Microbiol. 6(7), 825-831.
(105) Vaughan, P., Sedgwick, B., Hall, J., Gannon, J., and Lindahl, T. (1991)
Environmental mutagens that induce the adaptive response to alkylating agents in
Escherichia coli. Carcinogenesis. 12(2), 263-268.
(106) Ahmmed, Z., and Laval, J. (1984) Enzymatic repair of O-alkylated thymidine
residues in DNA: involvement of a 04-methylthymine-DNA methyltransferase
and a 02-methylthymine DNA glycosylase. Biochem. Biophys. Res. Commun.
120(1), 1-8.
(107) McCarthy, J. G., Edington, B. V., and Schendel, P. F. (1983) Inducible repair of
phosphotriesters in Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 80(24), 73807384.
(108) Morimoto, K., Dolan, M. E., Scicchitano, D., and Pegg, A. E. (1985) Repair of
06-propylguanine and 06-butylguanine in DNA by 06-alkylguanine-DNA
alkyltransferases from rat liver and E. coli. Carcinogenesis. 6(7), 1027-1031.
(109) Schendel, P. F. (1981) Inducible repair systems and their implications for
toxicology. Crit Rev. Toxicol. 8(4), 311-362.
180
- Chapter4 (110) Lindahl, T., Sedgwick, B., Demple, B., and Karran, P. (1983) Enzymology and
regulation of the adaptive response to alkylating agents. In Cellular Responses to
DNA Damage (Friedberg, E. C., and Bridges, B. A., Eds.) pp 241-253, Alan R.
Liss, Inc., New York.
(111) Robins, P., Harris, A. L., Goldsmith, I., and Lindahl, T. (1983) Cross-linking of
DNA induced by chloroethylnitrosourea is presented by 06-methylguanine-DNA
methyltransferase. Nucleic Acids Res. 11(22), 7743-7758.
(112) Ludlum, D. B., Mehta, J. R., and Tong, W. P. (1986) Prevention of 1-(3deoxycytidyl),2-(1-deoxyguanosinyl)ethane cross-link formation in DNA by rat
liver 06-alkylguanine-DNA alkyltransferase. Cancer Res. 46(7), 3353-3357.
(113) Lindahl, T., Demple, B., and Robins, P. (1982) Suicide inactivation of the E. coli
06-methylguanine-DNA methyltransferase. EMBOJ. 1(11), 1359-1363.
(114) Demple, B. (1990) Self-methylation by suicide DNA repair enzymes. In Protein
Methylation (Paik, W. K., and Kim, S., Eds.) pp 285-304, CRC Press, Inc., Boca
Raton, Fla.
(115) Dosanjh, M. K., Singer, B., and Essigmann, J. M. (1991) Comparative
mutagenesis of 06-methylguanine and 04-methylthymine in Escherichia coli.
Biochemistry. 30(28), 7027-7033.
(116) Cerda-Olmedo, E., Hanawalt, P. C., and Guerola, N. (1968) Mutagenesis of the
replication point by nitrosoguanidine: map and pattern of replication of the
Escherichia coli chromosome. J. Mol. Biol. 33(3), 705-719.
(117) Guerola, N., Ingraham, J. L., and Cerda-Olmedo, E. (1971) Induction of closely
linked multiple mutations by nitrosoguanidine. Nat. New Biol. 230(12), 122-125.
(118) Reed, J., and Hutchinson, F. (1987) Effect of the direction of DNA replication on
mutagenesis by N-methyl-N'-nitro-N-nitrosoguanidine in adapted cells of
Escherichia coli. Mol. Gen. Genet. 208(3), 446-449.
(119) Demple, B., Jacobsson, A., Olsson, M., Karran, M., and Lindahl, T. (1983)
Isolation of 06-methylguanine-DNA methyltransferase from E. coli. In DNA
Repair-Laboratory Manual of Research Procedures (Friedberg, E. C., and
Hanawalt, P. C., Eds.) pp 41-52, Marcel Dekker, Inc., New York.
(120) Demple, B., Jacobsson, A., Olsson, M., Robins, P., and Lindahl, T. (1982) Repair
of alkylated DNA in Escherichia coli. Physical properties of 06-methylguanineDNA methyltransferase. J. Biol. Chem. 257(22), 13776-13780.
(121) Lindahl, T. (1981) DNA methyl transferase acting on 06-methylguanine residues
in adapted E. coli. In Chromosome Damage and Repair (Seeberg, E., and Kleppe,
E., Eds.) pp 207-217, Plenum Publishing Corp., New York.
181
- Chapter4 (122) Teo, I., Sedgwick, B., Demple, B., Li, B., and Lindahl, T. (1984) Induction of
resistance to alkylating agents in E. coli: the ada+ gene product serves both as a
regulatory protein and as an enzyme for repair of mutagenic damage. EMBO J.
3(9). 2151-2157.
(123) Demple, B., Sedgwick, B., Robins, P., Totty, N., Waterfield, M. D., and Lindahl,
T. (1985) Active site and complete sequence of the suicidal methyltransferase
that counters alkylation mutagenesis. Proc. Natl. Acad. Sci. U. S. A. 82(9), 26882692.
(124) Nakabeppu, Y., Mine, Y., and Sekiguchi, M. (1985) Regulation of expression of
the cloned ada gene in Escherichia coli. Mutat. Res. 146(2), 155-167.
(125) Sedgwick, B., Robins, P., Totty, N., and Lindahl, T. (1988) Functional domains
and methyl acceptor sites of the Escherichia coli ada protein. J. Biol. Chem.
263(9), 4430-4433.
(126) Teo, I. A. (1987) Proteolytic processing of the Ada protein that repairs DNA 06methylguanine residues in E. coli. Mutat. Res. 183(2), 123-127.
(127)
Arikan, E., Kulkarni, M. S., Thomas, D. C., and Sancar, A. (1986) Sequences of
the E. coli uvrB gene and protein. Nucleic Acids Res. 14(6), 2637-2650.
(128) Akimaru, H., Sakumi, K., Yoshikai, T., Anai, M., and Sekiguchi, M. (1990)
Positive and negative regulation of transcription by a cleavage product of Ada
protein. J. Mol. Biol. 216(2), 261-273.
(129) Sedgwick, B. (1989) In vitro proteolytic cleavage of the Escherichia coli Ada
protein by the ompT gene product. J. Bacteriol. 171(4), 2249-2251.
(130) Moore, M. H., Gulbis, J. M., Dodson, E. J., Demple, B., and Moody, P. C. (1994)
Crystal structure of a suicidal DNA repair protein: the Ada 06-methylguanineDNA methyltransferase from E. coli. EMBO J. 13(7), 1495-1501.
(131) Vora, R. A., Pegg, A. E., and Ealick, S. E. (1998) A new model for how 06methylguanine-DNA methyltransferase binds DNA. Proteins. 32(1), 3-6.
(132) Daniels, D. S., and Tainer, J. A. (2000) Conserved structural motifs governing
the stoichiometric repair of alkylated DNA by 0(6)-alkylguanine-DNA
alkyltransferase. Mutat. Res. 460(3-4), 151-163.
(133) Brent, T. P., Dolan, M. E., Fraenkel-Conrat, H., Hall, J., Karran, P., Laval, L.,
Margison, G. P., Montesano, R., Pegg, A. E., Potter, P. M., and. (1988) Repair of
O-alkylpyrimidines in mammalian cells: a present consensus. Proc. Natl. Acad.
Sci. U. S. A. 85(6), 1759-1762.
(134) Rost, B., and Sander, C. (1992) Jury returns on structure prediction. Nature.
360(6404), 540.
182
Chapter 4 (135) Cheng, X., and Roberts, R. J. (2001) AdoMet-dependent methylation, DNA
methyltransferases and base flipping. Nucleic Acids Res. 29(18), 3784-3795.
(136)
Weinfeld,
M., Drake, A. F., Saunders,
J. K., and Paterson,
M. C.
(1985)
Stereospecific removal of methyl phosphotriesters from DNA by an Escherichia
coli ada+ extract. Nucleic Acids Res. 13(19), 7067-7077.
(137)
Lin, Y., Dotsch, V., Wintner, T., Peariso, K., Myers, L. C., Penner-Hahn,
J. E.,
Verdine, G. L., and Wagner, G. (2001) Structural basis for the functional switch
of the E. coli Ada protein. Biochemistry. 40(14), 4261-4271.
(138) Myers, L. C., Verdine, G. L., and Wagner, G. (1993) Solution structure of the
DNA methyl phosphotriester repair domain of Escherichia coli Ada.
Biochemistry. 32(51), 14089-14094.
(139)
Myers, L. C., Terranova, M. P., Nash, H. M., Markus, M. A., and Verdine, G. L.
(1992) Zinc binding by the methylation signaling domain of the Escherichia coli
Ada protein. Biochemistry. 31(19), 4541-4547.
(140)
Myers, L. C., Terranova, M. P., Ferentz, A. E., Wagner, G., and Verdine, G. L.
(1993) Repair of DNA methylphosphotriesters through a metalloactivated
cysteine nucleophile. Science. 261(5125), 1164-1167.
(141) Landini, P., and Volkert, M. R. (2000) Regulatory responses of the adaptive
response to alkylation damage: a simple regulon with complex regulatory
features. J. Bacteriol. 182(23), 6543-6549.
(142) Sakumi, K., and Sekiguchi, M. (1989) Regulation of expression of the ada gene
controlling the adaptive response. Interactions with the ada promoter of the Ada
protein and RNA polymerase. J. Mol. Biol. 205(2), 373-385.
(143) Teo, I., Sedgwick, B., Kilpatrick, M. W., McCarthy, T. V., and Lindahl, T.
(1986) The intracellular signal for induction of resistance to alkylating agents in
E. coli. Cell. 45(2), 315-324.
(144) He, C., Hus, J. C., Sun, L. J., Zhou, P., Norman, D. P., Dotsch, V., Wei, H.,
Gross, J. D., Lane, W. S., Wagner, G., and Verdine, G. L. (2005) A methylationdependent electrostatic switch controls DNA repair and transcriptional activation
by E. coli ada. Mol. Cell. 20(1), 117-129.
(145)
Springman, E. B., Angleton, E. L., Birkedal-Hansen,
H., and Van Wart, H. E.
(1990) Multiple modes of activation of latent human fibroblast collagenase:
evidence for the role of a Cys73 active-site zinc complex in latency and a
"cysteine switch" mechanism for activation. Proc. Natl. Acad. Sci. U. S. A. 87(1),
364-368.
183
- Chapter4 (146) Friedberg, E. C., Walker, G. C., Siede, W., Wood, R. D., Schultz, R. A., and
Ellenberger, T. DNA Repair and Mutagenesis (2005) ASM Press, Washington
D.C.
(147) Nakamura, T., Tokumoto, Y., Sakumi, K., Koike, G., Nakabeppu, Y., and
Sekiguchi, M. (1988) Expression of the ada gene of Escherichia coli in response
to alkylating agents. Identification of transcriptional regulatory elements. J. Mol.
Biol. 202(3), 483-494.
(148) Sedgwick, B. (1987) Molecular signal for induction of the adaptive response to
alkylation damage in Escherichia coli. J. Cell Sci. Suppl. 6:215-23., 215-223.
(149) Shevell, D. E., bou-Zamzam, A. M., Demple, B., and Walker, G. C. (1988)
Construction of an Escherichia coli K-12 ada deletion by gene replacement in a
recD strain reveals a second methyltransferase that repairs alkylated DNA. J.
Bacteriol. 170(7), 3294-3296.
(150) Saget, B. M., and Walker, G. C. (1994) The Ada protein acts as both a positive
and a negative modulator of Escherichia coli's response to methylating agents.
Proc. Natl. Acad. Sci. U. S. A. 91(21), 9730-9734.
(151) Rebeck, G. W., Coons, S., Carroll, P., and Samson, L. (1988) A second DNA
methyltransferase repair enzyme in Escherichia coli. Proc. Natl. Acad. Sci. U. S.
A. 85(9), 3039-3043.
(152) Rebeck, G. W., and Samson, L. (1991) Increased spontaneous mutation and
alkylation sensitivity of Escherichia coli strains lacking the ogt 06-methylguanine
DNA repair methyltransferase. J. Bacteriol. 173(6), 2068-2076.
(153) Potter, P. M., Wilkinson, M. C., Fitton, J., Carr, F. J., Brennand, J., Cooper, D. P.,
and Margison, G. P. (1987) Characterisation and nucleotide sequence of ogt, the
06-alkylguanine-DNA-alkyltransferase gene of E. coli. Nucleic Acids Res.
15(22), 9177-9193.
(154) Takano, K., Nakamura, T., and Sekiguchi, M. (1991) Roles of two types of 06methylguanine-DNA methyltransferases in DNA repair. Mutat. Res. 254(1), 3744.
(155) Potter, P. M., Brennand, J., and Margison, G. P. (1986) Lack of sequence
homology between a fragment of E. coli DNA encoding an 06-methylguanine
methyltransferase and the ada gene. Br. J. Cancer 54, 366-367.
(156) Sassanfar, M., Dosanjh, M. K., Essigmann, J. M., and Samson, L. (1991)
Relative efficiencies of the bacterial, yeast, and human DNA methyltransferases
for the repair of 06-methylguanine and 04-methylthymine. Suggestive evidence
for 04-methylthymine repair by eukaryotic methyltransferases. J. Biol. Chem.
266(5), 2767-2771.
184
- Chapter4 (157)
Nakabeppu, Y., Kondo, H., Kawabata, S., Iwanaga, S., and Sekiguchi, M. (1985)
Purification and structure of the intact Ada regulatory protein of Escherichia coli
K12. 06-methylguanine-DNA methyltransferase. J. Biol. Chem. 260(12), 72817288.
(158) Sedgwick, B. (1983) Molecular cloning of a gene which regulates the adaptive
response to alkylating agents in Escherichia coli. Mol. Gen. Genet. 191(3), 466472.
(159) Volkert, M. R., and Nguyen, D. C. (1984) Induction of specific Escherichia coli
genes by sublethal treatments with alkylating agents. Proc. Natl. Acad. Sci. U. S.
A. 81(13), 4110-4114.
(160)
Boyle, J. M., Durrant, L. G., Wild, C. P., Saffhill, R., and Margison, G. P. (1987)
Genetic evidence for nucleotide excision repair of 06-alkylguanine in mammalian
cells. J. Cell Sci. Suppl. 6:147-60., 147-160.
(161) Bronstein, S. M., Skopek, T. R., and Swenberg, J. A. (1992) Efficient repair of
06-ethylguanine, but not 04-ethylthymine or 02-ethylthymine, is dependent
upon 06-alkylguanine-DNA alkyltransferase and nucleotide excision repair
activities in human cells. Cancer Res. 52(7), 2008-2011.
(162)
Huang, J. C., Zamble, D. B., Reardon, J. T., Lippard, S. J., and Sancar, A. (1994)
HMG-domain proteins specifically inhibit the repair of the major DNA adduct of
the anticancer drug cisplatin by human excision nuclease. Proc. Natl. Acad. Sci.
U. S. A. 91(22), 10394-10398.
(163) Voigt, J. M., Van, H. B., Sancar, A., and Topal, M. D. (1989) Repair of 06methylguanine by ABC excinuclease of Escherichia coli in vitro. J. Biol. Chem.
264(9), 5172-5176.
(164) Samson, L., Han, S., Marquis, J. C., and Rasmussen, L. J. (1997) Mammalian
DNA repair methyltransferases shield 04MeT from nucleotide excision repair.
Carcinogenesis. 18(5), 919-924.
(165) Huang, J. C., Hsu, D. S., Kazantsev, A., and Sancar, A. (1994) Substrate
spectrum of human excinuclease: repair of abasic sites, methylated bases,
mismatches, and bulky adducts. Proc. Natl. Acad. Sci. U. S. A. 91(25), 1221312217.
(166) Edara, S., Kanugula, S., and Pegg, A. E. (1999) Expression of the inactive
C145A mutant human 06-alkylguanine-DNA alkyltransferase in E.coli increases
cell killing
and mutations
by N-methyl-N'-nitro-N-nitrosoguanidine.
Carcinogenesis. 20(1), 103-108.
185
CHAPTER 5
Effect of Mismatch Repair Proteins on Methyltransferase
Repair of 06-Methylguaninein Vivo
186
- Chapter5 5.1. ABSTRACT
DNA repair is essential for fighting the adverse effects of base damage to the cell.
One example of base damage, 06-methylguanine (06mG), stably pairs with thymine
during replication and thereby creates a promutagenic 06mG:T mismatch.
This
mismatch has also been linked with cellular toxicity. Therefore, in the absence of repair,
06mG:T mismatches can kill the cell, or, upon the next round of replication, result in G:C
--, A:T transition mutations. The Ada and Ogt methyltransferase (MTase) proteins can
both directly reverse the 06mG base damage to yield guanine. When the 06mG is in a
G:C pair, this reaction poses little problem for the cell. However, if the
6mG has
already been replicated, and an 06mG:T pair exists, the consequences for the cell are
much more complicated. If the 06mG:T pair is biochemically converted to a G:T pair,
the G:T mispair must still be corrected to G:C in order to avoid mutation. Two pathways
in E. coli can convert G:T mispairs to the native G:C base pairing. These are the Methyldirected Mismatch Repair (MMR) and the Very Short Patch Repair (VSPR) pathways.
The use by the cell of VSPR has distinct advantages. In this work, we hypothesized that
E. coli may coordinate MTase repair with VSPR as an efficient means to restore harmful
06mG:T mismatches to normal G:C base pairs. To test this hypothesis, we analyzed the
relative efficiencies of MTase repair of 06mG (by Ada, Ogt, and both) in cells that lacked
functional mismatch repair proteins MutS, MutL, MutH, or Vsr. We found that MTase
repair of 06mG is positively influenced by the MutS, MutL, and Vsr proteins.
In
contrast, cells that lacked MutH showed wild type levels of MTase repair. These results
are consistent with the notion that VSPR positively influences MTase repair of 06mG:T
mismatches to avoid the detrimental effects of this mispairing.
187
- Chapter5 5.2. INTRODUCTION
Alkylating agents produce a variety of DNA lesions by reacting at a variety of
positions on the DNA bases. The cellular impact of these lesions will depend on their
mutagenic and cytotoxic properties, as well as the extent to which they are repaired. One
particular type of DNA alkylation damage, O6-methylguanine (6mG), has been given
much attention due to its mutagenicity (1-4) and cytotoxicity (5-7). Extensive in vivo
mutagenesis studies done in our laboratory established that OmG codes as an adenine
during replication in E. coli nearly 100% of the time (8). As a result, this lesion is
responsible for G:C -+ A:T transition mutations following a second round of replication.
The O6mG:T base pair has also been shown to be a substrate for methyl-directed
mismatch repair (MMR) (9-11). It is hypothesized that the toxicity of O6mGresults from
reiterative and futile attempts to replace the mismatched T with a base that does not
recreate a substrate for another round of MMR (12). This process can lead to persistent
repair gaps and, ultimately, toxic double strand breaks. Therefore, repairing O6mG is
paramount to avoiding the lethality of alkylating agents.
In E. coli, repair of O6mG is primarily the responsibility of two methyltransferase
(MTase) proteins Ada and Ogt (6). Both of these proteins can repair O6mG directly by
transferring the methyl group to an active site cysteine residue (13-16).
This
methyltransfer is not reversible and therefore inactivates the protein (17). For this reason,
the MTase proteins are not true enzymes and are commonly referred to as "suicide
proteins". In the case of the Ada protein, repair of methylphosphotriesters, which are
normally formed alongside O6mG, leads to upregulation of Ada through a process called
the adaptive response (18). This response enables the repair of high levels O6mG
associated with alkylation pressure (14). In the case where MTase repair of 06mG
follows one round of replication, a mismatched G:T base pair results.
To prevent
mutation, this mismatch must be further processed before replication.
G:T mismatches are a substrate for two different types of Mismatch Repair (MR),
Methyl-directed Mismatch Repair (MMR) (19) and Very Short Patch Repair (VSPR)
188
- Chapter5 (20). Both systems are stimulated by the activities of the MutS and MutL mismatch
recognition proteins, although the downstream mechanisms of repair initiation and
completion are quite different.
MMR is only active for a short time period following replication, owing to the
hemimethylated state of Dam d(GATC) sites (21). Once the Dam methylase has had
enough time to methlyate the newly synthesized strand, MutH can no longer initiate
repair by incising the DNA backbone (22).
Therefore, MMR is time limited.
Additionally, incision by MutH at an adjacent d(GATC) site initiates repair as far as two
kilobases away from the mismatch (23). Following incision by MutH, strand removal is
accomplished through the concerted efforts of many proteins including DNA helicase II,
single stranded DNA binding protein, and Exo I, Exo VII, Exo X, or Rec J (24). DNA
Polymerase III must then resynthesize the DNA strand from the d(GATC) site to the
mismatch. DNA Ligase completes the repair event by sealing the DNA nick. In total,
repairing the G:T mismatch via MMR is of significant metabolic cost to the cell.
VSPR of G:T mismatches is initiated by the Vsr endonuclease, which in all cases
creates a single stranded nick on the 5' side of the mismatched T (25). Pol I then
removes and replaces a small number of bases ( < 10) 3' to the nick utilizing its 5' to 3'
exonuclease and polymerase activities (26). DNA Ligase then seals the nick to complete
the restoration of the native G:C pairing. Given that the repair patch created by VSPR is
much smaller than that for MMR, the VSPR pathway is significantly more efficient.
The primary purpose of VSPR is to repair G:T mismatches that arise through the
deamination of the Dcm product, 5-methylcytosine (27). Dcm methylates the 5 position
of the second cytosine of the sequence 5'- CCAGG -3' (28). Therefore Vsr most
commonly initiates repair on G:T mismatches in the 5'- CTAGG -3' sequence context
(where the T is mismatched with a G). However, Vsr has been shown to have significant
endonuclease activity for G:T mismatches in a variety of sequence contexts (29;30).
Therefore, VSPR could play a role in repairing G:T mismatches that arise through MTase
repair of O6mG:Tbase pairs, as well.
189
- Chapter5 -
Upon inspection, it appears the most efficient route to repairing O6mG:T base
pairs is through the methytransferase repair of O6mG followed by VSPR of the resulting
G:T mismatch. Additionally, it would seem that the proposed competition between MutS
and Ada for the O6mG:Tpair would be inefficient, and perhaps unlikely, for cell survival.
We therefore hypothesized that the MTases might act in conjunction with proteins in the
VSPR pathway, to restore cooperatively O6mG:Tmismatches to G:C base pairs.
To test our hypothesis that MTase repair can be positively affected by MR
proteins, we utilized a M13 based in vivo MTase assay in a number of cell strains. We
constructed a matrix of mutant E. coli that are defective in MTase activities (Ada, Ogt, or
both) and a single mismatch repair protein (MutS, MutL, Vsr, and MutH). In total 20 cell
strains were assayed for their relative MTase repair efficiency, allowing for a
comprehensive view of how MMR and VSPR proteins affect the repair of O6mG by
MTase proteins. Remarkably, both MutS and MutL were observed to play a cooperative
role with Ada and Ogt. Additionally, while cells deficient in MutH showed no difference
to wt, cells lacking Vsr showed a significant decrease in MTase repair efficiency. These
data represent the first direct examination of how MTase repair is affected by MR
proteins, and assist in our understanding of how repair systems interact in vivo.
190
- Chapter5 5.3. EXPERIMENTAL PROCEDURES
Preparation of E. coli Genomic DNA. Preparation of E. coli genomic DNA was carried
out essentially as described in Current Protocols (31). Briefly, 1 mL of cells from an
overnight culture was spun down and the resulting cell pellet was resuspended in 570 gIL
TE 8.0 (10 mM Tris, 1 mM EDTA). After the addition of 30 tL of 10% SDS and 1.8
units of Proteinase K (Roche), the solution was incubated at 37 °C for 1 h. This solution
was then supplemented with 100 gtL of 5 M NaCl, 80 gL of a 10% CTAC/0.7 M NaCl
solution, and incubated at 65 C for 10 minutes. This solution was extracted twice from
750 !IL of a 25:24:1 ratio of phenol to chloroform to isoamyl alcohol (Invitrogen). DNA
was precipitated from the final aqueous layer with 420 }xLof isopropanol, and the DNA
pellet was washed with 250 !aL of 70% Ethanol. The resulting DNA was dried in vacuo
for 5 minutes, resuspended in 100 .L TE, and quantified using UV spectrometry.
Preparation of Recombination Substrates. The TnlO marker, which codes for TetR_N
and TetR_C, was amplified from GM5555 (obtained from M. Marinus, U. Mass.
Worcester) genomic DNA using primers that contained flanking sequences to match the
target gene (Table 5.1). The PCR reaction was carried out according to Stratagene's
Opti-prime PCR Optimization Kit.
The combination of Taq polymerase (NEB),
buffering condition #11, and 2% DMSO as a reaction adjunct provided the best results.
The thermocycling parameters were as follows: 94 C for 90 sec, then thirty rounds of 95
°C for 30 sec, 56 C for 30 sec, 72 C for 60 sec, and finally 10 min at 72 C. The 2 kb
product, identified by separating a fraction of the reaction products on a 1% agarose gel,
was purified using the QIAquick PCR Purification kit.
Transformationof RecombinationSubstratesand Isolationof Recombinants. KM22 cells
(obtained from M. Marinus, U. Mass Worcester) (32) were prepared for transformation
by diluting 100 aL of an overnight culture in 10 mL LB plus 1 mM IPTG, growing (roller
drum, 37 C) until OD60 0 = 0.6, washing the cells twice with 15 mL cold water, and
resuspending the final cell pellet in 75 gpL1 mM MOPS/20% glycerol. Approximately
100 tL of cells and 500 ng of recombination substrate was combined in a chilled
191
- Chapter5 electroporation cuvette and a 2.5 kV pulse at 129 Ohms was applied using a Electro Cell
Manipulator 600 electroporation system (Bantex). Immediately after electroporation,
cells were transferred to a culture tube containing 3 mL LB and 1 mM IPTG and grown
at 37 °C as above. After three hours, cells were harvested by centrifugation, resuspended
in 100 FL LB, plated on a LB plate containing 4 Rg/mL Tetracycline, and incubated over
night at 37 °C. The next day, successful transductants were picked and grown over night
in liquid culture with 12 g/mL Tetracycline. Cultures were then analyzed individually
for the correct genotype by isolating their genomic DNA and performing PCR as
described above using primers to amplify the gene of interest (Table 5.2).
Preparation of Knock Out Plvir Lysates (aka "Up Infection"). A 1:100 dilution of
KM22 donor cells, which were grown over night in LB containing 12 ttg/mL
Tetracycline and 1 mM IPTG, was grown to mid log phase (OD600 =0.5) in 5 mL LB
supplemented with 5 mM CaC12. Serial dilutions of P1 vir (gift of Walker Lab, MIT),
ranging from 100 to 10-5 of stock (109 pfu/ gL), were prepared in LB. P1 infections (six
total) were prepared to contain 100 gL of cells and 10 gtL of one of the five P1 vir
dilutions. A control which had no virus was done in parallel for comparison. These
mixtures were incubated in a capped 1.5 mL tube at 37 C for 20 minutes. Infections
were then mixed with 3.5 mL of R-top agar (51 C)(per L: 10g Bacto tryptone, g Bacto
yeast extract, 8g Difco agar, 8g NaCl, add 2 mL 1 M CaC12 and 5 mL 20% glucose after
autoclaving) and immediately spread on R-plates (as R-top agar but 12 g Difco agar).
The agar layer solidified after 10 minutes and the plates were incubated at 37 C over
night.
The following day, virus was harvested from the plate displaying the most
discernable plaques. A spatula was used to scrape and transfer the agar layer to a 15 mL
centrifuge tube, and two 1 mL volumes of LB were used to collect all virus from the
plate. The tube containing the agar was supplemented with 300 CtLchloroform, vortexed
on high for 30 seconds to elute the virus, and spun at 7,000 xg for 10 minutes to pellet the
insoluble material. Approximately 1.75 mL of P1 vir was transferred to a sterile 2 mL
glass vial, supplemented with 50 1tL chloroform, and stored at 4 °C. Serial dilutions of
this virus (100 to 10-5) were used to infect recipient cells.
192
- Chapter5 Generation of Knock Out Transductants(aka "DownInfection"). A 1:100 dilution of
recipient cells was grown to mid log phase (OD6 0 0 =0.5) in 5 mL LB, supplemented with
500 gtLof a solution containing 1 M MgC12 and 50 mM CaCl2, vortexed, and incubated at
room temperature for 20 minutes. For infection, 100 jiL of cells and 10 [iL of a viral
dilution were incubated at 37 C for 20 minutes in a capped 1.5 mL tube. A control
which had no virus was done in parallel for comparison. An equal volume of 250 mM
sodium citrate was added to stop the infection, and cells were allowed to recover at 37 °C
L) was spread on a LB plate
for 3 hours.
The entire contents of each tube (200
containing 4
g/mL Tetracycline and incubated over night at 37 C. The next day,
successful transductants were picked and grown over night in liquid culture with 12
[tg/mL Tetracycline. The genomic DNA from individual cultures was then analyzed for
the correct genotype using PCR, agarose gel separation, and sequencing techniques as
described above.
One stock, possessing the correct genotype, was chosen for all
subsequent experiments.
Preparation of M13mp7L2 (M13) DNA. A 5 mL culture of NR9050 cells grown over
night in LB was diluted 1:5 in 5 mL of 2xYT and grown (roller drum, 37 C) for 1.5 h.
Once the cells reached mid-log phase, 2 mL were used to inoculate 1 L of 2xYT, in a 2 L
baffled flask, which then grew (275 rpm, 37 C) for 2.5 h. A 2 mL volume of saturated
M13mp7L2 progeny phage (gift of Sarah Delaney) was added to the cells which then
grew for an additional 9 h. To isolate progeny phage, the mixture was incubated at 0 °C
for 10 min, the cells were pelleted by centrifugation (9.5 krpm, 10 min), and the
supernatant was decanted into sterile tubes (6 tubes, 165 mL/tube). The phage in each
supernatant was precipitated by adding /4 volume (42 mL) of 20% PEG/2.5 M NaCl and
incubating over night at 4 C. Precipitated phage was collected by centrifugation (9.5
krpm, 10 min), resuspended in a total volume of 19 mL TE (10 mM Tris-HCI, 1 mM
EDTA
(pH
8.0)),
split
into
thirds,
and
extracted
5
times
with
3
mL
phenol:chloroform:isoamyl alcohol (25:24:1 v/v/v, Invitrogen). The aqueous phase (-18
mL) was applied to a hydroxylapatite column (1 g DNA grade resin (BioRad))
conditioned with 5 mL TE, and eluted using moderate air pressure. The column was
washed with 5 mL TE and M13 DNA was eluted using 20 mL total of 140 mM
193
- Chapter5 potassium phosphate buffer. Each of the 1 mL fractions was analyzed for DNA content
by separating 10 L on a 1% agarose gel in TAE, staining with ethedium bromide, and
visualizing the gel over a transilluminator. The 6 fractions containing the most amount of
M13 DNA were concentrated and dialyzed through 3 Centricon-100 columns (Amicon)
by centrifugation followed by 3 x 2 mL TE washes. The M13 DNA was collected in a
final volume of -250
L TE.
The concentration of genome was quantified by
spectrophotometry using the conversion factor 14 pmol/mL = 1 A260 unit. Typical yields
of M13 DNA were approximately 600 pmoles/L of culture.
Construction of 06mG - M13 Viral Genome. Genomes were constructed on a 24 pmole
scale. Single-stranded M13 DNA (2.4 pmole/gL) was linearized by cleavage with EcoRI
(70 U, NEB) for 8 h at 23 C in a total of 50 FL of the supplier's buffering conditions.
Cleavage occurs at a hairpin containing a single EcoRI site (33). The linearized M13
DNA was annealed with 1.25 equivalents of each scaffold 5'- GGT CTT CCA CTG AAT
CAT GGT CAT AGC -3' and 5'- AAA ACG ACG GCC AGT GAA TTG GAC GC -3'
(sequences that are partially complementary to the 5' and 3' sides of the insert and the
genomic DNA termini) by heating the mixture to 50 °C for 5 min and cooling linearly to
0 C over 50 min (33). The O6mG lesion containing insert, 5'- GAA GAC CTX GGC
GTC C -3' was phosphorylated on the 5'-terminus with T4 PNK (10 U, NEB) using the
supplier's buffering conditions supplemented with an excess of ATP (Roche).
The
phosphorylated insert (30 pmoles), the annealed scaffold mixture (24 pmol), 1 L of 100
mM ATP (Roche), and 2 pL of T4 DNA Ligase (800 U, NEB) were mixed in a final
reaction volume of 55 L and incubated at 16 °C for 20 h to recircularize the genome
(33-35). The scaffold DNA was removed using the exonuclease activity of T4 DNA
polymerase (36;37) by incubating the mixture at 37 °C for 6 h after the addition of 3 gL
enzyme (10.5 U, NEB). The genome construct mixture was brought to a final volume of
100 L with water, extracted twice with 100 jiL phenol:chloroform:isoamyl alcohol
(25:24:1 v/v/v, Invitrogen), and desalted using G-50 Sephadex Quick Spin Columns
(Roche).
194
- Chapter5 Quantification of Viral Genome.
Quantification of genome constructions was
accomplished by annealing 15 pmol of the 5'[3 2 P]-labeled 30 mer probe 5'- TCC CAG
TCA CGA CGT TGT AAA ACG ACG GCC -3' to 5 L of genome construct and a
range (3.9, 7.8, 15.6, 31.2, 62.5, 125, 250, and 500 fmoles) of wt M13 DNA standards.
The probe was designed to anneal in a region of the M13 remote from the cloning/insert
site. The annealing solutions consisted of 100 mM NaCl, 4.2% Ficoll (Amersham),
0.042% bromophenol blue (ICN), and 0.042% xylene cyanol FF (ICN) in a volume of
15.5 L. After incubating at RT for 1 h, the genome probe mixtures were run on a 1%
agarose gel in lx TAE buffer for 4 h at 100 V, after which the free probe front was
excised. The gel was run for an additional 3 h and then transferred onto a glass plate and
dried under a box fan for 36 h. The amount of circular genome was quantified by using
phosphorimagery to compare its signal intensity to the signal intensities for the standards.
Typical yields were in the range of 5% (1.2 pmoles), which was enough for hundreds of
electroporations.
Determinationof Relative Rates of O6mGRepair in Strains of Differing Genotype. To
prepare cells for electroporation, an over night grown in LB (containing 50
g/mL
Kanamycin, 12 g/mL Chloroamphenicol, or 12 tg/mL Tetracycline where applicable)
was diluted 1:50 in 10 mL LB (4 replicates), and grown to mid-log phase (OD60 0 = 0.5).
Cells were transferred into 15 mL tubes, harvested by centrifugation (6,000 xg, 10 min),
washed twice with 15 mL 4 °C water, and resuspended in 75 L 10% glycerol. Chilled
electrocuvettes were then supplemented with 100 pL of competent cells (-2 x 109 cells)
and -3.5 fmoles of the O6mG-M13 viral genome for transformation.
The cell/genome mixtures were electroporated (2.5 kV, 129 ohms in a 0.2-cm
cuvette) using a Electro Cell Manipulator 600 electroporation system (Bantex), and
transferred to 10 mL LB. Fractions of this dilution (20, 75, and 200 !IL) were transferred
to a tube containing 3.75 mL overlay mix, which was then vortexed and immediately
spread over the surface of a B-broth plate (per L; 10g tryptone, 8g NaCl, 20g bacto agar).
195
- Chapter5 Overlay mix was prepared by adding 375 IL of a solution containing 300 tL
plating bacteria (NR9050 over night diluted 1:5 in 2 x YT and grown on a roller drum at
37 °C for 1.5 h), 25 jiL 1% thiamine, 10 gL of 100 mM IPTG, and 40 L of 40 mg/mL
X-gal in DMF to 3 mL B-broth soft agar (10 g/L tryptone, 8 g/L NaCl, and 6 g/L) Bacto
agar) just before use.
Once completing all the electroporations and platings, the soft agar mix on the
top of each plate was allowed to cool for an additional 10 min on the bench. Plates were
then put at 37 °C, up side down, over night. After incubation for 16-20 h, the number of
light blue and dark blue plaques were counted and the ratio of darks to total was used to
calculate percent repair. Of the three dilutions plated for each electroporation, the one
which yielded the highest number of discernable plaques was counted (approximately
100 to 800 total plaques (Average = 421).
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- Chapter5 5.4. RESULTS
Genotypic Characterization of E. coli Strains. Before constructing the MMR protein
mutants, the parental C215 (wt), C216 (ogf.::kan), C217 (ada-,alkB:.Cam), and C218
(ogt-::kan, ada-::Cam) strains (8;38-40) (as FC215 derivatives) were characterized
genotypically. PCR was conducted on genomic DNA using primer sets (Table 5.2) that
annealed to ogtf or ada- gene sequences outside of the locations altered by antibiotic
resistance marker insertion (Figures 5.1 and 5.2). The PCR products obtained were
separated via 1% agarose gel in TAE and compared against their expected fragment sizes
(Table 5.3).
In all cases the size of the fragments obtained by PCR matched our
expectations.
Resulting PCR fragments were also sequenced using the 5' primer sequence
utilized for amplification.
Open reading frames within the sequencing data were
identified using NEB cutter (41) and analyzed using BLAST software (42). BLAST
results confirmed the presence of native gene sequences in the appropriate strains. For
the ogf strains, BLAST results yielded conserved domains matching the aminoglycoside
3'-phosphotransferase family, a member of which confers resistance to Kanamycin. For
the
ada- strains,
BLAST
results
yielded
conserved
domains
matching
the
chloramphenicol acetyltransferase protein.
Genotypic characterization for the mutH, mutL-, mutS, and vsf strains was
conducted as above. PCR was conducted on genomic DNA using primer sets (Table 5.2)
that annealed to mutH, mutL-, mutS, and vsr gene sequences outside of the locations
altered by antibiotic resistance marker insertion (Figures 5.3, 5.4, 5.5 and 5.6). In each
case, the size of the PCR product (Table 5.3) was easily matched to that of the native
gene (analyzed in parallel as a negative control) or the insertion deletion mutant. PCR
products from successful recombinants were analyzed, as above, by sequencing and
BLAST analysis. Results indicated that, as expected, the majority of the target gene was
indeed replaced by conserved domains matching the TetR_N and TetR_C proteins.
197
- Chapter5 -
Rates of EtmG Methyltransferase Repair.
The twenty cell strains assayed for their
relative rates of &0mG repair showed a wide range of MTase activity (Tables 5.4, 5.5,
5.6, and 5.7). Firstly, the data from the C215, C216, C217, and C218 parental strains
(Figure 5.7, black columns) demonstrates good correlation between percent repair of
06mG and MTase status of the cell. As expected, the C215 wt strain that expresses both
functional methyltransferases shows a high amount of lesion repair (96 ± 1%). The C216
strain which lacks Ogt protein shows significantly less MTase ability (69
i
3%), while
the C217 strain which lacks Ada protein shows near wt levels of MTase repair (95 ±- 1%).
The asymmetry in the results for these single knock outs were not surprising as the
normal cell contains roughly thirty Ogt molecules (15) and one molecule of Ada (43).
Therefore, cells which lack the predominant MTase under normal conditions, Ogt, are
expected to display the more significant decrease in MTase ability. Accordingly, the
C218 double mutant shows very poor repair of 06mG (6 ± 1%).
The parental strains bearing the mutHmutation (Figure 5.7, light grey columns)
showed MTase repair results nearly identical to those for the parental strains (Figure 5.7,
black columns). The percent repair data for C215H, C216H, C217H, and C218H strains
were 97 +t 1, 70 ± 2, 96 ± 2, and 6
i
1, respectively. These data indicate that the MutH
protein does not affect the MTase repair of 06mG by Ada or Ogt.
In contrast, mutS, and mutL- strains displayed levels of repair significantly
lower than those for the C215, C216, and C217 parental strains. The effect of knocking
out mutL- from the C215, C216, and C217 parental strains (Figure 5.7, grid columns)
resulted in a decrease in repair to -70% that of wild type (70, 75, and 66% of wt,
respectively). Though there was variation between the mutL mutants, there appeared to
be no significant differences between them. On the other hand, results for the mutS
knock outs showed more variation between the MTase backgrounds. Mutation of mutS
(Figure 5.7, dark gray columns) in the C215 background caused a modest decrease in
percent repair to 71% that for the parent strain. A more dramatic decrease was observed
by mutating mutS in the C216 background. Here the mutS mutant showed repair 62% of
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- Chapter5 that for the parental strain. The largest effect of mutating this protein was observed in the
C217 background where mutating mutS caused a drop in repair to only 41% that for the
parent cells. Cells lacking the functions of MutL and MutS clearly are less efficient at
repairing 06mG.
Remarkably, the vsr strains (Figure 5.7, white columns) also displayed less
efficient repair of 06mG as compared to their parent lines. Knocking out vsf from the
C215 background conferred a small decrease in repair to 84% that for the parent.
Absence of Vsr from the MTase single knockouts however dropped the levels of 06mG
repair more significantly. Mutating vsr in the C216 background dropped MTase repair to
59% that for the parent. The level of MTase repair in the vsr mutant form of C217 was
75% of that for the parent.
The level of 06mG repair for the C218 derivatives was, in all but one case,
consistently low. The exception was the C218L strain that displayed 44 + 4% repair of
O6mG. If reproducible, this result would be interesting from the standpoint of how the
absence of MutL protein confers a dramatic increase in repair of O6mG in cells lacking
both MTase proteins. An experiment to confirm of this result is underway.
199
- Chapter5 5.5 DIscUSSION
PCR mediated gene replacement (32) was used to construct sixteen mutant
forms of E. coli that contained differing MTase capacities and were deficient in either
mutH, mutL, mutS, or vsr. We chose to construct these mutants such that the vast
majority of the target gene would be replaced by the marker for tetracycline resistance.
We designed the recombination sequences such that approximately 150 bases on each
end of the target gene would be preserved (Figures 5.3 to 5.5, from the end of the gene to
the opposite end of recombination sequence). The genetic information central to the
recombination sequences, however, would be replaced by TetR upon recombination with
the targeted substrate generated by PCR (Figure 5.8) (32). Recombination, between the
target gene in the cell and the PCR generated substrate, is carried out by KM22 cells that
have recBCD replaced with A red (32). These cells facilitate the production of mutants
because no prior cloning of the target gene is necessary, and the frequency of
recombination is 10 to 100-fold higher than in rec proficient strains (44). Employing this
method allowed us to generate our mutant KM22 donor cells relatively quickly.
Subsequent transfer our alleles into recipient C215, C216, C217, and C218 strains was
carried out by generalized P1 vir techniques as described in the experimental procedures
section.
The assay we used to measure the relative efficiencies of MTase repair has been
described previously (8;33). Upon transformation of the O6mG-M13 genome into E.
coli, the ability of the cell to repair the lesion prior to replication determines if the codon
sequence containing the lesion encodes for a Trp or a stop codon (Figure 5.9). If repair
of @0mGdoes not precede replication, the replication machinery interprets the highly
mutagenic lesion as an A within a TAG stop codon. Subsequently, transcription of the afragment of f-Galactosidase is terminated.
If, however, 06mG is repaired before
replication, the codon in which it is located, TGG (Trp), allows read through and the afragment of P-Galactosidase is transcribed and translated. Subsequently, this fragment
associates with the co-fragment of the protein, provided by the host cell, and gives rise to
active P3-Galactosidase. This protein cleaves the chromogenic substrate X-gal to yield a
200
- Chapter5 dark blue plaque. Therefore, the percent of total plaques that are dark blue can be used to
relate the efficiency of repair.
The efficiency to which a O6mG lesion is repaired in E. coli depends on the
number of MTase molecules present in the cell and the affinities of those MTases for the
lesion. During exponential growth, approximately thirty molecules of Ogt (15) and one
molecule of Ada (43) are present.
On this basis, one would expect to observe a
significantly lower level of repair in the C216 background (ogt-::kan) as compared to the
C217 background (ada-::Cam). The affinity of Ada for 06mG, however, is greater than
that for the Ogt protein (45) and therefore the difference in repair observed for the ada
and ogt mutants reflects a multitude of parameters. Therefore, the mutant strains were
grouped according to the MTase background of the parent cell.
Interestingly, we observed data that were consistent with a small amount of
O6mG repair in the C218 (ada-, ogt) double mutant strain bearing no MTase activity.
There are two potential ways to rationalize this result. First, nucleotide excision repair
(NER) has been observed to repair these lesions (38;46-50) especially in the absence of
MTase activity.
Alternatively, the repair signal observed could be due to a small
population of Ml13 genome containing a G instead of O6mG. The synthetic route used to
produce the 16-mer ODN containing the 06mG lesions utilizes a deprotection step that is
known to yield small amounts of the unmodified oligonucleotide (51). A small amount
of viral genome containing G at the lesion site would manifest itself as "repair" by
yielding a dark blue plaque. Mass spectrometry analysis of the lesion containing 16-mer
confirmed the presence (-5%) of a species consistent with G at the lesion site. Therefore,
a small amount of systematic error is present in the repair percentages calculated, and
results in slightly elevated numbers for percent repair of O6mG.
Strains lacking mutS displayed a decrease in MTase repair. These results were
interesting as MutS binds to 06mG base pairs (10) and can presumably shield the lesion
from MTase repair. The mechanism by which the presence of MutS confers increased
MTase repair of O6mG therefore likely involves (1) identification of the lesion by MutS
201
- Chapter5 followed by (2) subsequent dissociation of MutS and (3) signaling the presence of a
mismatch to downstream repair proteins. The MutL protein has been shown to assist
MutS in mismatch presentation by stimulating the formation of an a-shaped DNA loop
containing the mismatch (52). The observation that mutL mutants display decreases in
MTase repair similar to the decreases for mutS mutants is in line with previous
observations that mutating either gene results in an identical defect in mismatch
recognition (53). It therefore appears that a novel role of the MutS and MutL mismatch
recognition proteins in assisting MTase repair has been discovered. More studies are
necessary to examine the exact mechanism of this effect.
It could be the case, for example, that MutS and MutL recruit Vsr to the 06mG
lesion site, and Vsr is responsible for stimulating MTase repair (Figure 5.10). Consistent
with this idea, strains that lack vsr showed a decrease in MTase repair (Figure 5.7, white
columns). A possible role for Vsr dependent stimulation of MTases could be to increase
the chances that Vsr initiates repair on the G:T mismatches that result from MTase repair
of a 06mG:T base pair. Subsequently, VSPR, and not MMR, restores mismatch to the
native G:C pairing with high efficiency. This model seems particularly attractive from
multiple perspectives. First, it is the most energetically favorable mechanism by which to
process cemG:T mismatches to G:C base pairs. Second, it might be that the cell has no
other choice. Given that 06mG:T mismatches arise through the replication of the lesion
containing strand, MMR would be targeted to the strand opposite the lesion and
reiteratively regenerate a MMR substrate. Eventually, if the cell does not die, the DNA
becomes fully methylated at the d(GATC) sites and MMR becomes inactive. At this
point, correct repair of 06mG:T mismatches depends on MTase repair followed by
VSPR. Third, it is likely that the efficiency of these two repair systems has helped define
cellular survival.
If cells have been continually faced with this scenario, it seems
plausible that cooperation between the MTase and VSPR proteins would evolve over
time to produce a cell that is more fit.
202
- Chapter5 5.6. FUTURE DIRECTIONS
Bacterial Two-Hybrid Studies between MTases and VSPR Proteins
A question raised from the data in Figure 5.7 is how the presence of MutS,
MutL, and Vsr confers increased repair of 06mG. One may suggest that MutS and MutL
assist MTase proteins in identification of 06mG lesions by presenting the lesion for repair
in the loop shaped structure reported by Allen (52). If this were the case, MutS and MutL
would stimulate (1) MMR by identifying mismatches on DNA and subsequently
activating the latent endonuclease activity of MutH at an unmethylated d(GATC) site
adjacent (by up to thousands of base pairs) to the mismatch (22), (2) VSPR by identifying
G:T mismatches on DNA and facilitating their recognition by Vsr for incision next to the
mismatched T (54), and (3) MTase repair by identifying 06mG mispairs and facilitating
their recognition by a MTase for direct reversal of the damage. In the first scenario, the
endonuclease activity of MutH is activated directly by MutS and MutL. In contrast, the
second two situations involve a mechanism whereby MutS and MutL assist the
efficiency, but not activity, of the downstream repair protein, and are consistent with the
observations that Vsr and the MTases are active in the absence of MutS and MutL. As an
alternative to presenting the lesion for recognition, MutS and MutL may assist the
efficiency of Vsr and/or the MTases directly through protein-protein interactions to aid
localization of these proteins at damaged DNA substrates.
One technology to study the extent to which proteins directly interact with each
other in vivo is the two-hybrid system (55). One such bacterial two-hybrid system
utilizes the reconstitution of the catalytic domain of Bordtella pertussis adenylate cyclase
(Figure 5.11) (56). In this system, the two proteins of interest are genetically fused to the
complementary T25 and T18 enzyme fragments that associate to produce cAMP. This
molecule then triggers transcriptional activation of genes such as lacZwhich are involved
in carbohydrate fermentation and produce a characteristic phenotype. In the absence of
binding affinity between the two proteins of interest the T25 and T18 fragments do not
associate for cAMP synthesis. However, as the binding affinity between the two proteins
203
- Chapter5 of interest increases, the amount of cAMP produced increases, as well. Therefore, this
system provides a tool to study protein-protein interactions qualitatively.
The Cupples laboratory has successfully utilized this system to study the extent
to which the proteins involved in VSPR interact with each other (57,58). Using j3galactosidase units as a measure, it was determined that MutL interacts with the Nterminus of Vsr.
It was also determined that the interaction between MutL and Vsr
inhibits the ability of MutL to dimerize, to interact with MutS and MutH, and to mediate
a previously unknown interaction between MutS and MutH.
In efforts to understand the extent to which these VSPR proteins interact with
the Ada and Ogt MTases, we have established a collaborative project with the Cupples
Lab. The T18 and T25 fusion vectors, each containing Ada and Ogt, will be constructed
and investigated as previously described (57;58) to address the possibility that these
proteins interact with MutS, MutL, and Vsr in vivo.
Toxicity Studies on Cells Proficient and Deficient in Vsr Protein
As previously described, 06mG is cytotoxic to E. coil in a manner that is
dependent on MR. A number of studies have been done to address the contribution of the
Dam, MutS, MutL, and MutH proteins in alleviating the lethality of the lesion. However
no studies have reported the effect of Vsr on cell survival, and one could imaging
multiple outcomes.
First, it could be that Vsr contributes to the lethality of 06mG:T
mismatches. Vsr may incise 5' to the mismatched T to initiate a round of VSPR that
would give rise to the original 06mG:T mismatch. Here, reiterative rounds of attempted
repair could continue indefinitely because VSPR does not become inactive like MMR
does once the DNA is fully Dam methylated. This futile cycling could in turn lead to
death of the cell through some unknown mechanism.
A second, and more likely,
scenario involves Vsr combating the cytotoxic properties of 06mG. Given Vsr increases
the efficiency of 06mG MTase repair, one may hypothesize the presence of Vsr would
204
- Chapter5 correlate with more efficient repair of the lethal 06mG lesion. A third possibility is that
the presence or absence of Vsr has no effect on the toxicity of 06mG.
As described in the discussion section, once MMR is inactive, MTase repair of
06mG:T to G:T is only useful (in avoiding a mutation during the next round of
replication) if the subsequent G:T can be corrected to G:C by VSPR. Therefore, there is
a rational basis for why the MTase proteins might be more inclined to repair 06mG:T if
Vsr is present for a concerted effort to restore the normal G:C pairing.
205
- Chapter 5 Figure 5.1. Sequence of ogt and location of peR primers.
analysis are shaded in gray.
1
61
121
181
241
301
361
421
481
ATGCTGAGAT TACTTGAAGA AAAAATTGCC
GATGAGCAAT TTCGCCTGCG GGCGGTTGAA
CTGCTGGACA TCCATTATCG CAAAGAAGGC
GGTTTAAGCG ACAAGCTTCG TGAATATTTT
CCCACTGCTA CGGGGGGGAC GCCATTTCAG
CCCTGCGGGC AGGTAATGCA TTACGGCCAA
GCGCGTGCCG TTGGTGCGGC AAACGGATCG
CGGGTTATTG GCCGAAACGG CACCATGACC
[..GGTT~rr~cJ-(f.CMT~GGTTATCTTTTG
206
ACGCCACTGG
TGGGAAGAGT
TATGAGCGCA
GCCGGTAATC
CGCGAAGTCT
CTGGCTGAGC
AATCCCATCA
GGATATGCAG
CTGTAA
Primer sequences used for
G~C~CTGTG~GGTGATTTG.O
ACAGCGAACG CATGGTGCAG
TTTCTGCCAC CAATCCAGGC
TTAGCATTAT TGATACGCTT
GGAAAACACT ACGCACTATC
AATTGGGCCG TCCTGGCGCG
GCATCGTCGT ACCTTGCCAT
GCGGAGTTCA GCGAAAAGAG
.............
- Chapter 5Figure 5.2. Sequence of ada/alkB and location of peR primers. Primer sequences used
for analysis are shaded in gray. The last base in ada is the first base of alkB. The co dons
that overlap are underlined.
1
61
121
181
241
301
361
421
481
541
601
661
721
781
841
901
961
1021
1081
1141
1201
1261
1321
1381
1441
1501
1561
1621
1681
ATGAAAAAAG CCACATGCTT~AACTGACGAT«-<?AACGCTGGC AA TCTGTCTT
CCGAATGCCG ACGGCGAATT CGTTTTCGCC GTGCGTACCA CAGGCATCTT
TCTTGCCGCG CCAGACATGC TTTGCGGGAA AACGTCTCCT TCTACGCAAA
GCACTCGCCG CTGGCTTTCG CCCCTGCAAA CGTTGTCAGC CAGAAAAAGC
CAACATCGGT TGGATAAAAT CACCCACGCG TGTCGACTGC TGGAACAGGA
ACGCTGGAAG CCTTAGCCGA CCAGGTGGCG ATGAGTCCAT TTCATCTACA
AAAGCGACTA CCGGAATGAC GCCTAAAGCC TGGCAACAGG CCTGGCGCGC
CGCGAATCGC TGGCGAAAGG GGAGAGCGTG ACGACGTCTA TTCTTAACGC
GACAGCAGCA GTTACTATCG CAAAGCTGAC GAAACGCTGG GCATGACGGC
CGTCACGGTG GCGAAAATCT GGCGGTGCGT TACGCGCTGG CTGATTGTGA
TGCCTGGTGG CAGAAAGCGA GCGGGGGATT TGCGCGATAT TGCTGGGCGA
ACACTAATCA GCGAGTTGCA GCAGATGTTT CCCGCTGCCG ACAACGCGCC
ATGTTTCAGC AACATGTGCG TGAAGTGATC GCCAGCCTCA ATCAACGCGA
ACGTTACCGC TGGACATTCG CGGCACTGCT TTTCAGCAAC AAGTCTGGCA
ACGATACCTT GCGGTGAAAC CGTCAGTTAT CAGCAACTGG CTAACGCCAT
AAAGCGGTAC GGGCCGTTGC CAGCGCCTGT GCCGCCAACA AGCTGGCTAT
TGTCATCGGG TGGTCCGTGG TGATGGCACA CTTTCCGGTT ACCGCTGGGG
AAAGCGCAAC TGCTGCGCCG CGAAGCTGAA AATGAGGAGA GGTAATGTTG
CCGATGCTGA ACCGTGGCAA GAGCCACTGG CGGCTGGTGC GGTAATTTTA
CTTTTAACGC TGCGGAGCAA CTGATCCGCG ATATTAATGA CGTTGCCAGC
TTCGCCAGAT GGTCACCCCC GGGGGATATA CCATGTCGGT GGCGATGACC
ATCTGGGCTG GACGACCCAT CGGCAAGGTT ATCTCTATTC GCCCATTGAT
ATAAACCGTG GCCCGCCATG CCACAGAGTT TTCATAATTT ATGTCAACGT
CGGCGGGCTA TCCAGATTTC CAGCCAGATG CTTGTCTTAT CAACCGCTAC
CGAAACTGTC GCTGCATCAG GATAAAGACG AACCGGATCT GCGCGCGCCA
TTTCTCTGGG CTTACCCGCG ATTTTTCAAT TTGGCGGCCT GAAACGAAAT
AACGTTTGTT GTTGGAACAT GGCGATGTGG TGGTATGGGG CGGTGAATCG
ATCACGGTAT TCAACCGTTG AAAGCGGGGT TTCATCCACT CACCATCGAC
Act'TGACAT'I';ICCGTGtJfOOEUt-oo.TMAAAAG
AATAA
207
AGCCCGCGAC
TTGCCGTCCG
TGCCAGCGAG
CAATGCCCAG
AACGCCTGTA
TCGGTTGTTT
TCGCCGTTTG
CGGATTCCCC
TAAACAATTC
GCTGGGTCGT
TGATGACGCG
TGCCGATCTG
TACGCCGCTG
GGCACTGCGC
CGGCAAACCG
CATAATACCC
CGTGTCGCGT
GATCTGTTTG
CGGCGTTTTG
CAGTCGCCGT
AACTGTGGGC
CCGCAAACAA
GCGGCTACGG
GCTCCTGGCG
ATTGTTTCTG
GATCCGCTCA
CGGCTGTTTT
TGCCGCTACA
- Chapter 5 Figure 5.3. Sequence of mutH and location of peR primers. Primer sequences used for
analysis are shaded in gray. Primer sequences used in production of recombination
substrate are shaded in black.
1
61
121
181
241
301
361
421
481
541
601
661
==.......
_......
~ .............
GCCAGCGCAG
ATCCCTGTGG
ACGGGCAATA
CTGTGGATAC
CCGTTGCTGT
TTAATGGATA
TTAC
GGCGAACGGA
CTACTGGCCC
;",;;;w,;;;,,;;;,.,;;g
GGAGTAAACC
ATAGTCTTGG
GCGGGGTGAC
CGGTTGAAGG
GGAGCCCGAA
TGATTGTTCT
TTCTGACGCT
GTCATTTTCT
GCTCTCTCCT
TACATTGGGA
CTGG
TGAGCAAGAT
TCGTCCGCTG
CTGGGAAACC
CGAGCGCAGC
TGAAGAGGAA
CGGTCAGGTT
GTTAGCGCAA
GCTGGTTACG
CTGGCTAGGT
ACTTAAAACT
TGCCCCGTTA
CAAACGCGTA
CGTTGGATCA
CTGGGAAGAA
CGGGGAGTAT
TGGTGCCCGG
GCCACGCGGC TTTTATTTGA AGAAGAATTT CACCAGTGCA
GATCCAGTAG
208
CCCGAAACTG
GAACTGGCGG
ATTGGCGTGT
TTTGCTGCTC
GAAACAACAT
AGCCACGTGC
ATCCCGCTGG
GACCGGCAGC
GAGCGGATTA
AAGAACAGTT
CACTTGTCGG
TACTGGAGAT
TGGGCGTGGA
TCGTTTGTGT
GCCACAAACT
CGCAGCGTCG
TACGCGAAGA
CCGCTCGTCA
- Chapter 5 Figure 5.4. Sequence of mutL and location of peR primers. Primer sequences used for
analysis are shaded in gray. Primer sequences used in production of recombination
substrate are shaded in black.
1
61
121
181
241
301
361
421
481
541
601
661
721
781
841
901
961
1021
1081
1141
1201
1261
1321
1381
1441
1501
1561
1621
1681
1741
1801
ATGCCAATTC~GGTCTTACC-GCGACAACTG
GAGCGACCTG CGTCGGTAGT CAAAGAACTA
GTGGG
TGCGGTATCA AAAAAGATGA GCTGGCGCTG
GCCTCTCTGG ACGATCTCGA AGCCATTATC
AGTATCAGTT CGGTTTCCCG CCTGACGCTC
TGGCAGGCCT ATGCCGAAGG GCGCGATATG
GTGGGGACGA CGCTGGAGGT GCTGGATCTG
CTGCGCACCG AGAAAACCGA ATTTAACCAC
GCGCGTTTCG ACGTCACGAT CAACCTGTCG
GCAGTGCCGG AAGGCGGGCA AAAAGAACGG
CTTGAACAAG CGCTGGCGAT TGAATGGCAA
GCCGATCCAA ATCACACCAC GCCCGCACTG
CGCATGATGC GCGATCGCCT GATCAATCAC
GGGGCCGATC AGCAACCGGC ATTTGTGTTG
GTCAACGTGC ACCCCGCCAA ACACGAAGTG
TTTATCTATC AGGGCGTGCT GAGCGTGCTA
GACGATGAAC CCCAACCTGC ACCGCGTTCC
AATCACTTTG CAGAACCGGC AGCTCGTGAG
GCATCAGGCA GTCGTCCGGC TGCCCCCTGG
CAAGGTGAAG TGTATCGCCA GCTTTTGCAA
CCGGAACCGC AGGAACCTGC ACTTGCGGCG
ATCGTCCATT CCGACTGTGC GTTGCTGGAG
CCAGTGGCAG AACGTTGGCT GCGTCAGGCA
GCCCAGCCGC TGCTGATTCC GTTGCGGCTA
GAAAAAGCGC AGTCTGCCCT GGCGGAATTG
GTGACCATCA GGGCAGTGCC TTTACCCTTA
GAACTGATAG GCTACCTGGC GAAGCAGTCC
ATTGCACGAA ATCTGAT G CGAACATGCG
CTGGCGGACG TGGAACGGTT ATGTCCGCAA
CAATCTGTTG ATTTACATCC GGGGATAAAA
209
GCGAACCAGA
GTGGAAAACA
GCGAAACTTA
GCGCTGGCTC
AGCCTGGGCT
ACTTCACGCA
AACGTGACGG
TTCTACAACA
ATTGATGAGA
CATAACGGTA
CGCTTAGGCG
CACGGCGATC
GCAGAAATTC
GCGATCCGCC
TATCTGGAGA
CGTTTCCATC
CAACAGCAAC
ATTCCGGAAA
CCGGTAGCTC
CCGAATGCGC
ACGCCCGCGC
AACAGTCAGA
CGCGACGGCA
CAATTGACGC
AAAGTTTCTG
GGTATTGATT
CGCCAACAAA
GTATTCGAAC
CAGTGGTCAA
CTTGTGAAAA
GGCCTGAAAG
TTGCCGCAGG
GCCTCGATGC
TCCGCATTCG
GTCATGCCAC
TTCGCGGTGA
CCGCAGAACA
TAAAACCGGC
CCCCGGCGCG
TCATCCGCCG
AAATTGTGCG
CGATTTGCGG
TCACGCTACG
AGTATTGCTA
AGGCCTGCGA
TCGACCCACA
AGTCGCGTCT
TGGAAACGCC
ACCGCGTGGC
CGCGCTACAC
AGCCAGGCTA
CGATGCAAAA
GTTTTGGTCG
ACATTTCACT
CGGGTGAAGC
CCGAAGAAAA
TCCAGTCAGA
ATTTACAAAT
CTGGCAATAT
TGGCACAGGC
CGCCGCCGGG
ATGAGTGA
TGAGGTGGTC
AGGTGCGACG
TGATAACGGC
CAGTAAAATC
GGCGCTGGCG
GCAGGAAGCC
GGCGCATCCT
GCGCAAATTC
CATTGCGCTG
TCAGTACCGC
CACCGCTTTT
CGGCTGGGTG
CGTGAACGGT
AGACAAACTG
TCAGGTGGAC
GGTGCATGAT
GCTACCGCTG
GGCGGGGCGC
TCCTGCGCCA
CCAGAAACAG
ATTAAAAGCG
GGTACTGACT
TTTATCCTTG
GCCCGTTTGC
ATCGGCATTA
TGCACAGCAT
CTTGATTCCT
TGCGCAGTGG
CATAACCCTG
TGGTCTGTTA
- Chapter 5 Figure 5.5. Sequence of mutS and location of peR primers. Primer sequences used for
analysis are shaded in gray. Primer sequences used in production of recombination
substrate are shaded in black.
1
61
121
181
241
301
361
421
481
541
601
661
721
781
841
901
961
1021
1081
1141
1201
1261
1321
1381
1441
1501
1561
1621
1681
1741
1801
1861
1921
1981
2041
2101
2161
2221
2281
2341
2401
2461
2521
ATGAGTGGAA~TAGAAAA~~CGACGCeGAT
AAAGCCCAGC ATCCCGAGAT CCTGCTGTTT
TA
TCGGCGGGAG AGCCGATCCC GATGGCGGGG
GCCAAACTGG TGAATCAGGG AGAGTCCGTT
ACCAGCAAAG GTCCGGTTGA GCGCAAAGTT
GATGAAGCCC TGTTGCAGGA GCGTCAGGAC
AAAGGTTTCG GCTACGCGAC GCTGGATATC
GCTGACCGCG AAACGATGGC GGCAGAACTG
GCAGAAGATT TTGCTGAAAT GTCGTTAATT
CTGTGGGAGT TTGAAATCGA CACCGCGCGC
GATCTGGTCG GTTTTGGCGT CGAGAACGCG
TTGCAGTATG CGAAAGATAC CCAACGTACG
GAACGTGAGC AGGACAGCAT CATTATGGAT
CAGAACCTGG CGGGTGGTGC GGAAAATACG
CCGATGGGCA GCCGTATGCT GAAACGCTGG
TTGCTTGAGC GCCAGCAAAC TATTGGCGCA
GTACTGCGTC AGGTCGGCGA CCTGGAACGT
CGCCCACGCG ATCTGGCCCG TATGCGCCAC
CAGTTAGAAA CTGTCGATAG TGCACCGGTA
GCCGAGCTGC GCGATCTGCT GGAGCGAGCA
GACGGTGGTG TTATCGCATC GGGCTATAAC
GACGGCGCGA CCGATTATCT GGAGCGTCTG
GACACGCTGA AAGTTGGCTT TAATGCGGTG
CAAAGCCATC TGGCACCCAT CAACTACATG
TACATCATTC CAGAGCTAAA AGAGTACGAA
CTGGCACTGG AAAAACAGCT TTATGAAGAG
GCGTTGCAAC AGAGCGCGAG CGCGCTGGCG
CGGGCCTATA CCCTGAACTA CACCTGCCCG
ACCGAAGGTC GCCATCCGGT AGTTGAACAA
CTGAATCTGT CGCCGCAGCG CCGCATGTTG
AGTACCTATA TGCGCCAGAC CGCACTGATT
CCGGCACAAA AAGTCGAGAT TGGACCTATC
GATGACCTGG CGTCCGGGCG CTCAACCTTT
TTACATAACG CCACCGAATA CAGTCTGGTG
ACCTACGATG GTCTGTCGCT GGCGTGGGCG
GCATTGACGT TATTTGCTAC CCACTATTTC
GGCGTCGCTA ACGTGCATCT CGATGCACTG
AGCGTGCAGG ATGGCGCGGC GAGCAAAAGC
GTGCCAAAAG AGGTTATTAA GCGCGCACGG
CCGAACG
GAAACTTCGC CTGCGGTCGA AGCTCTGGAA
CAGGCGCTGG AGTGGATTTA TCGCTTGAAG
210
ACGCCCATGA TGCAGCAGTA TCTCAGGCTG
TACCGGATGG GTGATTTTTA TGAACTGTTT
ACGCGGTGCT
ATTCCCTACC ATGCGGTGGA AAACTATCTC
GCCATCTGCG AACAAATTGG CGATCCGGCG
GTGCGTATCG TTACGCCAGG CACCATCAGC
AACCTGCTGG CGGCTATCTG GCAGGACAGC
AGTTCCGGGC GTTTTCGCCT GAGCGAACCG
CAACGCACTA ATCCTGCGGA ACTGCTGTAT
GAAGGCCGTC GCGGCCTGCG CCGTCGCCCG
CAGCAGTTGA ATCTGCAATT TGGGACCCGC
CCGCGCGGAC TTTGTGCTGC CGGTTGTCTG
ACTCTGCCGC ATATTCGTTC CATCACCATG
GCCGCGACGC GTCGTAATCT GGAAATCACC
CTGGCTTCTG TGCTCGACTG CACCGTCACG
CTGCATATGC CAGTGCGCGA TACCCGCGTG
TTGCAGGATT TCACCGCCGG GCTACAGCCG
ATTCTGGCAC GTCTGGCTTT ACGAACTGCT
GCTTTCCAGC AACTGCCGGA GCTGCGTGCG
CAGGCGCTAC GTGAGAAGAT GGGCGAGTTT
ATCATCGACA CACCGCCGGT GCTGGTACGC
GAAGAGCTGG ATGAGTGGCG CGCGCTGGCT
GAAGTCCGCG AGCGTGAACG TACCGGCCTG
CACGGCTACT ACATTCAAAT CAGCCGTGGG
CGTCGCCAGA CGCTGAAAAA CGCCGAGCGC
GATAAAGTTC TCACCTCAAA AGGCAAAGCA
CTGTTCGACC TGCTGTTGCC GCATCTGGAA
GAACTCGACG TGCTGGTTAA CCTGGCGGAA
ACCTTCATTG ATAAACCGGG CATTCGCATT
GTACTGAATG AGCCATTTAT CGCCAACCCG
ATCATCACCG GTCCGAACAT GGGCGGTAAA
GCGCTGATGG CCTACATCGG CAGCTATGTA
GATCGCATCT TTACCCGCGT AGGCGCGGCA
ATGGTGGAGA TGACTGAAAC CGCCAATATT
TTAATGGATG AGATCGGGCG TGGAACGTCC
TGCGCGGAAA ATCTGGCGAA TAAGATTAAG
GAGCTGACCC AGTTACCGGA GAAAATGGAA
GAGCACGGCG ACACCATTGC CTTTATGCAC
TACGGCCTGG CGGTTGCAGC TCTGGCAGGC
CAAAAGCTGC GTGAGCTGGA AAGCATTTCG
GAA
AATCTTGATC CGGATTCACT CACCCCGCGT
AGCCTGGTG~ AA
- Chapter 5Figure 5.6. Sequence of vsr and location of peR primers. Primer sequences used for
analysis are shaded in gray. Primer sequences used in production of recombination
substrate are shaded in black.
1
61
121
181
241
301
361
421
ATGGCC~CG;TTCACGA~AA
GATACGGCGA TAGAGAAGCG
GTTCAGGACG CCAGTCTGCC
ATATTTACCC ATGGCTGCTT
ACTCGAACCG AGTTCTGGCT
ATCAGTCGCT TGCAGGAACT
GGCCACTCGC
CCTCGCCAGT
CGGACGTCCG
CTGGCATCAT
GGAGAAGATA
CGGCTGGCGC
AGCAAAAATA
CTGTTAACCG
GATTTTGTCG
CATCACTGCT
GGTAAAAATG
GTATTGATTG
ACCGAGCGTC
GAAGGTGCCA GCGCGCAGAT CGACACGCAG GGGATTCATT
211
TGCGCGCGAT
GGCAGGGCCT
TTGATGAATA
ATCTGTTTAA
TTGAGCGCGA
TCTGGGAGTG
TGGAAGAGTG
TACTCGCTTG
TGCCACGCGT
GGCATTTCGC
TCGCTGCGTG
AGTGCCTGCG
TCGCCGCGAT
.tiWHW"~WH~
GATCTGCGGC
~
- Chapter 5 Figure 5.7. Relative efficiencies ofMTase repair in MR defective strains
100
_'\Nt"
I
I Vsr-
90
_MutH_MutL_MutS-
80
70
60
50
40
30
20
10
o
C215
C216
C218
C217
Parental Cell Type
C215
C216
C217
C218
wt
vsr
mutH
mutl
mutS
% Repair
wi
96
69
95
6
:t
Parent
C215
96
81
97
67
68
%of
parental
100
84
101
70
71
1
3
1
1
% Repair
vsr81
41
70
7
:t
% Repair
:t
3
2
4
1
wt
vsr
mutH
mutL
mutS
97
70
96
6
1
2
2
1
Parent
C216
69
41
70
52
43
% of
rJ parental
100
59
101
75
62
212
% Repair
:t
mutL-
mutH-
67
52
63
44
% Repair
:t
mulS-
1
3
3
4
wt
vsr
mutH
mutl
mutS
68
43
39
6
1
4
4
1
Parent
C217
95
70
96
63
39
%of
of parental
100
74
101
66
41
- Chapter 5 Figure 5.8. peR mediated gene replacement
Sequencematching5'
sideoftargetgP--
I
Resistancemarker-----0:quence
matching3'
sideoftargetgene
peR
from plasmid or
genomic DNA
Resistance marker
Electroporate
linear DNA
Resistance marker
E. coliKM22
'"
hyperrecombinogenic:
recBCD replaced with A, red
l
BMC Mol BioI. 2003 Dee 13;4(1):11.
Gene. 2000 Apr 4;246(1-2):321-30.
C
Resistance marker
vsr
213
/
vsr
J
- Chapter 5 -
Figure 5.9. In vivo assay for MTase repair
Not
Repaired
Repaired
STOP
UAG
-
UGG
= Trp
read
through
Active
~-Gal
X-Gal
from
the cell
DARK
~-Gal
CD-fragment
BLUE
214
LIGHT
BLUE
- Chapter 5 Figure 5.10. Hypothetical mechanism for stimulation of MTase repair by VSPR
proteins. (A) MutS and MutL identify the dmG mismatch and (B) present it in a looped
structure. (C) MutL stimulates the affinity of Vsr for the DNA. In the absence of a
substrate for Vsr, the protein assists in localizing the MTase to the dmG lesion site.
MutL
DNA
A
ctmG:T Mismatch
ADP
B
c
vsr~
•
215
MTase
- Chapter5 Figure 5.11. Bacterial two-hybrid system utilizing domains of adenylate cyclase. (A)
The fused T18 and T25 fragments of Bordetella pertussis adenylate cyclase exhibit a
constitutive enzymatic activity that results in cAMP synthesis. (B) However, when the
two fragments are co-expressed as independent polypeptides are unable to interact and
cAMP synthesis does not occur. (C) When the two fragments are co-expressed as fusion
proteins with X and Y, cAMP synthesis results from binding between X and Y.
Adenylate Cyclase
A
-c
N-
ATP
B
cAMP
N-
-C
No cAMP Produced
C
C
N
ATP
cAMP
216
- Chapter5 Table 5.1. PCR primers used for construction of recombination substrates.
vsr
5'- CCT CGC CAG TCT GTT AAC CGG GCA GGG CCT GGC ATT TCG CCT CGA
CAT CTT GGT TAC CGT GAA G -3'
5'- AAG CGC TTC ATC CGT CAG CTT CTC GCG CC ACG TAA CGC GCG CGG
AAT AAC ATC ATT TGG TGA C -3'
mutS
5'- GAC GAC GCA AAA CGC GCG TCG CAA CTG CTG GAT ATT TCA CTG ACC
AAC TCG ACA TCT TGG TTA CCG TGA AG -3'
5'- GGT ACT GAC AGC AAA GAC ATT TGC GTA CCA TCC ACT TGC GTA GCG
GCG CGC GGA ATA ACA TCA TTT GGT GAC -3'
mutL
5'- GCC TCG ATG CAG GTG CGA CGC GTA TCG ATA TTG ATA TCG AAC GCC
TCG ACA TCT TGG TTA CCG TGA AG -3'
5'- CAG GGT TAT GGC CTG TGC CAT TGA CCA CTG CGC ATG TTC GCT CGC
GGA ATA ACA TCA TTT GGT GAC -3'
mutH
5' - CAC TTG TCG GGC TGG TTA CGC CAG AGA ATT TAA AAC GCG ATA AAG
CTC GAC ATC TTG GTT ACC GTG AAG - 3'
5' - ATG GCT TCG GTA AGC GCT TTC GCA TTC GCT GCT TTC GGT CGT ATC
CGC GGA ATA ACA TCA TTT GGT GAC - 3'
217
- Chapter5 Table 5.2. PCR primers used for analyzing strain genotypes.
ada/alkB
5'- TGC TTA ACT GAC GAT CAA CGC TGG -3'
5'- TTA CCT GCC TGA CGG AAT GTC AG -3'
mutH
5'- ATG TCC CAA CCT CGC CCA CT -3'
5'- ACT GGC CCG TCA TTT TCT GAT CCA GTA G -3'
mutL
5'- ATG CCA ATT CAG GTC TTA CCG CC -3'
5'- CAT CTT TCA GGG CTT TTA TCG CCG G -3'
mutS
5'- GAG TGC AAT AGA AAA TTT CGA CGC CC -3'
5'- ACA CCA GGC TCT TCA AGC GAT AAA TC -3'
ogt
5'- TCC ACT GTG GGT GAT TTG C
5'- TCA TGG CGC AAT AAC CAC TC -3'
vsr
5'- TTT GAA TTC ATG ATG GCC GAC GTT CAC GAT -3'
5'- GAC AAG CTT TCA AGC GAG TAA ATG AAT CCC -3'
218
- Chapter5 Table 5.3. Sizes of gene and PCR amplification products.
Gene Size
PCR Product Sizes
KO Product Size
ada
mutH
1065
690
-1700
-700
-2000
-2200
mutL
mutS
ogt
vsr
1848
2562
516
471
-1850
-2550
-450
-450
-2200
-2200
-2100
-2200
* - Note that the expected size of the PCR product may not match the size of the gene if
the primer sequences used do not match the 3' and 5' most ends of the target gene. See
Figure 5.1 to 5.6 for locations of the sequences used for amplification.
219
- Chapter5 Table 5.4. Results from MTase assay on C215 derivatives.
Cell
Type
Volume
Plated
Average
Standard
Deviation
95
96
96
97
96
1
82
78
84
78
81
3
96
98
96
97
97
1
67
1
68
1
Dilution
Factor
Light
Blue
Dark
Blue
Events
Plated
Total
Events
%
Repaired
C215-1
200
50
19
395
414
20700
C215-2
C215-3
C215-4
200
200
50
50
25
15
568
376
593
391
29650
19550
200
50
16
549
565
28250
C215V-1
C215V-2
C215V-3
C215V-4
75
75
75
133
133
133
64
82
76
295
297
390
359
379
466
47867
50533
62133
75
133
90
322
412
54933
C215H-1
C215H-2
200
200
50
50
25
10
656
411
681
421
34050
21050
C215H-3
C215H-4
200
50
23
602
75
133
9
306
625
315
31250
42000
C215L-1
C215L-2
C215L-3
C215L-4
75
75
75
133
133
133
133
145
146
288
285
310
421
430
456
56133
57333
60800
75
133
157
319
476
63467
68
66
68
67
C215S-1
C215S-2
C215S-3
C215S-4
200
50
75
133
350
314
409
525
455
602
26250
60667
30100
67
69
68
281
411
54800
68
200
50
175
141
193
75
133
130
220
- Chapter5 Table 5.5. Results from MTase assay on C216 derivatives.
Cell
Type
Volume
Plated
Dilution
Factor
Light
Blue
Dark
Blue
Events
Plated
Total
Events
%
Repaired
C216-1
C216-2
C216-3
C216-4
200
200
200
200
50
50
50
50
137
144
152
161
346
373
315
320
483
517
467
481
24150
25850
23350
24050
C216V-1
C216V-2
C216V-3
C216V-4
200
200
200
200
50
50
50
50
168
319
439
372
107
240
274
272
275
559
713
644
C216H-1
C216H-2
C216H-3
C216H-4
200
200
200
200
50
50
50
50
48
102
133
131
111
226
317
343
C216L-1
C216L-2
C216L-3
C216L-4
200
75
75
75
50
133
133
133
283
302
217
185
C216S-1
C216S-2
C216S-3
C216S-4
200
200
200
200
50
50
50
50
139
145
228
169
Average
Standard
Deviation
72
72
67
67
69
3
13750
27950
35650
32200
39
43
38
42
41
2
159
328
450
474
7950
16400
22500
23700
70
69
70
72
70
1
255
375
251
206
538
677
468
391
26900
90267
62400
52133
47
55
54
53
52
3
82
127
189
129
221
272
417
298
11050
13600
20850
14900
37
47
45
43
43
4
221
- Chapter5 Table 5.6. Results from MTase assay on C217 derivatives.
Cell
Type
Volume
Plated
Dilution
Factor
Light
Blue
Dark
Blue
Events
Plated
Total
Events
%
Repaired
C217-1
200
200
50
50
19
25
326
473
345
498
17250
24900
94
200
50
15
338
353
17650
200
50
21
345
366
18300
96
94
C217V-1
C217V-2
C217V-3
C217V-4
200
200
200
200
50
50
50
50
176
165
258
269
472
423
545
510
648
588
803
779
32400
29400
40150
38950
73
72
C217H-1
200
200
200
200
50
50
50
50
7
9
6
17
382
181
184
316
389
190
190
333
19450
9500
9500
16650
98
C217L-1
C217L-2
C217L-3
C217L-4
200
200
50
50
106
67
178
113
284
180
14200
9000
200
200
50
50
88
202
128
386
216
588
10800
29400
63
63
59
C217S-1
C217S-2
C217S-3
C217S-4
200
50
116
58
174
8700
33
200
50
234
178
412
20600
200
50
144
90
234
11700
200
50
364
259
623
31150
43
38
42
C217-2
C217-3
C217-4
C217H-2
C217H-3
C217H-4
222
Average
Standard
Deviation
95
1
70
3
96
2
63
3
39
4
95
68
65
95
97
95
66
- Chapter5 Table 5.7. Results from MTase assay on C218 derivatives.
Average
Standard
Deviation
7
7
5
6
6
1
24500
23600
27050
19850
6
9
6
7
7
1
278
231
343
462
13900
11550
17150
23100
7
6
7
6
6
1
66
47
69
91
132
105
161
227
6600
5250
8050
11350
50
45
43
40
44
4
9
32
38
22
158
507
563
520
7900
25350
28150
26000
6
6
7
4
6
1
Cell
Type
Volume
Plated
Dilution
Factor
Light
Blue
Dark
Blue
Events
Plated
Total
Events
%
Repaired
C218-1
200
200
200
200
50
50
50
50
418
474
234
407
31
37
12
27
449
511
246
434
22450
25550
12300
21700
C218V-1
C218V-2
C218V-3
C218V-4
200
200
200
200
50
50
50
50
460
430
506
369
30
42
35
28
490
472
541
397
C218H-1
C218H-2
C218H-3
C218H-4
200
200
200
200
50
50
50
50
258
218
320
436
20
13
23
26
C218L-1
C218L-2
C218L-3
C218L-4
200
200
200
200
50
50
50
50
66
58
92
136
C218S-1
200
200
200
200
50
50
50
50
149
475
525
498
C218-2
C218-3
C218-4
C218S-2
C218S-3
C218S-4
223
- Chapter5 5.7. REFERENCES
(1) Loechler, E. L., Green, C. L., and Essigmann, J. M. (1984) In vivo mutagenesis
by 06-methylguanine built into a unique site in a viral genome. Proc. Natl. Acad.
Sci. U. S. A. 81(20), 6271-6275.
(2) Hill-Perkins, M., Jones, M. D., and Karran, P. (1986) Site-specific mutagenesis
in vivo by single methylated or deaminated purine bases. Mutat. Res. 162(2), 153163.
(3) Bhanot, 0. S., and Ray, A. (1986) The in vivo mutagenic frequency and
specificity of 06-methylguanine in phi X174 replicative form DNA. Proc. Natl.
Acad. Sci. U. S. A. 83(19), 7348-7352.
(4) Singer, B., and Dosanjh, M. K. (1990) Site-directed mutagenesis for quantitation
of base-base interactions at defined sites. Mutat. Res. 233(1-2), 45-51.
(5) Karran, P., and Marinus, M. G. (1982) Mismatch correction at 06-methylguanine
residues in E. coli DNA. Nature. 296(5860), 868-869.
(6) Takano, K., Nakamura, T., and Sekiguchi, M. (1991) Roles of two types of 06methylguanine-DNA methyltransferases in DNA repair. Mutat. Res. 254(1), 3744.
(7) Rebeck, G. W., and Samson, L. (1991) Increased spontaneous mutation and
alkylation sensitivity of Escherichia coli strains lacking the ogt 06-methylguanine
DNA repair methyltransferase. J. Bacteriol. 173(6), 2068-2076.
(8) Delaney, J. C., and Essigmann, J. M. (2001) Effect of sequence context on 0(6)methylguanine repair and replication in vivo. Biochemistry. 40(49), 14968-14975.
(9) Calmann, M. A., Evans, J. E., and Marinus, M. G. (2005) MutS inhibits RecAmediated strand transfer with methylated DNA substrates. Nucleic Acids Res.
33(11), 3591-3597.
(10) Rasmussen, L. J., and Samson, L. (1996) The Escherichia coli MutS DNA
mismatch binding protein specifically binds 0(6)-methylguanine DNA lesions.
Carcinogenesis. 17(9), 2085-2088.
(11) Pauly, G. T., Hughes, S. H., and Moschel, R. C. (1994) Response of repaircompetent and repair-deficient Escherichia coli to three 06-substituted guanines
and involvement of methyl-directed mismatch repair in the processing of 06methylguanine residues. Biochemistry. 33(31), 9169-9177.
(12) Karran, P., and Bignami, M. (1994) DNA damage tolerance, mismatch repair and
genome instability. Bioessays. 16(11), 833-839.
224
Chapter5 (13) Shevell, D. E., bou-Zamzam, A. M., Demple, B., and Walker, G. C. (1988)
Construction of an Escherichia coli K-12 ada deletion by gene replacement in a
recD strain reveals a second methyltransferase that repairs alkylated DNA. J.
Bacteriol. 170(7), 3294-3296.
(14) Karran, P., Lindahl, T., and Griffin, B. (1979) Adaptive response to alkylating
agents involves alteration in situ of 06-methylguanine residues in DNA. Nature.
280(5717), 76-77.
(15) Potter, P. M., Wilkinson, M. C., Fitton, J., Carr, F. J., Brennand, J., Cooper, D. P.,
and Margison, G. P. (1987) Characterisation and nucleotide sequence of ogt, the
06-alkylguanine-DNA-alkyltransferase gene of E. coli. Nucleic Acids Res.
15(22), 9177-9193.
(16) Rebeck, G. W., Coons, S., Carroll, P., and Samson, L. (1988) A second DNA
methyltransferase repair enzyme in Escherichia coli. Proc. Natl. Acad. Sci. U. S.
A. 85(9), 3039-3043.
(17) Lindahl, T., Demple, B., and Robins, P. (1982) Suicide inactivation of the E. coli
06-methylguanine-DNA methyltransferase. EMBO J. 1(11), 1359-1363.
(18) Jeggo, P. (1979) Isolation and characterization of Escherichia coli K-12 mutants
unable to induce the adaptive response to simple alkylating agents. J. Bacteriol.
139(3), 783-791.
(19) Su, S. S., and Modrich, P. (1986) Escherichia coli mutS-encoded protein binds to
mismatched DNA base pairs. Proc. Natl. Acad. Sci. U. S. A. 83(14), 5057-5061.
(20) Lieb, M. (1983) Specific mismatch correction in bacteriophage lambda crosses
by very short patch repair. Mol. Gen. Genet. 191(1), 118-125.
(21) Meselson, M.
(1988) Methyl-directed repair of DNA mismatches. In
Recombination of the Genetic Material (Low, K. B., Ed.) pp 91-113, Academic
Press, Inc., San Diego, Calif.
(22) Au, K. G., Welsh, K., and Modrich, P. (1992) Initiation of methyl-directed
mismatch repair. J. Biol. Chem. 267(17), 12142-12148.
(23) Wagner, R., Jr., and Meselson, M. (1976) Repair tracts in mismatched DNA
heteroduplexes. Proc. Natl. Acad. Sci. U. S. A. 73(11), 4135-4139.
(24) Burdett, V., Baitinger, C., Viswanathan, M., Lovett, S. T., and Modrich, P.
(2001) In vivo requirement for RecJ, ExoVII, ExoI, and ExoX in methyl-directed
mismatch repair. Proc. Natl. Acad. Sci. U. S. A. 98(12), 6765-6770.
(25) Hennecke, F., Kolmar, H., Brundl, K., and Fritz, H. J. (1991) The vsr gene
product of E. coli K-12 is a strand- and sequence-specific DNA mismatch
endonuclease. Nature. 353(6346), 776-778.
225
- Chapter5 (26) Dzidic, S., and Radman, M.
(1989) Genetic requirements for hyperrecombination by very short patch mismatch repair: involvement of Escherichia
coli DNA polymerase I. Mol. Gen. Genet. 217(2-3), 254-256.
(27) Lieb, M., and Bhagwat, A. S. (1996) Very short patch repair: reducing the cost of
cytosine methylation. Mol. Microbiol. 20(3), 467-473.
(28) May, M. S., and Hattaman, S.
(1975) Deoxyribonucleic acid-cytosine
methylation by host- and plasmid-controlled enzymes. J. Bacteriol. 122(1), 129138.
(29) Lieb, M., and Rehmat, S. (1995) Very short patch repair of T:G mismatches in
vivo: importance of context and accessory proteins. J. Bacteriol. 177(3), 660-666.
(30) Glasner, W., Merkl, R., Schellenberger, V., and Fritz, H. J. (1995) Substrate
preferences of Vsr DNA mismatch endonuclease and their consequences for the
evolution of the Escherichia coli K-12 genome. J. Mol. Biol. 245(1), 1-7.
(31) Current Protocols in Molecular Biology(2001) John Wiley & Sons, Inc.,
Hoboken, N.J.
(32) Murphy, K. C., Campellone, K. G., and Poteete, A. R. (2000) PCR-mediated
gene replacement in Escherichia coli. Gene. 246(1-2), 321-330.
(33) Delaney, J. C., and Essigmann, J. M. (1999) Context-dependent mutagenesis by
DNA lesions. Chem. Biol. 6(10), 743-753.
(34) Henderson, P. T., Delaney, J. C., Gu, F., Tannenbaum, S. R., and Essigmann, J.
M. (2002) Oxidation of 7,8-dihydro-8-oxoguanine affords lesions that are potent
sources of replication errors in vivo. Biochemistry. 41(3), 914-921.
(35)
Henderson,
P. T., Delaney, J. C., Muller, J. G., Neeley, W. L., Tannenbaum,
S.
R., Burrows, C. J., and Essigmann, J. M. (2003) The hydantoin lesions formed
from oxidation of 7,8-dihydro-8-oxoguanine are potent sources of replication
errors in vivo. Biochemistry. 42(31), 9257-9262.
(36) Moriya, M. (1993) Single-stranded shuttle phagemid for mutagenesis studies in
mammalian cells: 8-oxoguanine in DNA induces targeted G.C-->T.A
transversions in simian kidney cells. Proc. Natl. Acad. Sci. U. S. A. 90(3), 11221126.
(37) Moriya, M., Zhang, W., Johnson, F., and Grollman, A. P. (1994) Mutagenic
potency of exocyclic DNA adducts: marked differences between Escherichia coli
and simian kidney cells. Proc. Natl. Acad. Sci. U. S. A. 91(25), 11899-11903.
(38)
Samson, L., Han, S., Marquis, J. C., and Rasmussen,
L. J. (1997) Mammalian
DNA repair methyltransferases shield 04MeT from nucleotide excision repair.
Carcinogenesis. 18(5), 919-924.
226
- Chapter5 (39) Mackay, W. J., Han, S., and Samson, L. D. (1994) DNA alkylation repair limits
spontaneous base substitution mutations in Escherichia coli. J. Bacteriol. 176(11),
3224-3230.
(40) Cupples, C. G., and Miller, J. H. (1989) A set of lacZ mutations in Escherichia
coli that allow rapid detection of each of the six base substitutions. Proc. Natl.
Acad. Sci. U. S. A. 86(14), 5345-5349.
(41) Vincze, T., Posfai, J., and Roberts, R. J. (2003) NEBcutter: A program to cleave
DNA with restriction enzymes. Nucleic Acids Res. 31(13), 3688-3691.
(42) Altschul, S. F., Gish, W., Miller, W., Myers, E. W., and Lipman, D. J. (1990)
Basic local alignment search tool. J. Mol. Biol. 215(3), 403-410.
(43) Rebeck, G. W., Smith, C. M., Goad, D. L., and Samson, L.
(1989)
Characterization of the major DNA repair methyltransferase activity in unadapted
Escherichia coli and identification of a similar activity in Salmonella
typhimurium. J. Bacteriol. 171(9), 4563-4568.
(44) Murphy, K. C. (1998) Use of bacteriophage lambda recombination functions to
promote gene replacement in Escherichia coli. J. Bacteriol. 180(8), 2063-2071.
(45) Sassanfar, M., Dosanjh, M. K., Essigmann, J. M., and Samson, L. (1991)
Relative efficiencies of the bacterial, yeast, and human DNA methyltransferases
for the repair of 06-methylguanine and 04-methylthymine. Suggestive evidence
for 04-methylthymine repair by eukaryotic methyltransferases. J. Biol. Chem.
266(5), 2767-2771.
(46) Boyle, J. M., Durrant, L. G., Wild, C. P., Saffhill, R., and Margison, G. P. (1987)
Genetic evidence for nucleotide excision repair of 06-alkylguanine in mammalian
cells. J. Cell Sci. Suppl. 6:147-60., 147-160.
(47) Bronstein, S. M., Skopek, T. R., and Swenberg, J. A. (1992) Efficient repair of
06-ethylguanine, but not 04-ethylthymine or 02-ethylthymine, is dependent
upon 06-alkylguanine-DNA alkyltransferase and nucleotide excision repair
activities in human cells. Cancer Res. 52(7), 2008-2011.
(48) Huang, J. C., Zamble, D. B., Reardon, J. T., Lippard, S. J., and Sancar, A. (1994)
HMG-domain proteins specifically inhibit the repair of the major DNA adduct of
the anticancer drug cisplatin by human excision nuclease. Proc. Natl. Acad. Sci.
U. S. A. 91(22), 10394-10398.
(49) Samson, L., Thomale, J., and Rajewsky, M. F. (1988) Alternative pathways for
the in vivo repair of 06-alkylguanine and 04-alkylthymine in Escherichia coli:
the adaptive response and nucleotide excision repair. EMBO J. 7(7), 2261-2267.
227
- Chapter5 (50) Voigt, J. M., Van, H. B., Sancar, A., and Topal, M. D. (1989) Repair of 06methylguanine by ABC excinuclease of Escherichia coli in vitro. J. Biol. Chem.
264(9), 5172-5176.
(51) Borowy-Borowski, H., and Chambers, R. W. (1987) A study of side reactions
occurring during synthesis of oligodeoxynucleotides containing 06alkyldeoxyguanosine residues at preselected sites. Biochemistry. 26(9), 24652471.
(52) Allen, D. J., Makhov, A., Grilley, M., Taylor, J., Thresher, R., Modrich, P., and
Griffith, J. D. (1997) MutS mediates heteroduplex loop formation by a
translocation mechanism. EMBO J. 16(14), 4467-4476.
(53) Schaaper, R. M., and Dunn, R. L. (1987) Spectra of spontaneous mutations in
Escherichia coli strains defective in mismatch correction: the nature of in vivo
DNA replication errors. Proc. Natl. Acad. Sci. U. S. A. 84(17), 6220-6224.
(54) Bhagwat, A. S., and Lieb, M. (2002) Cooperation and competition in mismatch
repair: very short-patch repair and methyl-directed mismatch repair in Escherichia
coli. Mol. Microbiol. 44(6), 1421-1428.
(55) Fields, S., and Song, 0. (1989) A novel genetic system to detect protein-protein
interactions. Nature. 340(6230), 245-246.
(56) Karimova, G., Pidoux, J., Ullmann, A., and Ladant, D. (1998) A bacterial twohybrid system based on a reconstituted signal transduction pathway. Proc. Natl.
Acad. Sci. U. S. A. 95(10), 5752-5756.
(57) Mansour, C. A., Doiron, K. M., and Cupples, C. G. (2001) Characterization of
functional interactions among the Escherichia coli mismatch repair proteins using
a bacterial two-hybrid assay. Mutat. Res. 485(4), 331-338.
(58)
Monastiriakos,
S. K., Doiron, K. M., Siponen, M. I., and Cupples, C. G. (2004)
Functional interactions between the MutL and Vsr proteins of Escherichia coli are
dependent on the N-terminus of Vsr. DNA Repair (Amst). 3(6), 639-647.
228
CHAPTER 6
Cloning,,Expression, and Purification of Truncated
Hormone Receptors
229
- Chapter6 6.1. INTRODUCTION
The final two projects are ones that are not related to mismatch repair. My skills
in protein purification were needed on another project involving the synthesis of agents
that kill cells by blocking DNA repair. My advisor asked me to include these chapters, as
they represent important contributions to the lab research portfolio.
6.2. NUCLEAR HORMONE RECEPTORS
Nuclear hormone receptors (NHR) are ligand inducible transcription factors that
regulate gene expression (1). Members of this family include the estrogen receptor (ER),
androgen receptor (AR), progesterone receptor (PR), and glucocorticoid receptor (GR)
proteins. In the absence of hormones, these receptors are associated with heat shock
proteins and are not active (1;2). However, upon binging to their cognate ligand, these
proteins undergo a series of events including conformational changes, dissociation from
heat shock complexes, dimerization, phosphorylation, nuclear translocation, and binding
to promoter sequences in DNA called hormone response elements (1-4). Promoter
binding is followed by the recruitment of coactivators, corepressors, and transcriptional
machinery for regulation of target genes (2).
The observation that a number of cancer types over express hormone receptors
makes these proteins good therapeutic targets (5-9). Indeed, many cancer treatments act
by interrupting hormone dependent transcription and cell proliferation, and a number of
these treatments are highly cancer specific because of the fidelity of ligand binding by
certain hormone receptors. Other cancer drugs produce their cytotoxic effects through
covalent damage to DNA (10;11). Our work aims to combine the specificity of steroid
binding with the toxicity of DNA damage to produce a potent drug that is selectively
lethal to cancer cells (Figure 6.1).
230
- Chapter6 In principle, tethering a synthetic ligand for a cancer specific receptor to a
chemical functionality that crosslinks DNA allows a bifunctional approach to slowing
cancer progression. First, the bifunctional compound may bind the cognate hormone
receptor and antagonize the receptors role in malignant growth. Second, the ability of the
compound to recruit a receptor to the site of DNA damage and render the damage
undetectable, allows time for interstrand crosslinks to form. We hypothesize that toxic
crosslinks will form preferentially on DNA in cancer cells due to the abundance of
camouflaging receptor proteins. In contrast, non-cancer cells containing relatively less
receptor protein will selectively detect and repair the monoadduct before formation of a
toxic crosslink.
The ability of such bifunctional agents to possess selective toxicity, on the basis
of the cell receptor status, has been demonstrated by our lab (12-15). The ability of lead
compounds, both in solution and bound to duplex DNA, to bind hormone receptors was
also investigated biochemically. This section describes my work towards these ends.
Four truncated human receptor genes were cloned and three receptor ligand
binding domains were expressed in S21 insect cells using a baculovirus. This chapter
contains the methods by which these genes were cloned and these proteins expressed and
purified. Chapters 7 and 8 contain the publications to which my work contributed.
231
- Chapter6 6.3. EXPERIMENTAL PROCEDURES
Preparation of the hER-LBD.
The human estrogen receptor ligand binding domain
(hER-LBD) was obtained by PCR amplification of a segment of the hER gene contained
within the plasmid pSV2NeoCMV-hER (16). Primers 5'-GGA TCC ATG TCT GCT
GGA GAC ATG AGA GCT GCC -3' and 5'-CCG CTC GAG TCA GAC TGT GGC
AGG GAA ACC CTC -3' were used to amplify a 0.94 kb segment containing amino
acids 282 to 595 of the hER. Using the BamH I and Xho I restriction sites contained
within these primers the amplified segment was inserted into complementary sites of
pSK278 (17) (provided by Sang Seok Koh, Whitehead Institute, Figure 6.2) using T4
DNA ligase.
Following transformation into competent E. coli DH5ac and restriction
analysis of DNA from isolated clones, the recombinant transfer vector (pSK278-hERLBD, Figure 6.3) and pBacPAC6 viral DNA (Clonetech, Palo Alto, CA) were
cotransfected into S21 cells (Invitrogen, Carlsbad, CA).
Recombinant virus was
collected, purified via plaque assay and used to reinfect S21 cells from which cell
extracts were prepared. Expression of the expected 34 kDa protein containing the hERLBD fused to a FLAG® peptide was confirmed by Western blot analysis. The hER-LBD
was purified from S21 cell lysates by immunoaffinity chromatography using ANTIFLAG® M2-affinity gel (Sigma). Following adsorption to the gel, the 34 kDa hER-LBD
protein was recovered by elution with FLAG® -peptide (Sigma) The estradiol binding
activity of the purified hER-LBD was confirmed using a competitive assay with [3 H]17p-estradiol (18).
232
- Chapter6 Preparation of the hAR-LBD.
The human androgen receptor ligand binding domain
(hAR-LBD) was obtained by PCR amplification of a segment of the hAR gene contained
within the plasmid pCMV3.1 AR (19). Primers 5'-AAT GCG GCC GCT ACT CTG
GGA GCC CGG AA -3' and 5'-GGG TTA ATT AAG GCT CAC TGG GTG TGG AAA
TAG AT -3' were used to amplify a 0.89 kb segment containing amino acids 624 to 919
of the hAR. Using the Not I and Pac I restriction sites contained within these primers the
amplified segment was inserted into complementary sites of pSK278 (17) (provided by
Sang Seok Koh, Whitehead Institute, Figure 6.2) using T4 DNA ligase.
Following
transformation into competent E. coli DH5c and restriction analysis of DNA from
isolated clones, the recombinant transfer vector (pSK278-hAR-LBD, Figure 6.4) and
pBacPAC6 viral DNA (Clonetech, Palo Alto, CA) were cotransfected into S21 cells
(Invitrogen, Carlsbad, CA). Recombinant virus was collected, purified via plaque assay
and used to reinfect Sf21 cells from which cell extracts were prepared. Expression of the
expected 33 kDa protein containing the hAR-LBD fused to a FLAG® peptide was
confirmed by Western blot analysis. The hAR-LBD was purified from Sf21 cell lysates
by immunoaffinity chromatography using ANTI-FLAG® M2-affinity gel (Sigma).
Following adsorption to the gel, the 33 kDa hAR-LBD protein was recovered by elution
with FLAG® -peptide (Sigma) The R1881 binding activity of the purified hAR-LBD was
confirmed using a competitive assay with [17a-methyl-3 H]-(R1881), analogous to that
for the estrogen receptor (18).
233
- Chapter6 Preparation of the hPR-LBD. The human progesterone receptor ligand binding domain
(hPR-LBD) was obtained by PCR amplification of a segment of the hPR gene contained
within the plasmid pCR®3.1 hPR-B (provided by Weigel Lab, Baylor College of
Medicine). Primers 5'- AAC GGA TCC ATG GAA GGG CAG CAC AAC TAC -3' and
5'-CCG CTC GAG GCC TCA CTT TTT ATG AAA GAG -3' were used to amplify a
1.0 kb segment containing amino acids 595 to 933 of the hPR. Using the BamH I and
Xho I restriction sites contained within these primers the amplified segment was inserted
into complementary sites of pSK278 (17) (provided by Sang Seok Koh, Whitehead
Institute, Figure 6.2) using T4 DNA ligase. Following transformation into competent E.
coli DH5a and restriction analysis of DNA from isolated clones, the recombinant transfer
vector (pSK278-hPR-LBD, Figure 6.5) and pBacPAC6 viral DNA (Clonetech, Palo Alto,
CA) were cotransfected into Sf21 cells (Invitrogen, Carlsbad, CA). Recombinant virus
was collected, purified via plaque assay and used to reinfect Sf21 cells from which cell
extracts were prepared. Expression of the expected 38 kDa protein containing the hPRLBD fused to a FLAG® peptide was confirmed by Western blot analysis. The hPR-LBD
was purified from Sf21 cell lysates by immunoaffinity chromatography using ANTIFLAG® M2-affinity gel (Sigma). Following adsorption to the gel, the 38 kDa hPR-LBD
protein was recovered by elution with FLAG® -peptide (Sigma)
The progesterone
binding activity of the purified hPR-LBD was confirmed using a competitive assay with
[3H]-progesterone, analogous to that for the estrogen receptor (18).
234
- Chapter6 Preparation of the hGR-LBD Transfer Vector. The human glucocorticoid receptor ligand
binding domain (hGR-LBD) was obtained by PCR amplification of a segment of the hGR
gene contained within the plasmid OB7 (20). Primers 5'- AAT GCG GCC GCT GAA
GCT CGA AAA ACA AA -3' and 5'- GGC TTA ATT AAC GCG TCA CTT TTG ATG
AAA C -3' were used to amplify a 0.87 kb segment containing amino acids 489 to 777 of
the hGR. Using the Not I and Pac I restriction sites contained within these primers the
amplified segment was inserted into complementary sites of pSK278 (17) (provided by
Sang Seok Koh, Whitehead Institute, Figure 6.2) using T4 DNA ligase. The recombinant
transfer vector (pSK278-hGR-LBD, Figure 6.6) was transformed into competent E. coli
DH5ct and restriction analyses were conducted to identify successful clones.
235
- Chapter 6-
Figure 6.1. Model for mechanism of bifunctional toxicants
I
Repair
Enzyme
I
I
I
Protein
Recognition
Domain
\
+' •
II
I
~I
\
\
~
I
I
I
I
I
I
I
I
I
I
I
I
I
I
~
Adduct is Repaired and
Rendered Non-Toxic
Complex is
Toxic to Cell
NON-CANCEROUS
CELL
CANCEROUS
CELL
236
- Chapter6 Figure 6.2. Restriction map of pSK278
Brr
Bpn
Pvul
Sca
SgrAI 2526
237
- Chapter6 Figure 6.3. Restriction map of pSK278-hER-LBD
Pvul 504
Sca 1493t
Eco01091
4
Agel 3196
Alel 3423
238
Chapter6 Figure 6.4. Restriction map of pSK278-hAR-LBD
Eco01091 4420
t'l
I
J'tUU
239
- Chapter6 Figure 6.5. Restriction map of pSK278-hPR-LBD
Rcm 77
2319
PspXI
Xho I
Sbfl
Pstl
Not I
Eag I
Pac I
2343
Alel 3498
M
240
i /'
- Chapter6 Figure 6.6. Restriction map of pSK278-hGR-LBD
Hl
IRI
51
Pvt
I
EcoO
'A/el 3389
· ,,
241
--
- Chapter6 -
6.4. REFERENCES
(1) McKenna,
N. J., Lanz, R. B., and O'Malley,
B. W.
(1999) Nuclear receptor
coregulators: cellular and molecular biology. Endocr. Rev. 20(3), 321-344.
(2) McKenna, N. J., and O'Malley, B. W. (2002) Combinatorial control of gene
expression by nuclear receptors and coregulators. Cell. 108(4), 465-474.
(3) Couse, J. F., and Korach, K. S. (1999) Estrogen receptor null mice: what have we
learned and where will they lead us? Endocr. Rev. 20(3), 358-417.
(4) Hall, J. M., and McDonnell, D. P. (1999) The estrogen receptor beta-isoform
(ERbeta) of the human estrogen receptor modulates ERalpha transcriptional
activity and is a key regulator of the cellular response to estrogens and
antiestrogens. Endocrinology. 140(12), 5566-5578.
(5) Ferno, M., Borg, A., Johansson,
U., Norgren,
A., Olsson, H., Ryden, S., and
Sellberg, G. (1990) Estrogen and progesterone receptor analyses in more than
4,000 human breast cancer samples. A study with special reference to age at
diagnosis and stability of analyses. Southern Swedish Breast Cancer Study Group.
Acta Oncol. 29(2), 129-135.
(6) Bonkhoff, H., Fixemer, T., Hunsicker, I., and Remberger, K. (2001) Progesterone
receptor expression in human prostate cancer: correlation with tumor progression.
Prostate. 48(4), 285-291.
(7) Brolin, J.,
Skoog,
L., and Ekman,
P.
(1992)
Immunohistochemistry
and
biochemistry in detection of androgen, progesterone, and estrogen receptors in
benign and malignant human prostatic tissue. Prostate. 20(4), 281-295.
(8) Latil, A., Bieche, I., Vidaud, D., Lidereau, R., Berthon, P., Cussenot, O., and
Vidaud, M. (2001) Evaluation of androgen, estrogen (ER alpha and ER beta), and
progesterone receptor expression in human prostate cancer by real-time
quantitative reverse transcription-polymerase chain reaction assays. Cancer Res.
61(5), 1919-1926.
(9) Netto, G. J., Cheek, J. H., Zachariah, N. Y., Romer, J. C., Lee, C. F., Schoenfeld,
R., and Chakmakjian, Z. H. (1990) Steroid receptors in benign mastectomy
tissue. Am. J. Clin. Pathol. 94(1), 14-17.
(10) Teicher, B. A. Antitumor alkylating agents. In CancerPrinciples and Practice
(1997) Lippincott-Raven: New York.
(11) Hurley, L. H. (2002) DNA and its associated processes as targets for cancer
therapy. Nat. Rev. Cancer. 2(3), 188-200.
242
- Chapter6 (12)
Rink, S. M., Yarema, K. J., Solomon, M. S., Paige, L. A., Tadayoni-Rebek,
B. M.,
Essigmann, J. M., and Croy, R. G. (1996) Synthesis and biological activity of
DNA damaging agents that form decoy binding sites for the estrogen receptor.
Proc. Natl. Acad. Sci. U. S. A. 93(26), 15063-15068.
(13)
Marquis, J. C., Hillier, S. M., Dinaut, A. N., Rodrigues, D., Mitra, K., Essigmann,
J. M., and Croy, R. G. (2005) Disruption of gene expression and induction of
apoptosis in prostate cancer cells by a DNA-damaging agent tethered to an
androgen receptor ligand. Chem. Biol. 12(7), 779-787.
(14)
Mitra, K., Marquis,
J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
(15) Sharma, U., Marquis, J. C., Nicole, D. A., Hillier, S. M., Fedeles, B., Rye, P. T.,
Essigmann, J. M., and Croy, R. G. (2004) Design, synthesis, and evaluation of
estradiol-linked genotoxicants as anti-cancer agents. Bioorg. Med. Chem. Lett.
14(14), 3829-3833.
(16) Green, S., Walter, P., Kumar, V., Krust, A., Bornert, J. M., Argos, P., and
Chambon, P. (1986) Human oestrogen receptor cDNA: sequence, expression and
homology to v-erb-A. Nature. 320(6058), 134-139.
(17) Koh, S. S., Hengartner,
C. J., and Young, R. A.
(1997) Baculoviral
transfer
vectors for expression of FLAG fusion proteins in insect cells. Biotechniques.
23(4), 622-627.
(18) Obourn, J. D., Koszewski, N. J., and Notides, A. C. (1993) Hormone- and DNAbinding mechanisms of the recombinant human estrogen receptor. Biochemistry.
32(24), 6229-6236.
(19) Tilley, W. D., Marcelli, M., Wilson, J. D., and McPhaul, M. J. (1989)
Characterization and expression of a cDNA encoding the human androgen
receptor. Proc. Natl. Acad. Sci. U. S. A. 86(1), 327-331.
(20) Hollenberg, S. M., Weinberger, C., Ong, E. S., Cerelli, G., Oro, A., Lebo, R.,
Thompson, E. B., Rosenfeld, M. G., and Evans, R. M. (1985) Primary structure
and expression of a functional human glucocorticoid receptor cDNA. Nature.
318(6047), 635-641.
243
CHAPTER7
A Rationally Designed Genotoxin that Selectively Destroys
Estrogen Receptor-Positive Breast Cancer Cells
Kaushik Mitra, John C. Marquis, Shawn M. Hillier, Peter T. Rye, Beatriz Zayas, Annie S.
Lee, John M. Essigmann, and Robert G. Croy
Department of Chemistryand Divisionof Bioengineeringand EnvironmentalHealth,
MassachusettsInstitute of Technology,Cambridge,Massachusetts02139
Received October 22, 2001
244
- Chapter77.1. ABSTRACT
We describe a novel strategy to increase the selective toxicity of genotoxic
compounds. The strategy involves the synthesis of bifunctional molecules capable of
forming DNA adducts that have high affinity for specific proteins in target cells. It is
proposed that the association of such proteins with damaged sites in DNA can
compromise protein function and/or DNA repair resulting in increased toxicity. We
describe the synthesis of a bifunctional compound consisting of an aniline mustard linked
to the 7a position of estradiol. This novel compound can form covalent DNA adducts
that have high affinity for the estrogen receptor. Breast cancer cells that express high
levels of the estrogen receptor showed increased sensitivity to the cytotoxic effects of the
new compound.
245
- Chapter 77.2 MANUSCRIPT
The persistence of genetic damage produced by alkylating agents (1-8) as well as
the antagonism of essential biochemical processes such as transcription can have lethal
consequences for malignant cells (9;10). Both mechanisms have been identified in
studies to uncover the reasons for the efficacy of cisplatin in the treatment of several
cancers (11-1g). We describe a synthetic strategy to create bifunctional molecules that
produce DNA adducts capable of binding the estrogen receptor (ER), which is aberrantly
expressed in many breast cancer cells (20). It is speculated that DNA adducts that form
complexes with the ER will be poorly repaired in these cells because they are
camouflaged from detection by DNA repair enzymes. Consequently, the DNA lesions
persist. Furthermore, the DNA adducts would be expected to act as "molecular decoys"
capable of displacing the ER from its natural targets and antagonizing its role in
malignant growth. In healthy cells, where the abundance of the ER is minimal, no such
ER-DNA adduct complexes will be present, and the cell should survive (21).
In this report we describe the design and synthesis of compound 1, a bifunctional
agent that can form covalent DNA adducts capable of binding the ER with high affinity
and specificity. We show that 1 has selective toxicity toward ER+ breast cancer cells
compared to ER- cells in vitro.
Compound 1 consists of a bis-chloroethyl aniline mustard as the DNA alkylating
unit tethered to estradiol, the natural ligand for the ER. The site of substitution of
estradiol in 1 was based on reports that relatively large alkyl groups can be attached at the
7a position with retention of high affinity for the ER (22-24). The synthetic strategy for
1 is shown in Scheme 7.1. Compound 7, a key compound in the synthesis, was prepared
by a modification of a published strategy (25;26). Briefly, 3 was functionalized with a 6carbon chain at the 7-position in a-stereochemistry to provide the alkenyl steroid 4.
Efficient reduction of the 6-oxo group in 4 was achieved with Et3SiH/BF 3.Et 2O; however,
this treatment also caused the loss of 3,17-tetrahydropyranoxy (THP) groups producing
diol 5. The 3,17-OHs of 5 were reprotected with THP groups to afford 6, followed by
oxidation of the alkene at the terminus of the linker to provide alcohol 7. Steroid alcohol
246
- Chapter 77 was converted to bromide 8, which was subsequently allowed to react with a protected
ethanolamine to give 9. Compound 9 was desilylated with tetrabutylammonium fluoride
(TBAF) and converted to a carbonate intermediate with p-nitrophenyl chloroformate.
The carbonate was coupled to (N,N-bis-2-chloroethylaminophenyl)propylamine that was
prepared from chlorambucil via the Curtius reaction(27). Deprotection of the product in
HCl/methanol furnished 1.
The affinity of 1 for the ER was first determined. Using a competitive binding
assay (28) with [3 H]-1713estradiol, compound 1 was found to have a relative binding
affinity (RBA) for the calf uterine ER of 30; RBA of estradiol = 100.
Next, the ability of 1 to modify DNA covalently was investigated. Plasmid DNA
was incubated with 100 pM [C]-I
(obtained by reacting the carbonate intermediate with
4-(N,N-bis-2-chloroethylaminophenyl)-[1-' 4 C] propylamine in step xii in Scheme 6.1;
additional details in Supporting Information) at 37 C for up to 6 h. After unbound 1 was
removed by phenol-CHCl3 extraction and ethanol precipitation, the radioactivity
associated with DNA was measured.
The amount of radioactivity bound to DNA
increased at a constant rate over the 6-h period indicating the formation of covalently
bound 1 (see Supporting Information).
On the basis of previous studies on DNA
alkylation by nitrogen mustards (29-31), it is likely that covalent adducts of 1 are formed
primarily at the N 7 atom of guanines. To investigate the identity of the covalent adducts,
1-modified DNA was treated with 0.1 N HCl, the major product from the digested DNA
was isolated by reversed-phase HPLC and analyzed by full scan electrospray mass
spectrometry to yield a prominent molecular ion signal at M + H+/z 813.5051. This mass
is consistent with a chemical structure in which one ethylene arm of the mustard of 1 is
attached to guanine and the other arm contains a -OH substituted for the Cl atom. DNA
modified with d4-1 (deuterium labeled 1) and analyzed using the same conditions for
hydrolysis, HPLC and mass spectrometry revealed a molecular ion signal at M+H+/z
817.5.
This result is consistent with d4 -1 attached to guanine as described.
preparation of d4-1 is described in Supporting Information.)
247
The
- Chapter7The affinity of DNA adducts of 1 for the ER was investigated using an
Electrophoretic Mobility Shift Assay (EMSA). Substrates for this assay were prepared
by
reaction
of
5' [32 p]-labeled
self-complementary
oligonucleotide
5'-
d(ATTATTGGCCAATAAT) with 1 for 4 h at 37 C. To assess quantitatively the level
of covalent modification under these conditions, the DNA was treated with piperidine (1
M, 90 C, 1 h) and the products were separated on a denaturing 20% polyacrylamide gel
(Figure 6.1A). This analysis found that 50% of the 16-mer oligo was cleaved, forming
two products identical with those produced by the Maxam-Gilbert sequencing reaction
for guanine (data not shown). EMSA was performed by combining the modified
[ 3 2 p]_
labeled 16-mer with a 30 kD fragment of the ER containing the ligand binding site for
estradiol (ER-LBD). The 30 kD ER-LBD was obtained by expression of a gene fragment
containing amino acids 282 to 595 of the human estrogen receptor in Sf21 cells using the
baculovirus expression system. See Supporting Information).
Analysis on a 4%
nondenaturing polyacrylamide gel (0.5X TBE) revealed that approximately 50% of the
16-mer formed a slowly migrating band (Figure 6.1B; lane 2). The amount of retarded
band corresponded closely with the level of modification of the 16-mer as revealed by
piperidine treatment. Furthermore, no slowly migrating band was seen with unmodified
16-mer (Figure 7.1B, lane 1) or with 16-mer modified with the nitrogen mustard
chlorambucil (see Supporting Information).
To examine the identity of the complex,
increasing amounts of 17f3-estradiol were added to the mixture containing the ER-LBD
and 1-modified 16-mer. As shown in Figure 7.1B (lanes 3-8), addition of 17f3-estradiol
resulted in disruption of the ER-LBD-DNA complex. These in vitro results indicated that
ER-LBD selectively forms a complex with 1-modified oligo, and raises the strong
possibility of formation of similar complexes in cells.
Clonal survival assays were performed in ER+ and ER- breast cancer cell lines to
determine if ER status affected the sensitivity of cells to the toxicity of 1. To reveal
clearly the effect of DNA adducts, cells were exposed to 1 for 2 h after which fresh drugfree medium was added to the cell cultures. This procedure minimized the possibility
that unreacted 1 could function as a receptor antagonist and thereby inhibit growth or
survival. Under these conditions 1 was found to possess a significantly lower EC5 0 in the
248
- Chapter7ER+ cell line MCF-7 (EC50= 3.5 pM) as compared to the ER- cell line MDA-MB231
(EC50 = 9.2 pM) (Figure 7.2). In contrast to these results, chlorambucil under the same
between these two cell lines (Figure 7.2).
conditions did not show any difference in EC50o
The fact that ER status affected the EC5 0 of 1, but not of chlorambucil, is
consistent with the role of the ER as an effector of selective toxicity of 1. Covalent 1guanine adducts have been identified in DNA isolated from treated cells (unpublished).
Therefore, 1 is stable to cell culture conditions and can form DNA adducts in cells
identical with those formed in vitro. We are currently investigating the role of 1-DNA
adducts in vivo to determine if they are less efficiently repaired in ER+ cells and to
evaluate whether the association of the ER with DNA adducts of 1 is directly responsible
for the greater sensitivity of MCF-7 cells.
Acknowledgment
We thank John Wishnok and W. Sara Gardinar-Stillwell for helpful discussions. We are
grateful to the National Cancer Institute (1-RO1-CA77743-04) for support of this work,
to the Susan G. Komen Foundation for a fellowship to K.M. (PDF00-000679), to the
National Institutes of Health for support of the NMR facility (1SlORR13886-01), and to
the BEH Mass Spectrometry Laboratory for use of their instruments.
Supporting Information Available
Procedures for the synthesis of 1, hER-LBD expression and purification, RBA
experiments, DNA-damaging studies, gel shift experiments and toxicity experiments
(PDF). This material is available free of charge via the Internet at http://pubs.acs.org.
249
- Chapter7Scheme 7.1.
Synthesis of compound 1.
a
a
Conditions: (i) DHP, pyridinium p-
toluenesulfonate, reflux, 92%; (ii) n-BuLi, KOtBu, B(OMe) 3, H 20 2; (iii) PCC, 59%; (iv)
6-iodohexene, KOtBu, Et3B, 47%; (v) CH 3 COCl/MeOH, Et 3SiH, BF 3.Et 2O, 74%; (vi)
77%; (vii) BH 3 .THF, KOH/H 20 2, 66%; (viii)
DHP, pyridinium p-toluenesulfonate,
methane sulfonyl chloride, LiBr, 86%; (ix) Ph 2 P(O)NH-CH 2-CH 2-OtBDMS, NaH,
catalyst tetra-n-butylammonium bromide,
DIEA;
(xii)
nitrophenylc hloroformate,
73%; (x) TBAF, 73%; (xi) p4-(N,N-bis-2-chloroethylaminophenyl)-
propylamine, DIEA; (xiii) H+, 60%.
OR
OR
iv
RO
C
RO
3 R=TH
3R=THP
4
0
4
IR
vii
S
6: R=THP
ix
N -,OTBDMS
8
I
HPh"
,N
1
250
*H
1
0
,O
N
9
¾ 1N-.ci
- Chapter 7Figure 1.1. (A) Piperidine treatment of self-complementary
deoxy oligonucleotide 5'd(AATATTGGCCAATATT)
treated with 1. Lane 1: untreated oligonucleotide, Lane 2:
oligonucleotide + 200 pM 1. (B) Retarded mobility of oligonucleotide-l in the presence
of ER-LBD illustrated by EMSA, and disappearance of the retarded band by competition
with estradiol. Lane 1: untreated oligonucleotide + ER-LBD, Lane 2: oligonucleotide
modified by 200 pM 1 + ER-LBD, Lanes 3-8: modified oligonucleotide + ER-LBD + 10300 nM estradiol.
B
A
12
Estradiol
12345678
-
.....
........- ._~ -.... -- -
251
..-
Chapter 7-
Figure 7.2. Survival of MCF-7 (ER+) (, 0) and MD-MB231 (ER-) (., o) cells treated
with 1 (closed symbols) or chlorambucil (open symbols).
100
1
I
Contan
252
of (1)
- Chapter 77.3. REFERENCES
(1) Dolan, E. M. (1997) Inhibition of DNA repair as a means of increasing the
antitumor activity of DNA reactive agents. Adv. Drug Del. Rev. 26, 105-118.
(2) aoui-Jamali, M., Loubaba, B. B., Robyn, S., Tapiero, H., and Batist, G.
(1994)
Effect of DNA-repair-enzyme modulators on cytotoxicity of L-phenylalanine
mustard and cis-diamminedichloroplatinum (II) in mammary carcinoma cells
resistant to alkylating drugs. Cancer Chemother. Pharmacol. 34(2), 153-158.
(3) Frankfurt, O. S., Seckinger, D., and Sugarbaker, E. V. (1993) Inhibition of DNA
repair in cells treated with a combination of alkylating agents. Anticancer Res.
13(4), 947-952.
(4) Frankfurt, O. S. (1991) Inhibition of DNA repair and the enhancement of
cytotoxicity of alkylating agents. Int. J. Cancer. 48(6), 916-923.
(5) Ross, W. E., Ewig, R. A., and Kohn, K. W.
(1978) Differences
between
melphalan and nitrogen mustard in the formation and removal of DNA cross-links.
Cancer Res. 38(6), 1502-1506.
(6) Hansson, J., Lewensohn, R., Ringborg, U., and Nilsson, B. (1987) Formation and
removal of DNA cross-links induced by melphalan and nitrogen mustard in
relation to drug-induced cytotoxicity in human melanoma cells. Cancer Res.
47(10), 2631-2637.
(7) De, S., I, McHugh, P. J., Clingen, P. H., and Hartley, J. A. (2000) Defining the
roles of nucleotide excision repair and recombination in the repair of DNA
interstrand cross-links in mammalian cells. Mol. Cell Biol. 20(21), 7980-7990.
(8) Hansson, J., Keyse, S. M., Lindahl, T., and Wood, R. D. (1991) DNA excision
repair in cell extracts from human cell lines exhibiting hypersensitivity to DNAdamaging agents. Cancer Res. 51(13), 3384-3390.
(9) Zhai, X., Beckmann, H., Jantzen, H. M., and Essigmann, J. M. (1998) Cisplatin-
DNA adducts inhibit ribosomal RNA synthesis by hijacking the transcription
factor human upstream binding factor. Biochemistry. 37(46), 16307-16315.
(10) Lerner, L. J., and Jordan, V. C. (1990) Development of antiestrogens and their
use in breast cancer: eighth Cain memorial award lecture. Cancer Res. 50(14),
4177-4189.
(11)
Zhai, X., Beckmann, H., Jantzen, H. M., and Essigmann, J. M. (1998) Cisplatin-
DNA adducts inhibit ribosomal RNA synthesis by hijacking the transcription
factor human upstream binding factor. Biochemistry. 37(46), 16307-16315.
(12) Chu, G. (1994) Cellular responses to cisplatin. The roles of DNA-binding
proteins and DNA repair. J. Biol. Chem. 269(2), 787-790.
253
- Chapter7(13) Toney, J. H., Donahue, B. A., Kellett, P. J., Bruhn, S. L., Essigmann, J. M., and
Lippard, S. J. (1989) Isolation of cDNAs encoding a human protein that binds
selectively
to
DNA
modified
by
the
anticancer
drug
cisdiamminedichloroplatinum(II). Proc. Natl. Acad. Sci. U. S. A. 86(21), 8328-8332.
(14) Brown, S. J., Kellett, P. J., and Lippard, S. J. (1993) Ixrl, a yeast protein that
binds to platinated DNA and confers sensitivity to cisplatin. Science. 261(5121),
603-605.
(15) Trimmer, E. E., Zamble, D. B., Lippard, S. J., and Essigmann, J. M.
(1998)
Human testis-determining factor SRY binds to the major DNA adduct of cisplatin
and a putative target sequence with comparable affinities. Biochemistry. 37(1),
352-362.
(16)
Huang, J. C., Zamble, D. B., Reardon, J. T., Lippard, S. J., and Sancar, A. (1994)
HMG-domain proteins specifically inhibit the repair of the major DNA adduct of
the anticancer drug cisplatin by human excision nuclease. Proc. Natl. Acad. Sci. U.
S. A. 91(22), 10394-10398.
(17) Jamieson, E. R., and Lippard, S. J. (1999) Structure, Recognition, and Processing
of Cisplatin-DNA Adducts. Chem. Rev. 99(9), 2467-2498.
(18) McA'Nulty, M. M., and Lippard, S. J. (1996) The HMG-domain protein Ixrl
blocks excision repair of cisplatin-DNA adducts in yeast. Mutat. Res. 362(1), 7586.
(19)
Szymkowski, D. E., Yarema, K., Essigmann, J. M., Lippard, S. J., and Wood, R.
D. (1992) An intrastrand d(GpG) platinum crosslink in duplex M13 DNA is
refractory to repair by human cell extracts. Proc. Natl. Acad. Sci. U. S. A. 89(22),
10772-10776.
(20) Ferno, M., Borg, A., Johansson, U., Norgren, A., Olsson, H., Ryden, S., and
Sellberg, G. (1990) Estrogen and progesterone receptor analyses in more than
4,000 human breast cancer samples. A study with special reference to age at
diagnosis and stability of analyses. Southern Swedish Breast Cancer Study Group.
Acta Oncol. 29(2), 129-135.
(21) Rink, S. M., Yarema, K. J., Solomon, M. S., Paige, L. A., Tadayoni-Rebek,
B. M.,
Essigmann, J. M., and Croy, R. G. (1996) Synthesis and biological activity of
DNA damaging agents that form decoy binding sites for the estrogen receptor.
Proc. Natl. Acad. Sci. U. S. A. 93(26), 15063-15068.
(22) Bucourt, R., Vignau, M., and Torelli, V. (1978) New biospecific adsorbents for
the purification of estradiol receptor. J. Biol. Chem. 253(22), 8221-8228.
(23) Bowler, J., Lilley, T. J., Pittam, J. D., and Wakeling, A. E. (1989) Novel steroidal
pure antiestrogens. Steroids. 54(1), 71-99.
254
- Chapter 7(24) DaSilva, J. N., and van Lier, J. E. (1990) Synthesis and structure-affinity of a
series of 7 alpha-undecylestradiol derivatives: a potential vector for therapy and
imaging of estrogen-receptor-positive cancers. J. Med. Chem. 33(1), 430-434.
(25) Skaddan, M. B., Wust, F. R., and Katzenellenbogen, J. A. (1999) Synthesis and
Binding Affinities of Novel Re-Containing 7alpha-Substituted Estradiol
Complexes: Models for Breast Cancer Imaging Agents. J. Org. Chem. 64(22),
8108-8121.
(26) Tedesco, R., Fiaschi, R., and Napolitano, E. (1995) 6-Oxoestradiols from
Estradiols: Exploiting Site Selective Metalation of Aralkyl Systems with
Superbases. Synthesis., 1493-1495.
(27) Valu, K. K., Gourdie, T. A., Boritzki, T. J., Gravatt, G. L., Baguley, B. C., Wilson,
W. R., Wakelin, L. P., Woodgate, P. D., and Denny, W. A. (1990) DNA-directed
alkylating agents. 3. Structure-activity relationships for acridine-linked aniline
mustards: consequences of varying the length of the linker chain. J. Med. Chem.
33(11), 3014-3019.
(28)
Korenman, S. G. (1970) Endocrinology87,
1119-1123.
(29) Kohn, K. W., Hartley, J. A., and Mattes, W. B. (1987) Mechanisms of DNA
sequence selective alkylation of guanine-N7 positions by nitrogen mustards.
Nucleic Acids Res. 15(24), 10531-10549.
(30) Povirk, L. F., and Shuker, D. E. (1994) DNA damage and mutagenesis induced
by nitrogen mustards. Mutat. Res. 318(3), 205-226.
(31)
Osborne, M. R., Wilman, D. E., and Lawley, P. D. (1995) Alkylation of DNA by
the nitrogen mustard bis(2-chloroethyl)methylamine. Chem. Res. Toxicol. 8(2),
316-320.
255
CHAPTER 8
Design, synthesis, and evaluation of estradiol-linked
genotoxicants as anti-cancer agents
U. Sharma, J. C. Marquis, A. Nicole Dinaut, S. M. Hillier, B. Fedeles, P. T. Rye, J. M.
Essigmann, and R. G. Croy
Departmentof Chemistryand BiologicalEngineeringDivision,MassachusettsInstitute
of Technology, Cambridge, MA 02139, USA
Received 27 February 2004; Revised 21 April 2004; accepted21 April 2004. Available
online2 June 2004.
256
- Chapter8 8.1. ABSTRACT
A series of bifunctional compounds was prepared consisting of 17P estradiol
linked to a DNA damaging N,N-bis-(2-chloroethyl)aniline. The objective of our studies
was to determine the characteristics of the linker that permitted both reaction with DNA
and binding of the resultant covalent adducts to the estrogen receptor.
Linker
characteristics were pivotal determinants underlying the ability of the compounds to kill
selectively breast cancer cells that express the estrogen receptor.
yI)
H
I
Linker
(1)
257
- Chapter8 -
8.2.
INTRODUCTION
Many anti-tumor drugs produce their cytotoxic effect through covalent damage to
DNA (1;2). Repair enzymes that remove such covalent lesions, restoring the integrity of
genetic information, limit the effectiveness of these drugs. Inhibition of DNA repair in
cancer cells is an attractive yet not significantly explored strategy to potentiate the
therapeutic effects of alkylating chemotherapeutic drugs (3;4). We have previously
described the synthesis and biological activity of novel bifunctional compounds that form
covalent DNA adducts with high affinity for the estrogen receptor (ER) (5;6). The
compounds were designed to test the hypothesis that adduct-ER complexes would be
concealed from DNA repair proteins and therefore be refractory to repair.
The
compounds would thus exhibit greater toxicity in cancer cells that over express the ER
protein.
The original series of compounds consisted of an N,N-bis-chloroethylaniline
connected to a 2-phenylindole (2PI) group by alkyl-amino-carbamate linkers of various
lengths
(7).
Investigations with these derivatives identified several molecular
characteristics of the linker that were important for the compound to react with DNA and
bind to the ER. A compound with greater affinity for the ER was obtained by replacing
the 2PI group with that of 17f3-estradiolconnected via the 7a position (8). Both the 2PI
and estradiol-based compounds showed increased toxicity toward breast cancer cells that
express the ER.
Evidence was presented that the differential toxicity of the DNA
damaging agents was not due to an anti-hormonal mechanism.
We investigated the molecular features of the linker that are essential for the
biological activities of the estradiol-linked compound 1 (Figure 8.1) by varying the
structure of the linker connecting the N,N-bis-(2-chloroethyl)aniline and estradiol moiety.
The roles of the amino and carbamate groups in the linker were of particular interest
because their substitution with different chemical groups might lead to a less complex
synthetic process and increase overall yields.
The modified linkers retained the six
carbon alkyl chain that appears, based on our earlier work, to be essential for the attached
ligand to fit into the estradiol binding site of the ER (9). Amino, amido, and guanidino
258
- Chapter8 groups were incorporated into the region connecting the hexanyl-substituted estradiol and
the aniline group.
We report here the synthesis and physical properties of these
molecules and their cytotoxic effects toward breast cancer cells.
259
- Chapter8 8.3. EXPERIMENTAL PROCEDURES
The synthesis of compound 1 was described in our previous report (10). The
syntheses utilized 3,17p-bis-(2-tetrahydropyranyloxy)-7a-(6-hydroxyhexan- 1-yl)-estra1,3,5(10)triene 2 as the starting compound (see box in Scheme 7.1); its preparation has
also been described (11). Construction of linkers proceeded by linear additions to 2 with
final addition of the N,N-bis-(2-chloroethyl)aniline moiety. Compound 5 was prepared
by conversion alcohol 2 to the bromide, which was allowed to react with a protected
ethanolamine providing 3 as described in Ref. (12). The Mitsunobo reaction (13) then
was used to couple 1,3-bis-(tert-butoxycarbonyl)-2-methyl-2-thiopseudourea
with 3.
Reaction of the resulting product 4 with excess (N,N-bis-2-chloroethylaminophenyl)propylamine followed by acid deprotection produced 5. Procedures described by Linney
et al.(14) were applied to incorporate an N,N-disubstituted guanidine moiety into the
linker. The preparation of 7 proceeded with the initial reaction of 2 with 1,3-bis(tertbutoxycarbonyl)-2-methyl-2-thiopseudourea
under Mitsunobu conditions (15).
The
resulting product 6 was then allowed to react with an excess of (N,N-bis-2chloroethylaminophenyl)-propylamine
followed by acid deprotection to furnish 7.
Compound 9 was prepared by conversion of 2 to the p-nitrophenyl carbonate 8, which
was then allowed to react with (N,N-bis-2-chloroethylaminophenyl)-propylamine (16).
Removal of THP groups under acidic conditions produced 9. Compound 10, the starting
material for compounds 12-15, was prepared by hydrazinolysis of the phthalimide of 2
that was formed under Mitsunobu conditions (17). Compound 12 containing two amide
groups was synthesized by first reacting 10 with the NHS ester of 4(tertbutoxycarbonylamino)butyric acid. Following removal of the THP and Boc groups the
terminal amino group was allowed to react with the NHS ester of chlorambucil producing
12. Compound 13 in which the linker contains two amino groups was produced by
reduction of 12 with borane dimethylsulfide complex (18). The preparation of 14 was
also accomplished by conversion of 2 to the phthalimide via Mitsunobo conditions with
subsequent hydrazinolysis to obtain the amine 10. The NHS ester of chlorambucil was
then allowed to react with the terminal amine, producing 14. Compound 15, containing a
secondary amino group in the linker, was prepared by reduction of the amide in 14 using
borane dimethylsulfide complex (18).
260
- Chapter8 8.4. RESULTS AND DISCUSSION
In the initial characterization of the biochemical properties of new compounds 5,
7, 9, 12-15 we evaluated their affinities for the ER. A radiometric competitive binding
assay (19) with the rabbit uterine ER was used to determine the relative binding affinity
(RBA) of each compound for the ER as compared with estradiol; RBA=100. All of the
compounds exhibited some affinity for the ER. The data in Table 7.1 show that the new
compounds have RBA values for the ER ranging from 6 to 40.
Among the new
compounds, 15 containing a single amino group in the linker had an RBA of 40, which is
comparable to 1.
Although it is apparent that the original combination of the positively charged
secondary amine with the neutral carbamyl group (compound 1) results in a bifunctional
compound with excellent affinity for the rabbit uterine ER, compounds 7, 12, and 13 also
have good affinities. These molecules were viewed as valuable assets as we move ahead
toward probing structure-activity
relationships and the biochemical
mechanisms
underlying the biological activity of 1. It is likely that our 7a-linked estradiol compounds
adopt a binding mode similar to that identified for the 7a-undecylamide estradiol analog
ICI 164,384 (20). The positioning and orientation of the estradiol moiety of ICI-164,384
within the hydrophobic binding cavity of the ER is directed by its 7a side chain, which
protrudes out of a hydrophobic channel extending from the binding pocket.
surface of the LBD, a
90 °
At the
flexion of the undecyl chain enables the remainder of the linker
to track closely with the surface contours of the LBD (21). The low RBAs of compounds
5, 9, and 14 may result from surface interactions adopted by the linkers in these
molecules that do not permit optimal alignment of the estradiol moiety within the binding
cavity.
The reactivity of each compound with DNA was assessed by its ability to produce
piperidine
labile
sites
in
the
self
complementary
deoxyoligonucleotide
5-
d(AATATTGGCCAATATT). The results in Table 8.1 (column 3) indicate the percent of
the oligomer that was cleaved by piperidine.
Compound 9 in which the alkyl linker
contains a single carbamyl group produced the lowest level of modification (i.e., 3%
261
- Chapter8 cleaved by piperidine). Compound 14 containing an amido instead of the carbamyl group
produced approximately five times the number of DNA adducts (14% cleaved by
piperidine). High levels of reactivity toward DNA were observed with compounds with
linkers containing secondary amino groups. The combination of the amino and carbamyl
groups in the linker of 1 resulted in a 10-fold increase in reactivity over 9 in which the
linker contains only the carbamyl group. The reactivity of 1 was similar to that of 15 in
which the linker contains a single secondary amine suggesting that the charged amino
group is the major determinant of reaction rate.
Compound 13 in which the linker
contains a diamine -NH-(CH 2)4 -NH-CH 2 - was the most reactive (79% cleaved by
piperidine). The same is likely the case for molecules 5 and 7 in which the strongly basic
guanidino groups would be cationic under assay conditions. It is likely that the cationic
nature of these molecules gives them a high reactivity with DNA by localizing the
reactive alkylating group in the vicinity of nucleophilic atoms. A similar result has been
reported for a conjugate of chlorambucil with the polyamine spermidine (22).
Using an electrophoretic gel mobility shift assay (23), we observed that covalent
DNA adducts of 1, 5, 7, 13, and 15 form complexes with the portion of the ER containing
the ligand binding domain (ER-LBD) (Table 1, column 4).
Under conditions that
allowed complex formation, addition of the ER to the modified DNAs resulted in the
appearance of a slowly migrating band by electrophoresis that was eliminated by addition
of excess competitor, estradiol (data not shown). The results in Table 1 (column 4)
indicate that the extent of complex formation for 1, 7, 13, and 15 were correlated with the
RBAs of the unreacted compounds. The exception was compound 5 in which the linker
contained both amino and guanidino groups. In this case, despite its low RBA, virtually
all of the modified oligonucleotide formed a slowly migrating band. We do not know the
basis for this unexpected finding.
Log P and logD values can be predictive of aqueous solubility, absorption, and
permeability (24). The lipophilicities of 1, 5, 7, 9, 12-15 were assessed using an HPLC
method to estimate the logP of the neutral form of each compound. Log D values at
pH 7.4 were estimated using an equation derived by Horvith et al.(25) for basic
compounds. The logD values in Table 1 indicated that the aqueous solubilities of the
262
- Chapter8 eight compounds span approximately a 2500-fold range under physiological conditions.
The compounds with logD values >5 (compounds 9 and 14) had both low affinities for
the ER and low reactivity with DNA.
Compounds containing charged groups with
calculated logD values <3 generally had the highest affinities for the ER along with the
greatest reactivities toward DNA. These relationships, however, did not prove to be
reliable predictors of biological activities in cytotoxicity assays against breast cancer
cells.
The lethal effects of our new compounds were investigated in the MCF-7 (ER+)
and MDA-MB231 (ER-) breast cancer cell lines. The data shown in Figure 8.2 and
Table 8.1 indicate that most but not all of the modifications that were introduced in the
linker resulted in decreased toxicity toward both cell lines. The low toxicity of 5 and 7,
which contain guanidinium groups, may be related to either their poor uptake by cells or
their rapid excretion once absorbed (26). Despite showing reactivity toward DNA in
vitro, neither compound showed significant toxicity at the highest dose; that is, 20 g.M.
Lack of uptake may also be responsible for the low toxicity of 9, 12, and 14, which have
high logD values that are not predictive of good absorption (24). Further work is
warranted to determine if cellular uptake is indeed limiting for these compounds.
As previously reported, 1 was significantly more toxic toward MCF-7 cells than
MDA-MB231 cells (23). Compounds 13 and 15 containing amino groups showed
toxicity similar to that of 1. Both of these compounds also showed greater toxicity toward
the ER-positive MCF-7 cells than toward the ER-negative MDA-MB231 cells. This
result was consistent with our intended mechanisms, since the RBAs and reactivities with
DNA of 13 and 15 imply greater toxicity on ER-positive cells. It is interesting that the
results of the electrophoretic mobility shift assay indicate that DNA adducts of 1 have the
greatest affinity for the ER-LBD; compound 1 also shows the largest differential toxicity
between the two cell lines.
263
- Chapter8 8.5. CONCLUSION
We designed and synthesized a series of estradiol-aniline mustard-linked
bifunctional molecules that differ in their relative affinities for the ER and their capacities
to covalently modify DNA. The selective cytotoxic effects of 3 of these compounds (1,
13, 15) toward ER-positive breast cancer cells correlated with their ability to react with
DNA and their affinity for the ER, as well as favorable solubility. We are directing
research to determine if the interaction of the ER with DNA adducts formed by these
compounds is responsible for their selective effects.
264
- Chapter8 8.6. ACKNOWLEDGEMENTS
Research was supported by NIH grant CA77743, the Natural Sciences and
Engineering Research Council of Canada (PDF-242490-2001 Fellowship to AND), an
Award from the Susan B. Komen Foundation, and NIH grant SS10RR13386-01 for
support of the NMR and the mass spectrometry facility at MIT.
265
- Chapter8Figure 8.1. Structure and molecular features of lead compound.
bis(2-chloroethyl)
aniline
H
N-(CH2)3
A
(CH 2 )2C1
N1~~~~~~~
(CH2 )2 CI
Linker
(1)
266
- Chapter8 Table 8.1. RBA=relative binding affinity for the rabbit uterine ER as compared to
estradiol (RBA=100); % Oligo MODIFIED=percent of 16-mer cleaved by piperidine
after treatment with test compound for 24 h at 37 °C; % Oligo SHIFTED=percent of
covalently modified 16-mer that formed a slowly migrating complex with the ER-LBD
under electrophoresis; ED30=concentration of compound that resulted in 30% clonal
survival of cells exposed for 2 h.
Compound
1
5
7
9
12
13
14
15
RBA
46
10
28
6
29
29
13
40
Oligomodified
Oligoshiftd
C()
°()
45
29
58
3
39
79
14
44
93
93
38
ND
ND
38
ND
67
ogP
5.32
6.27
5.67
5.61
4.85
5.32
5.07
5.96
267
logD
222
3.12
2.52
5.61
4.85
222
5.07
2.87
ED30(uM)
MCF-7
5.1
>20
>20
>20
5.1
>20
5.1
MDA-MB231
9.6
>20
>20
>20
>20
7.9
>20
86
- Chapter8 Scheme 8.1. Synthesis of estradiol-linked aniline mustards. The starting compound 2 is
shown in the box (center). Reagents and conditions: (a) methanesulfonyl chloride, DIEA,
THF; (b) LiBr, TEA, DMF, 60 °C; (c) Ph 2P(O)NHCH 2CH 2 OTBDMS, NaH, TBAB, PhH, 60 °C; (d) TBAF, THF; (e) Boc-NHC(-N-Boc)SCH 3, PPh3, DIPAD, THF; (f) 4-(N,Nbis-2-chloroethylamino-phenyl)-propylamine, THF/H20 (90:10), reflux; (g) HCI/dioxane,
CH 2C12; (h) p-nitrophenylchloroformate, DIEA, THF; (i) phthalimide, DIPAD, PPh3,
THF; (j) Hydrazine, EtOH, reflux; (k) 4-(N-Boc)n-butyl(N-hydroxysuccinimide)ester,
TEA, DMF; (1) 4-(N,N-bis-chloroethylamino-phenyl)butyl(N-hydroxysuccinimide)ester,
TEA, DMF; (m) BH3 S(CH3) 2, THF, HC1; (n) HCI, THF. DIEA=diisopropylethyl amine;
TEA=triethyl amine; TBAB=tetrabutylammonium bromide; TBAF=tetrabutylammonium
fluoride;
DIPAD=diisopropylethyl
azadicarboxylate;
DMF=dimethylformamide;
THF=tetrahydrofuran.
Ri-C,_NL(CH
oR.(C·~NcH
1
g t
1 2R
2
mt
k
I,n
2)a-NHBoc
|,
a-d
R,-(CH2)-NH 2
IJ
10
,I2
-
14
0
h
R1-(C
2
eJ N.-Bo
c
(
R1-(CH2)-N
i Boc
t,g
6
RI-(2C)-N-(CH2)2-N1
A_.
Boc
NH
NH
1
Ri-(H2C)-W(CH -N
2)2 N-(CH
RI-(CH2H-N(CH.2)4-R2
15
13
(CH.2-OH
Je 3
NBc
O,, (Phh
r f,g
H12-
(CH,)4N-(CH24--R
R-(CH,-
12
HL0
R1-(CH2) 6-N-L(CH
11
R--(HC)-N
I
-(CH
Ri4(CH2-0-U-.W(CHa-
R2
9
NH
2)-R 2
Ri-(CH2)-N 11N-(CH
2)3-R2
5
7
R1= ,,,.<1
R2 =---N-(CH2)2C
.OX
X = TPH 2,3,4,6,6
X = H 5,7,9,12,13,14,1!
xo
|
268
2
- Chapter8 Figure 8.2. Survival of MCF-7 (ER+) and MDA-MB23 1 (ER-) breast cancer cells after
2 h exposure to estradiol-linked toxicants.
MCF-7
MDA-MB231
1
'Ft
.2
;n
DD
c
a)
(D
Concentration
-
AmineCarb (1) --
AmineGua(S)--
Diamine (13)
FM
Guanidine(7)
Amine (15)
-
269
'
Amide (14)
Carbamate (9) -4-Diamide
m
Chlorambucil (CMB)
(12)
Chapter8 8.7. REFERENCES
(1) Teicher, B. A. (1997) Antitumor alkylating agents. In Cancer Principles and
Practice (DeVita, V. T., Hellman, S., and Rosenberg, S. A., Eds.) pp 405-417,
Lippincott-Raven, New York.
(2) Hurley, L. H. (2002) DNA and its associated processes as targets for cancer
therapy. Nat. Rev. Cancer. 2(3), 188-200.
(3) Barret, J. M., and Hill, B. T. (1998) DNA repair mechanisms associated with
cellular resistance to antitumor drugs: potential novel targets. Anticancer Drugs.
9(2), 105-123.
(4) Dolan, E. M. (1997) Inhibition of DNA repair as a means of increasing the
antitumor activity of DNA reactive agents. Adv. Drug Del. Rev. 26, 105-118.
(5)
Rink, S. M., Yarema, K. J., Solomon, M. S., Paige, L. A., Tadayoni-Rebek,
B. M.,
Essigmann, J. M., and Croy, R. G. (1996) Synthesis and biological activity of
DNA damaging agents that form decoy binding sites for the estrogen receptor.
Proc. Natl. Acad. Sci. U. S. A. 93(26), 15063-15068.
(6) Mitra, K., Marquis,
J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
(7) Rink, S. M., Yarema, K. J., Solomon, M. S., Paige, L. A., Tadayoni-Rebek,
B. M.,
Essigmann, J. M., and Croy, R. G. (1996) Synthesis and biological activity of
DNA damaging agents that form decoy binding sites for the estrogen receptor.
Proc. Natl. Acad. Sci. U.S. A. 93(26), 15063-15068.
(8) Mitra, K., Marquis,
J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
(9) Rink, S. M., Yarema, K. J., Solomon, M. S., Paige, L. A., Tadayoni-Rebek,
B. M.,
Essigmann, J. M., and Croy, R. G. (1996) Synthesis and biological activity of
DNA damaging agents that form decoy binding sites for the estrogen receptor.
Proc. Natl. Acad. Sci. U. S. A. 93(26), 15063-15068.
(10)
Mitra, K., Marquis,
J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
270
- Chapter8 (11) Mitra, K., Marquis, J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
(12) Korenman, S. G. (1970) Endocrinology87, 1119-1123.
(13)
Linney, I. D., Buck, I. M., Harper, E. A., Kalindjian,
S. B., Pether, M. J.,
Shankley, N. P., Watt, G. F., and Wright, P. T. (2000) Design, synthesis, and
structure-activity relationships of novel non-imidazole histamine H(3) receptor
antagonists. J. Med. Chem. 43(12), 2362-2370.
(14)
Linney,
I. D., Buck, I. M., Harper, E. A., Kalindjian,
S. B., Pether, M. J.,
Shankley, N. P., Watt, G. F., and Wright, P. T. (2000) Design, synthesis, and
structure-activity relationships of novel non-imidazole histamine H(3) receptor
antagonists. J. Med. Chem. 43(12), 2362-2370.
(15) Linney,
I. D., Buck, I. M., Harper, E. A., Kalindjian,
S. B., Pether, M. J.,
Shankley, N. P., Watt, G. F., and Wright, P. T. (2000) Design, synthesis, and
structure-activity relationships of novel non-imidazole histamine H(3) receptor
antagonists. J. Med. Chem. 43(12), 2362-2370.
(16) Mitra, K., Marquis, J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
(17) Hughes, D. L. Organic Reactions (1992) John Wiley and Sons, New York.
(18) Brozwn, H. C., and Heim, P. J. (1973) Selective reductions. XVIII. Fast reaction
of primary, secondary, and tertiary amides with diborane. Simple, convenient
procedure for the conversion of amides to the corresponding amines. J. Org.
Chem. 38(5), 912-916.
(19) Korenman, S. G. (1970) Endocrinology87, 1119-1123.
(20)
Pike, A. C., Brzozowski, A. M., Walton, J., Hubbard, R. E., Thorsell, A. G., Li,
Y. L., Gustafsson, J. A., and Carlquist, M. (2001) Structural insights into the
mode of action of a pure antiestrogen. Structure. 9(2), 145-153.
(21)
Pike, A. C., Brzozowski, A. M., Walton, J., Hubbard, R. E., Thorsell, A. G., Li,
Y. L., Gustafsson, J. A., and Carlquist, M. (2001) Structural insights into the
mode of action of a pure antiestrogen. Structure. 9(2), 145-153.
(22)
Holley,
J. L., Mather, A., Wheelhouse,
R. T., Cullis, P. M., Hartley, J. A.,
Bingham, J. P., and Cohen, G. M. (1992) Targeting of tumor cells and DNA by a
chlorambucil-spermidine conjugate. Cancer Res. 52(15), 4190-4195.
271
- Chapter8 (23) Mitra, K., Marquis, J. C., Hillier, S. M., Rye, P. T., Zayas, B., Lee, A. S.,
Essigmann, J. M., and Croy, R. G. (2002) A rationally designed genotoxin that
selectively destroys estrogen receptor-positive breast cancer cells. J. Am. Chem.
Soc. 124(9), 1862-1863.
(24)
Lipinski,
C. A., Lombardo,
F., Dominy,
B. W., and Feeney,
P.
(1997)
Experimental and computational approaches to estimate solubility and
permeability in drug discovery and development settings. J. Adv. Drug Del. Rev.
23, 3-25.
(25) Horvath, Cs., Melander, W., and Molnar, I. (1977) Anal. Chem. 49, 865.
(26) Zhang, L., Brett, C. M., and Giacomini, K. M. (1998) Role of organic cation
transporters in drug absorption and elimination. Annu. Rev. Pharmacol. Toxicol.
38:431-60., 431-460.
272
PETER T. RYE
(617)253-6752
ptrye@mit.edu
Massachusetts Institute of Technology
77 Massachusetts Avenue, 56-669
Cambridge, MA, 02139
EDUCATION
Massachusetts Institute of Technology - Department of Chemistry
Doctor of Philosophy in Chemistry, June 2006. GPA: 4.8/5.0
Syracuse University - College of Arts and Sciences
B.S. Biochemistry, May 1999. Magna cum laude with honors. GPA: 3.8/4.0
Minors: Quantitative Analysis and Philosophy
PUBLICATIONS
Design, synthesis, and evaluation of estradiol-linked genotoxicants as anti-cancer agents. U. Sharma,
J. C. Marquis, A. Nicole Dinaut, S. M. Hillier, B. Fedeles, P. T. Rye, J. M. Essigmann, and R. G.
Croy. Bioorganic & Medicinal Letters (2004) 14/14, 3829-3833.
Intermetal Coupling in [(rlS-C5 R 5)Fe(dppe)] 2(Ji-CH=CHCH=CH) and in their Dication and
Monocation, Mixed-Valence Forms. Chung, M.-C.; Gu, X.; Etzenhouser, B. A.; Spuches, A. M.; Rye,
P. T.; Seetharaman, S. K.; Rose, D. J.; Zubieta, J.; Sponsler, M. B. Organometallics (2003) 22, 3485-
3494.
A Rationally Designed Genotoxin that Selectively Destroys Estrogen Receptor-Positive Breast Cancer
Cells, Mitra, K.; Marquis, J. C.; Hillier, S. M.; Rye, P. T.; Zayas, B.; Lee, A. S.; Essigmann, J. M.;
Croy, R. G.; J. Am. Chem. Soc.; (Communication); 2002; 124(9); 1862-1863.
HONORS/AWARDS
Karl Taylor Compton Prize for outstanding contributions to the MIT community, May 2006.
Best Poster Award, The 7 th Annual Northeastern Student Chemistry Research Conference, April 2005.
The William L. Stewart Jr. Institute Award for devotion to student life at MIT, April 2004.
Fellowship funding from The National Institutes of Health, September 2000 - January 2005.
Outstanding Teaching Awards from the MIT Chemistry Department, Fall 1999 and Spring 2000.
Phi Beta Kappa National Honors Society, September 1998.
Who's Who Among Students in American Universities and Colleges, 1997-1998.
Lt. Adolph Ornstein Science Scholarship, January 1998, January 1999, and September 1999.
Syracuse University Upper, May 1999, and Lower Division Honors, May 1997.
Golden Key National Honor Society, 1997.
Phi Eta Sigma National Honor Society, September 1996.
LEADERSHIP/EXTRACURRICULUR
Elected to the MIT Corporation Joint Advisory Committee, October 2004.
Secured funding for, and conducted, a series of conflict negotiation seminars, 2002.
Directed a workshop for Northeastern University's Center for Effective Teaching, September 2001.
Mentored undergraduates through The MIT Minorities Summer Research Program, 2001 and 2002.
Instructed specific lectures for MIT summer biochemistry programs, July 2001.
Founded Chemistry REFS (Resources for Easing Friction and Stress) Program at MIT, February 2001.
Certified as a mediator for the Commonwealth of Massachusetts, June 2000.
President of Watson Hall at Syracuse University, January - May 1997.
273