University of Iowa Iowa Research Online eses and Dissertations Summer 2013 Studies of Proliferating Cell Nuclear Antigen Mutant Proteins Defective in Translesion Synthesis and Mismatch Repair Lynne Margaret Dieckman University of Iowa Copyright 2013 Lynne Margaret Dieckman is dissertation is available at Iowa Research Online: hps://ir.uiowa.edu/etd/4839 Follow this and additional works at: hps://ir.uiowa.edu/etd Part of the Cell Biology Commons Recommended Citation Dieckman, Lynne Margaret. "Studies of Proliferating Cell Nuclear Antigen Mutant Proteins Defective in Translesion Synthesis and Mismatch Repair." PhD (Doctor of Philosophy) thesis, University of Iowa, 2013. hps://doi.org/10.17077/etd.5pihvjjx
Transcript
University of IowaIowa Research Online
Theses and Dissertations
Summer 2013
Studies of Proliferating Cell Nuclear AntigenMutant Proteins Defective in Translesion Synthesisand Mismatch RepairLynne Margaret DieckmanUniversity of Iowa
Copyright 2013 Lynne Margaret Dieckman
This dissertation is available at Iowa Research Online: https://ir.uiowa.edu/etd/4839
Follow this and additional works at: https://ir.uiowa.edu/etd
Part of the Cell Biology Commons
Recommended CitationDieckman, Lynne Margaret. "Studies of Proliferating Cell Nuclear Antigen Mutant Proteins Defective in Translesion Synthesis andMismatch Repair." PhD (Doctor of Philosophy) thesis, University of Iowa, 2013.https://doi.org/10.17077/etd.5pihvjjx
MUTANT PROTEINS DEFECTIVE IN TRANSLESION SYNTHESIS
AND MISMATCH REPAIR
by
Lynne Margaret Dieckman
A thesis submitted in partial fulfillment of the requirements for the Doctor of Philosophy degree in Molecular and Cellular Biology in the Graduate College of The University of
Iowa
August 2013
Thesis Supervisor: Associate Professor M. Todd Washington
Graduate College The University of Iowa
Iowa City, Iowa
CERTIFICATE OF APPROVAL
PH.D. THESIS
This is to certify that the Ph.D. thesis of
Lynne Margaret Dieckman
has been approved by the Examining Committee for the thesis requirement for the Doctor of Philosophy degree in Molecular and Cellular Biology at the August 2013 graduation.
Thesis Committee: M. Todd Washington, Thesis Supervisor Marc Wold Kris DeMali John Dagle Jon Houtman
ii
ACKNOWLEDGMENTS
First and foremost, I would like to thank Mr. Joel Hutchison for playing an
integral role in establishing my interest and foundation in science when I was a high
school student. Under his instruction, I gained the knowledge and confidence necessary
to succeed as a college student and persevere as a graduate student. I would not be in
science today if it had not been for his constant direction and encouragement to pursue a
career path in chemistry. Even though his classes only lasted for four years of my
educational life, he has continued to be a cherished mentor since then and will continue to
be for the rest of my life.
I would like to thank all of my family and friends for their continued support and
reassurance throughout my graduate school career. I am extremely lucky to have such an
amazing group of people who have been there for me under any circumstance. Just to
name a couple, my mom has been the most loving and supportive parent that I could ask
for. And Tyson Shepherd has been a fantastic help in and out of the lab with all of our
entertaining and valuable discussions.
Lastly, I would like to thank my thesis advisor, Dr. M. Todd Washington, for
providing an environment adequate for me to learn and grow during both my Master’s
and Ph.D. work. His confidence in my abilities facilitated my development as a scientist,
and I would not be where I am today if it were not for his continued support and guidance
throughout the years. He was a fantastic and respected mentor during graduate school
and will always remain a valued friend.
iii
ABSTRACT
Proliferating cell nuclear antigen (PCNA) is a versatile protein involved in all
pathways of DNA metabolism. It is best known as a processivity factor for classical
polymerases, which synthesize DNA on non-damaged templates during DNA replication
(ex: pol δ). Non-classical polymerases, on the other hand, are those that synthesize DNA
on damaged templates (ex: pol η). PCNA also functions in repair, recombination, and
most other DNA-dependent cellular processes. A number of separation of function
mutant PCNA proteins have been identified, suggesting that PCNA could be a valuable
target to manipulate DNA metabolism. This thesis focuses on the study of PCNA mutant
proteins that affect translesion synthesis (TLS) and mismatch repair (MMR).
During TLS, the process by which DNA polymerases replicate through DNA
lesions, PCNA recruits and stabilizes polymerases at the replication fork. TLS requires
the monoubiquitylation of PCNA, and PCNA and ubiquitin-modified PCNA (Ub-PCNA)
stimulate TLS by classical and non-classical polymerases. Two mutant forms of yeast
PCNA, one with an E113G substitution and one with a G178S substitution, support
normal cell growth but inhibit TLS. To better understand the role of PCNA in TLS, I re-
examined the structures of both mutant PCNA proteins and identified substantial
disruptions of the subunit interface that forms the PCNA trimer. This resulted in reduced
trimer stability in the mutant proteins. The mutant forms of PCNA and Ub-PCNA do not
stimulate TLS of an abasic site by either classical pol or non-classical pol . Normal
replication by pol was also impacted, but normal replication by pol was much less
affected. These findings support a model in which reduced trimer stability causes these
mutant PCNA proteins to occasionally undergo conformational changes that compromise
their ability to stimulate TLS by both classical and non-classical polymerases.
During MMR, PCNA recruits and coordinates proteins involved in the mismatch
recognition, excision, and resynthesis steps. Previously, two mutant forms of PCNA
iv
were identified that cause defects in MMR with little if any other defects. These are the
C22Y and C81R mutant PCNA proteins. In order to understand the structural and
mechanistic basis by which these two substitutions in PCNA proteins block MMR, we
solved the X-ray crystal structures of both mutant proteins and carried out further
biochemical studies. I found that these amino acid substitutions lead to distinct structural
changes in PCNA. The C22Y substitution alters the positions of the -helices lining the
central hole of the PCNA ring, whereas the C81R substitution creates a distortion in the
-sheet at the PCNA subunit interface. I conclude that the structural integrity of the -
helices lining the central hole and the -sheet at the subunit interface are both necessary
to form productive complexes with MutS and mismatch-containing DNA.
As described above, my studies focused on four amino acid substitutions in
PCNA that disrupt TLS and MMR: the E113G and G178S substitutions cause defects in
TLS while the C22Y and C81R substitutions cause defects in MMR. The structures of
these mutant PCNA proteins revealed that three of the four substitutions caused
disruptions near the subunit interface of PCNA. To further examine the importance of
this region, we generated random mutations of the PCNA subunit interface and
performed in vivo genetics assays and in vitro biochemical assays to examine their effects
on TLS and MMR. We determined that the subunit interface of PCNA is very dynamic
and that small changes at this interface can cause drastically different effects on TLS and
MMR. Moreover, we suggest that the integrity of the subunit interface as well as the
nearby β-strands in domain A are crucial for proper PCNA function in vivo and in vitro.
v
TABLE OF CONTENTS
LIST OF TABLES ........................................................................................................... viii LIST OF FIGURES ........................................................................................................... ix LIST OF ABBREVIATIONS .......................................................................................... xiii CHAPTER 1 INTRODUCTION .........................................................................................1
DNA Metabolism and Carcinogenesis .............................................................................1 DNA replication, repair, and recombination ................................................................1 DNA mismatches and mistmatch repair .......................................................................5 Mutagenesis and translesion synthesis .........................................................................7
DNA Polymerases ............................................................................................................8 Overview of DNA polymerases ...................................................................................8 Classical DNA polymerases .........................................................................................9 Classical DNA polymerase δ ......................................................................................10 Non-classical DNA polymerases ................................................................................12 Non-classical DNA polymerase η ..............................................................................13
Eukaryotic MutS Homologues MSH2 and MSH6 .........................................................15 Structure and function of the MSH2 and MSH6 proteins .........................................15 Interactions with PCNA and other repair proteins .....................................................17
Proliferating Cell Nuclear Antigen.................................................................................19 Structure and function of PCNA ................................................................................19 Role of PCNA in regulating DNA metabolism ..........................................................21
Post-Translational Modifications of PCNA ...................................................................24 Overview of PCNA modifications .............................................................................24 Monoubiquitylation of PCNA ....................................................................................24 Polyubiquitylation of PCNA ......................................................................................28 Sumoylation of PCNA ................................................................................................29
Structures of PCNA Complexes .....................................................................................30 Insights from structures of PCNA bound to PIP peptides ..........................................30 Structures of PCNA bound to full-length proteins .....................................................33 Low resolution structures of PCNA complexes .........................................................34 Conclusions from structural studies with PCNA........................................................35
Interactions of Y-family Polymerases with PCNA and Ubiquitylated PCNA ...............36 Interactions with un-modified PCNA .........................................................................37 Interactions with ubiquitin-modified PCNA ..............................................................38
Polymerase Switching and the Tool Belt Model ............................................................40 Polymerase switching during translesion synthesis ...................................................40 The tool belt model of polymerase switching ............................................................41
Mutant PCNA Proteins ...................................................................................................43 Mutant PCNA proteins defective in translesion synthesis .........................................43 Mutant PCNA proteins defective in mismatch repair ................................................45
SYNTHESIS BY DNA POLYMERASE η AND BY DNA POLYMERASE δ ...........76
Abstract ..........................................................................................................................76 Introduction ....................................................................................................................77 Materials and Methods ...................................................................................................79
vi
Protein expression and purification ............................................................................79 DNA and nucleotide substrates ..................................................................................80 PCNA trimer stability assays......................................................................................80 Enzyme-linked immunosorbent assays ......................................................................81 Polymerase processivity assays ..................................................................................81 Polymerase activity assays .........................................................................................82
Results ............................................................................................................................83 The E113G and G178S mutant PCNA proteins have altered subunit interfaces .......83 Trimer stability of the E113G and G178S mutant PCNA proteins ............................84 Interactions of the E113G mutant PCNA protein with DNA polymerases ................86 Impact of the E113G mutant PCNA protein on the activity of pol η .........................86 Impact of the E113G mutant PCNA protein on the catalytic efficiency of pol η ......88 Impact of the E113G mutant PCNA protein on the activity of pol δ .........................89 Impact of the E113G mutant PCNA protein on the catalytic efficiency of pol δ .......90 Impact of the G178S mutant PCNA protein on the activity of pol δ .........................91
Discussion ......................................................................................................................91 Inhibition of TLS in yeast by mutant PCNA proteins ................................................91 Mechanism of TLS inhibition by mutant PCNA proteins ..........................................92 Inhibition of TLS is caused by PCNA trimer instability ............................................93 Model of TLS inhibition by PCNA trimer instability ................................................94
CHAPTER 3 DISTINCT STRUCTURAL ALTERATIONS IN PCNA BLOCK DNA MISMATCH REPAIR .......................................................................................113
Abstract ........................................................................................................................113 Introduction ..................................................................................................................114 Materials and Methods .................................................................................................116
Protein expression and purification ..........................................................................116 DNA and nucleotide substrates ................................................................................116 Crystallization of the C22Y and C81R mutant proteins ...........................................117 Data collection and structural determination ............................................................117 PCNA trimer stability assays....................................................................................118 Polymerase δ activity assays ....................................................................................118 Enzyme-linked immunosorbent assays ....................................................................118 Sedimentation assays ................................................................................................119
Results ..........................................................................................................................120 Structure of the C22Y mutant PCNA protein...........................................................120 Structure of the C81R mutant PCNA protein ...........................................................121 Stability of the mutant PCNA proteins .....................................................................121 Impact of the mutant PCNA proteins on DNA polymerase δ activity .....................122 Interactions of the mutant PCNA proteins with MutSα ...........................................123 Interactions of the mutant PCNA proteins with MutSα and DNA ...........................124
CHAPTER 4 IDENTIFICATION AND CHARACTERIZATION OF RANDOM MUTATIONS OF THE PCNA SUBUNIT INTERFACE ..............................................142
Abstract ........................................................................................................................142 Introduction ..................................................................................................................142 Materials and Methods .................................................................................................146
Protein expression and purification ..........................................................................146 DNA and nucleotide substrates ................................................................................146 PCNA trimer stability assays....................................................................................147
Genetic analyses of random PCNA interface mutant proteins .................................148 Trimer stability of the PCNA interface mutant proteins ..........................................149 Impact of the PCNA interface mutant proteins on the activity of pol η ...................150 Impact of the PCNA interface mutant proteins on the activity of pol δ ...................151 Structures of the PCNA interface mutant proteins ...................................................153
Discussion ....................................................................................................................154 CHAPTER 5 THE C-TERMINAL REGION OF DNA POLYMERASE η IS INTRINSICALLY DISORDERED AND REQUIRED FOR INTERACTION WITH PCNA AND MONOUBIQUITYLATED PCNA ................................................168
Abstract ........................................................................................................................168 Introduction ..................................................................................................................168 Materials and Methods .................................................................................................170
Protein expression and purification ..........................................................................170 Protein disorder prediction studies ...........................................................................171 Nuclear magnetic resonance spectroscopy ...............................................................171 Enzyme-linked immunosorbent assays ....................................................................172 Isothermal titration calorimetry experiments ...........................................................172 Crystallization of the Ub-PCNA-CTR fusion protein ..............................................173
Results ..........................................................................................................................173 The C-terminal region of pol η is intrinsically disordered .......................................173 Binding studies of the CTR of pol η with PCNA, ubiquitin, and Ub-PCNA ...........174 Crystallography studies of the complex of the CTR of pol η and Ub-PCNA ..........176
Integrity of the PCNA Interface during Translesion Synthesis ....................................192 Overview of studies with the E113G mutant PCNA proteins ..................................192 Role of Ub-PCNA during TLS .................................................................................193 Possible insights from studies with the E113G mutant PCNA and Ub-PCNA proteins ...................................................................................................194
Defects in Mismatch Repair Caused by Distinct Structural Alterations in PCNA ......195 Overview of studies with the C22Y and C81R mutant PCNA proteins...................195 How do the C22Y and C81R substitutions prevent productive PCNA complex formation during MMR? ..........................................................................................196
Importance of the Subunit Interface of PCNA for Translesion Synthesis and Mismatch Repair ..........................................................................................................198
Overview of studies with random PCNA interface mutant proteins ........................198 Shearing of the PCNA interface ...............................................................................200 Growth phenotypes of the S177G and G178S mutant PCNA proteins ....................201 PCNA-dependent processes and PCNA substitutions that disrupt them..................202
PCNA and Its Interactions with Intrinsically Disordered Proteins ..............................203 Overview of studies with the C-terminal region of pol η .........................................203 Regulation of PCNA interactions .............................................................................204
Table 1.1. Classification of DNA polymerases .................................................................55 Table 2.1. Distances between potential hydrogen bond donor and acceptor
atoms at the PCNA subunit interface ................................................................97 Table 2.2. Percentage of PCNA proteins in the monomeric state as determined
by size exclusion chromatography ..................................................................101 Table 2.3. Processivity of pol η on non-damaged and damaged DNA ...........................105 Table 2.4. Steady state kinetics of nucleotide incorporation by pol η .............................107 Table 2.5. Processivity of pol δ on non-damaged and damaged DNA ...........................109 Table 2.6. Steady state kinetics of nucleotide incorporation by pol δ .............................111 Table 3.1. Data collection and refinement statistics ........................................................129 Table 3.2. Relative DNA synthesis by pol δ in the presence of PCNA
mutant proteins ................................................................................................136 Table 4.1. Summary of genetic studies with the PCNA interface mutant proteins .........159 Table 4.2. Distances between potential hydrogen bond donor and acceptor
atoms at the PCNA subunit interface ..............................................................165 Table 4.3. Summary of in vivo and in vitro studies with the PCNA interface mutant
Figure 1.1. Model of DNA replication in eukaryotes ........................................................50 Figure 1.2. Common types of DNA damage .....................................................................51 Figure 1.3. Possible DNA mismatches ..............................................................................52 Figure 1.4. Model of mismatch repair in eukaryotes ........................................................53 Figure 1.5. Model of translesion synthesis .......................................................................54 Figure 1.6. Mechanism of DNA polymerization by polymerases ....................................56 Figure 1.7. Structure of the catalytic subunit (Pol3) from DNA polymerase δ
from yeast bound to DNA ..............................................................................57 Figure 1.8. Structure of DNA polymerase from the bacteriophage RB69........................58 Figure 1.9. Structure of DNA polymerase η from yeast ...................................................59 Figure 1.10. Structural model of the full-length pol η .......................................................60 Figure 1.11. Structure of the ubiquitin-binding zinc-finger (UBZ) of pol η .....................61 Figure 1.12. Structure of the human MutSα dimer bound to a G:T mispair ......................62 Figure 1.13. Structure of yeast PCNA ...............................................................................63 Figure 1.14. Structure of yeast PCNA bound to DNA .....................................................64 Figure 1.15. Structure of ubiquitin-modified PCNA ........................................................65 Figure 1.16. Overlay of the two positions occupied by ubiquitin in the crystal
structure of Ub-PCNA .................................................................................66 Figure 1.17. The potential “ubiquitin-switch” on PCNA .................................................67 Figure 1.18. Overlay of the structures of ubiquitylated and sumoylated PCNA ..............68 Figure 1.19. Structures of PCNA bound to PIP peptides ..................................................69 Figure 1.20. Structure of PCNA bound to FEN1 ...............................................................70 Figure 1.21. The structured and unstructured regions of Y-family polymerases ..............71 Figure 1.22. Structural models of the full-length Y-family polymerases ..........................72 Figure 1.23. Structural model of full length pol bound to
Figure 1.24. The tool belt model of translesion DNA synthesis ........................................74 Figure 1.25. Structures of the G178S and E113G mutant PCNA proteins ........................75 Figure 2.1. The subunit interface of the wild-type and mutant PCNA proteins ...............96 Figure 2.2. Analysis of the wild-type and mutant PCNA proteins by native gel
electrophoresis ................................................................................................98 Figure 2.3. Analysis of the wild-type and mutant PCNA proteins by size exclusion
chromatography ..............................................................................................99 Figure 2.4. Stability of the wild-type and mutant Ub-PCNA proteins ............................102 Figure 2.5. Interaction of the wild-type and mutant PCNA and Ub-PCNA
proteins with pol .......................................................................................103 Figure 2.6. Processive DNA synthesis by pol in the presence of the wild-type
and mutant PCNA proteins ...........................................................................104 Figure 2.7. Steady state kinetics of pol in the presence of the wild-type
and mutant PCNA proteins ..........................................................................106
Figure 2.8. Processive DNA synthesis by pol in the presence of the wild-type and mutant PCNA proteins ...........................................................................108
Figure 2.9. Steady state kinetics of pol in the presence of the wild-type and
mutant PCNA proteins .................................................................................110 Figure 2.10. Processive DNA synthesis by pol δ in presence of the E113G
and G178S mutant PCNA proteins ...........................................................112 Figure 3.1. Structure of the C22Y mutant PCNA protein................................................130 Figure 3.2. Structure of the C81R mutant PCNA protein ................................................131 Figure 3.3. Analysis of the PCNA proteins by native gel electrophoresis ......................132 Figure 3.4. Analysis of the PCNA proteins by size exclusion chromatography..............133 Figure 3.5. Analysis of the C81R mutant PCNA protein by size exclusion
chromatography ...........................................................................................134 Figure 3.6. DNA synthesis by pol in the presence of the PCNA proteins ....................135
Figure 3.7. Interactions of the PCNA proteins with MutS ............................................137 Figure 3.8. Sedimentation analysis of the interactions of the PCNA
proteins with MutS and mistmatched DNA ...............................................138 Figure 3.9. Sedimentation analysis of the interactions of the PCNA
proteins with MutS and homoduplex DNA ..............................................140
xi
Figure 4.1. The amino acid residues that comprise the PCNA subunit interface ...........158 Figure 4.2. Analysis of the wild-type and mutant PCNA proteins by native gel
electrophoresis ..............................................................................................160 Figure 4.3. Analysis of the PCNA interface mutant proteins by size exclusion
chromatography ............................................................................................161 Figure 4.4. DNA synthesis by pol η in the presence of the PCNA mutant proteins .......162 Figure 4.5. DNA synthesis by pol on a non-damaged template in the presence
of the PCNA mutant proteins .......................................................................163 Figure 4.6. DNA synthesis by pol on a template containing an abasic site in the
presence of the PCNA mutant proteins ........................................................164
Figure 4.7. Structure of the V180A mutant PCNA protein ............................................166 Figure 5.1. The structured and unstructured regions of yeast pol η ................................179 Figure 5.2. The
1H-
15N heteronuclear single quantum coherence (HSQC)
spectrum of the CTR of pol η ......................................................................180 Figure 5.3. Analysis of the interaction of the PCNA and Ub-PCNA proteins
with pol using ELISA ..............................................................................181 Figure 5.4. Analysis of the interaction of the full-length pol and the
CTR of pol η with Ub-PCNA .......................................................................182 Figure 5.5. Analysis of the interaction of the CTR of pol with PCNA and
ubiquitin using ITC ......................................................................................183 Figure 5.6. Analysis of the interaction of the CTR of pol with
Ub-PCNA using ITC ....................................................................................184 Figure 5.7. The
1H-
15N HSQC spectra of the CTR of pol η bound to PCN ..................185
Figure 5.8. The
1H-
15N HSQC spectra of the CTR of pol η bound to Ub-PCNA ..........187
Figure 5.9. Production and purification of the Ub-PCNA-CTR fusion protein ..............189 Figure 5.10. Crystallization of the Ub-PCNA-CTR fusion protein ................................190 Figure 6.1. Positions of all PCNA substitutions used for studies ..................................207 Figure 6.2. Close-up of positions of all PCNA substitutions used for studies ...............208 Figure 6.3. Potential positions of the ubiquitin moiety on PCNA and
possible roles of these positions during TLS ................................................209 Figure 6.4. SAXS analysis of the E113G mutant Ub-PCNA protein .............................210 Figure 6.5. Non-denaturing gel electrophoresis of wild-type PCNA .............................211
xii
Figure 6.6. B-factors of the PCNA interface .................................................................212
xiii
LIST OF ABBREVIATIONS
8-oxoG – 7,8-dihydro-8-oxoguanine
BER – base excision repair
Can-R – canavanine-resistant
CTR – C-terminal region
dNTP – deoxyribonucleotide
DSB – double-strand break
ELISA – enzyme-linked immunosorbent assay
EM – electron microscopy
EXOI – exonuclease I
FAD – flavin adenine dinucleotide
FEN1 – flap endonuclease I
HSQC – heteronuclear single quantum coherence
HU – hydroxyurea
IDCL – interdomain connector loop
IDL – insertion/deletion loop
ITC – isothermal titration calorimetry
MCM – mini chromosome maintenance
MLH – MutL homologue
MMR – mismatch repair
MMS – methyl methanesulfonate
MSH – MutS homologue
NER – nucleotide excision repair
NHEJ – nonhomologous end joining
NMR – nuclear magnetic resonance
NTR – N-terminal region
PAD – polymerase associated domain
xiv
PAGE – polyacrylamide gel electrophoresis
PCNA – proliferating cell nuclear antigen
PIP – PCNA interacting peptide
Pol – polymerase
PUb-PCNA – polyubiquitylated PCNA
RFC – replication factor C
RPA – replication protein A
SAXS – small angle X-ray scattering
ssDNA – single-stranded DNA
SUMO – small ubiquitin-like modifier
TLS – translesion synthesis
TT dimer – thymine-thymine dimer
UBM – ubiquitin-binding motif
Ub-PCNA – monoubiquitylated PCNA
UBZ – ubiquitin-binding zinc-binding
UV – ultraviolet radiation
XP-V – xeroderma pigmentosum variant
α – alpha
δ – delta
ε – epsilon
ζ – zeta
η - eta
ι – iota
κ – kappa
1
CHAPTER 1
INTRODUCTION
(The section entitled “Structures of PCNA Complexes” has been published in
Dieckman, L.M., Freudenthal, B.D., and Washington, M.T. (2012) Subcell Biochem.
(Springer) 62: 281-299. The section entitled “Interactions of Y-family Polymerases with
PCNA and Ubiquitylated PCNA” is to be published in Pryor, J.M., Dieckman, L.M.,
Boehm, E.M., and Washington, M.T. (2013) Nucl. Acids Mol. Biol. (Springer) (In
press).)
DNA Metabolism and Carcinogenesis
DNA replication, repair, and recombination.
All living organisms inherit their genetic information in the form of DNA. In
order for each daughter cell to acquire a complete genome from generation to generation,
this material must be duplicated during every round of cell division. The process of
duplicating DNA is called DNA replication and involves the coordinated action of
numerous enzymes and proteins. Figure 1.1 shows a model of DNA replication in
eukaryotes. In brief, the process is initiated by the separation of the two complementary
DNA strands in an ATP-dependent fashion by the mini chromosome maintenance
(MCM) complex, the major DNA helicase in eukaryotic organisms. Any single-stranded
DNA generated during replication is protected and prevented from re-annealing by
replication protein A (RPA), a heterotrimeric single-stranded DNA binding protein that
binds DNA in a non-specific manner. From here, DNA synthesis begins bidirectionally,
and replication is divided into leading strand and lagging strand synthesis. All nucleotide
synthesis in cells is carried out in the 5ꞌ to 3ꞌ direction by enzymes called polymerases. In
eukaryotes, the polymerase that accounts for leading strand replication is polymerase ε
2
(pol ε). Lagging strand synthesis is more complex than leading strand synthesis due to
the inability of polymerases to add nucleotides to DNA in a 3ꞌ to 5ꞌ direction. To
overcome this obstacle, lagging strand synthesis is performed in short fragments and,
subsequently, the DNA fragments are joined together. This procedure is initiated by the
introduction of RNA primers by polymerase α (pol α), an RNA primase, and the
remaining DNA replication on this strand is then carried out by polymerase δ (pol δ).
Lagging strand synthesis is completed by the concerted actions of flap endonuclease 1
(FEN1) and a DNA ligase, which remove the RNA primer and seal the gap between the
resulting two fragments of DNA in an ATP-dependent manner. Polymerases are
recruited to the replication fork by a replication accessory factor called proliferating cell
nuclear antigen (PCNA), a homotrimeric sliding clamp that prevents polymerases from
dissociating from the DNA during replication. PCNA also interacts with and coordinates
the actions of all of these proteins involved in lagging strand synthesis at the replication
fork.
It is imperative that DNA be replicated accurately and efficiently to avoid any
type of alteration in the DNA sequence from one cell division to the next. However,
errors occur during each round of replication at a rate of 10-9
[1]. In addition, our
genome is constantly being attacked by byproducts of cellular processes as well as by
environmental agents. Common examples of DNA damage are shown in Figure 1.2.
Exposure to ultraviolet light causes adjacent thymine residues to covalently bond to one
another to create thymine dimers or (6-4) photoproducts [2]. Ionizing radiation also
causes numerous types of DNA damage, including deleterious double-stranded breaks
[3]. Chemical factors, present in both the environment and originating within the cell
frequently damage DNA as well. For example, oxygen free radicals that are released as
by-products of chemical reactions during metabolism in cells cause damage such as 7,8-
dihydro-8-oxoguanine (8-oxoG) [4, 5]. These lesions occur between 1000 and 2000
times per cell per day [6]. In addition to the lesions induced by damaging agents, other
3
types of damage result from spontaneous processes. For instance, uracil (which is
considered damage in the context of DNA) results from the deamination of a cytosine.
Similarly, an abasic site results from the hydrolysis of the N-glycosidic bond that attaches
the base to the sugar-phosphate backbone. This type of damage can occur at an
astounding rate of 10,000 sites per human cell per day [6, 7].
Fortunately, cells have evolved to deal with the presence of DNA replication
errors and lesions. In order to preserve their genetic information, they possess several
biological responses to DNA damage. Two major categories of pathways exist to repair
incorrect or damaged DNA: DNA repair and DNA damage tolerance. There are several
types of DNA repair mechanisms, including direct reversal of the damage, base and
nucleotide excision repair (BER and NER), mismatch repair, and double-strand break
repair, which includes both nonhomologous end joining (NHEJ) and homologous
recombination. The two main types of DNA damage tolerance pathways that exist are
translesion synthesis - the best studied DNA damage tolerance pathway – and the error-
free pathway. The focus of this thesis will be on both mismatch repair and translesion
synthesis and specific proteins involved in these pathways, which will be discussed
below.
Some lesions are repaired by direct reversal of the damaged base. One example
of this is the repair of ultraviolet light-induced damage by a single enzyme (photolyase)
existing in bacteria, plants, and animals. These enzymes utilize energy from light to
activate the flavin adenine dinucleotide (FAD) and break the covalent bonds of the
pyrimidine dimers. Photolyases, however, are not found in placental mammals, and so
these organisms rely on excision repair for removal of the lesion [8].
In general, excision repair requires that the damaged nucleotide(s) are removed,
that the resulting gap is filled using the opposite strand as a template, and finally that the
DNA is sealed by ligation. During BER, the damage is recognized and removed by a
DNA glycosylase [9]. This leaves an abasic site, which is then removed in two steps by
4
an AP endonuclease and an AP lyase [10, 11]. The resulting single nucleotide gap is
filled by a DNA polymerase and subsequently sealed by a DNA ligase [11-13]. Bulky
lesions that cause distortion in the DNA double helix are removed by NER [14]. NER
begins with recognition of the damaged base(s) by an enzyme complex and, unlike BER,
several DNA bases are removed on either side of the lesion, leaving a 27-29 nucleotide
gap. Similar to BER, the gap is filled by a DNA polymerase and sealed by a DNA ligase
[15].
Double-strand breaks (DSB) in DNA occur through several means. The main
cause of DSBs is exposure to ionizing radiation. In addition to this, reactive oxygen
species and chemotherapeutic agents that generate oxidative free radicals also cause
DSBs [16-18]. Moreover, when the replication machinery encounters certain forms of
DNA damage, the replication fork can collapse and form a DSB. These DSBs can be
extremely harmful to the DNA as they are capable of leading to chromosomal
rearrangements and, if left unrepaired, cell death.
Due to the extremely detrimental consequences of DSBs, it is imperative that cells
have a means of fixing them, which can occur through one of two methods – NHEJ and
homologous recombination. In humans, NHEJ is the preferred method, whereas
homologous recombination predominates in yeast. Homologous recombination is
initiated by the formation of a DSB. This is followed by resection, in which sections of
DNA at the 5ꞌ end of the break are processed to give 3ꞌ single-stranded tails. Resection
then stimulates strand invasion, where the single-stranded tails invade and align with
their complementary strands on the homologous chromosome and initiate DNA synthesis
by a DNA polymerase to fill in the degraded DNA [19, 20]. This type of DNA repair is
often accurate as it is able to use an existing complementary strand as a template for
nucleotide incorporation. Conversely, during NHEJ, the DNA ends of DSBs are directly
ligated together without the use of a homologous template. When perfectly compatible
overhangs are present after generation of the DSB, NHEJ accurately repairs the break.
5
When these overhangs are incompatible, however, NHEJ can lead to chromosomal
translocations, telomere fusion [21], and a gain or loss of nucleotides anywhere in the
genome.
DNA mismatches and mismatch repair.
Purine bases only form appropriate hydrogen bonding and structural geometry
when base-paired with their complementary pyrimidine. Inappropriate insertion of a
pyrimidine opposite another pyrimidine forms energetically unfavorable pairing because
the bases are not large enough to form the necessary hydrogen bonds. An example of this
situation is shown in Figure 1.3. Purine-purine mispairing, on the other hand, is
energetically unfavorable because the bases are too bulky and create steric clashes within
the double helix. Even though the incorporation of a mismatched base pair is highly
unfavorable, they are still produced frequently during DNA metabolism. DNA
mismatches arise from the inaccurate insertion, deletion, or mis-incorporation of
nucleotides during DNA replication, repair, and recombination [22, 23]. Like DNA
damage, mismatched base pairs must be removed to maintain genome integrity.
Recognition and repair of these mispairs is accomplished by mismatch repair (MMR)
proteins, thereby reducing mutation rates. Defects in MMR are most known for their
correlation with sporadic and hereditary human cancers, including hereditary non-
polyposis colorectal cancer [24-26].
The MMR pathway has been studied extensively in E. coli. In this system, the
homodimer MutS recognizes base-base mispairs and small nucleotide insertion/deletion
mispairs [25]. MutL, which also acts as a homodimer [27], then interacts with MutS and
enhances mismatch recognition [28]. This complex recruits and activates MutH in an
ATP-dependent manner. The process of MMR is initiated when MutH specifically nicks
the newly synthesized strand of DNA [25, 28]. The bacterial genome is hemimethylated
immediately after DNA replication, and thus the newly synthesized strand is defined by
6
the absence of methylation. This allows for the helicase UvrD to unwind the duplex
DNA from the nick towards the mismatch [29] and for subsequent degradation of the
single-stranded DNA by an exonuclease (EXOI, EXOX, EXOVII, or RecJ depending on
the location of the mismatch). The resulting gap is then resynthesized by DNA
polymerase III and sealed by DNA ligase [25, 30, 31].
The general scheme for which MMR occurs in eukaryotes is shown in Figure 1.4.
While MutS and MutL homologues are present in eukaryotes, no MutH homologue has
been identified. The eukaryotic MutS homologues MSH2, MSH3, and MSH6 exist as
heterodimers. Where the MSH2-MSH6 heterodimer (denoted MutSα) is the most
abundant and involved in the recognition of base-base mismatches and small
insertion/deletion loops (IDLs) [32-34], the MSH2-MSH3 heterodimer (denoted MutSβ)
is involved in the recognition and repair of longer IDLs [32, 34]. MutL homologues
MLH1, MLH2, MLH3, and PMS1 also function as heterodimers. The MLH1-PMS1
complex carries out the majority of MMR in yeast cells, while MLH1-MLH2 and MLH1-
MLH3 have been implicated in playing a minor role in the repair of mismatches [35, 36].
Like the MutL and MutS complex in E. coli, the MLH1-PMS1 heterodimer
interacts with either the MutSα or MutSβ complex at the site of the mismatched DNA
template [37, 38]. Although MMR in yeast and humans is not as well-defined as in
bacteria, the main proteins involved in strand degredation, resynthesis, and ligation have
been identified as exonuclease I (EXOI) [39-42], DNA polymerase δ [43], replication
protein A (RPA) [44, 45], replication factor C (RFC) [46, 47], high-mobility group box 1
[48], and DNA ligase I [49].
Unlike E. coli, eukaryotic DNA does not exhibit the presence of hemi-methylated
DNA to help discriminate strand specificity during MMR. It has been suggested that the
recognition of the newly synthesized daughter strand is determined by the DNA
biotins on both ends. The X represents the position of a non-damaged G or an abasic site.
The primer strand was a 31-mer oligodeoxynucleotide with the sequence 5'-TCG CAG
GTA GCC AGC CTC GCA CCC GTC CAA C. The primer strand was 5'-32
P-end-
labeled with T4 polynucleotide kinase and (γ-32
P)ATP and annealed to the template
strand at 1 µM in 25 mM TrisCl, pH 7.5, and 100 mM NaCl at 90oC for 2 min and slowly
cooled to 30oC. A mixture of all four dNTPs (10 mM each) was purchased from New
England Biolabs.
PCNA trimer stability assays.
For non-denaturing polyacrylamide gel electrophoresis (PAGE), the wild-type
and S177G, G178S, S179T, V180A, and I181R mutant PCNA proteins (0.1 to 1.0
mg/ml) were incubated in 25 mM TrisCl, pH 7.4, 150 mM NaCl, 0.01% bromophenol
blue, and 10% glycerol for 5 min and then run on a TrisCl pre-cast 4-20% gradient non-
denaturing polyacrylamide gel (Bio-Rad) at 4oC at a current of 25 mA using 25 mM Tris,
pH 8.3, and 0.2 M glycine running buffer. Protein bands were visualized using
Coomassie blue staining. For size exclusion chromatography, wild-type and mutant
PCNA proteins were diluted to 10 mg/ml in 25 mM Tris, pH 7.4, 150 mM NaCl, 5%
glycerol and run on a 120 ml HiLoad 16/60 Superdex 200 PG column (GE Healthcare).
Polymerase activity assays.
Running start assays were performed as described previously [161, 305].
Reactions were carried out in the absence of PCNA and in the presence of 90 nM wild-
type or mutant PCNA proteins (trimer concentration), and contained 20 nM pol or pol
η, 25 nM DNA, and 100 µM of each of the four dNTPs. Reactions were stopped after 1
and 2 minutes in the case of pol δ and after 30 min in the case of pol η. The extension
products were analyzed on a 15% polyacrylamide sequencing gel containing 8 M urea.
148
Results
Genetic analyses of random PCNA interface
mutant proteins.
Previous work in our lab has suggested that amino acid substitutions at the
subunit-subunit interface of PCNA that cause defects in TLS and MMR cause structural
perterbations at the interface (Chapter 3 and [219, 241, 305]). To determine the
importance of the PCNA interface in these DNA replication and repair processes, Todd
Washington generated several random mutations at the subunit interface using site-
directed mutagenesis with primers containing randomized nucleotides at each of the
individual codons. The mutations used for the studies presented herein are shown in
Figure 4.1. To examine how these mutant proteins affect the function of PCNA in TLS
and MMR in vivo, Christine and Viana carried out cell survival and mutagenesis studies
with yeast cells expressing PCNA containing each of these individual interface
mutations. PCNA containing the K164R mutation was used as a negative control in these
studies because it cannot be ubiquitylated at residue 164 and is therefore defective in
promoting TLS. As shown in Table 4.1, column I, several of these interface mutants
displayed normal cell growth, while four (the S177G, G178M, G178L, and V180A
substitutions) grew slower than wild-type PCNA. Interestingly, all PCNA mutants
generated were more sensitive to UV exposure than wild-type PCNA (Table 4.1, column
II).
They next assayed the PCNA interface mutants for induced mutations after UV
exposure (Table 4.1, column III). In this assay, cells expressing the mutant PCNA
proteins were plated on synthetic complete medium lacking arginine and containing
canavanine to determine the frequency of CAN1S to can1
r forward mutations. Only three
substitutions in PCNA (S177G, G178M, and V180A) had normal levels of induced
mutagenesis, suggesting that all other substitutions are defective in TLS in vivo. Lastly,
149
to examine the effect of these PCNA interface mutants on MMR, they performed a
spontaneous mutagenesis assay in which mutagenesis in cells was measured in the
absence of any DNA-damaging agents (Table 4.1, column IV). Results showed that the
E113G, Y114A, G178S, G178M, and V180A mutant PCNA proteins have increased
rates of spontaneous mutations and are therefore likely defective in MMR. Overall, the
genetic studies suggest that these PCNA interface mutant proteins have differing effects
on TLS and MMR.
From these in vivo results, we identified five PCNA interface mutants that
encompass an array of phenotypes and span the β-strand βD2 to isolate and use as a
representative sample for in vitro assays. All subsequent biochemical studies were
performed with these five PCNA interface mutant proteins – S177G, G178S, S179T,
V180A, and I181R.
Trimer stability of the PCNA interface mutant proteins.
Since I have observed reduced trimer stability with several other mutant PCNA
proteins that disrupt the subunit interface, I examined the ability of the five PCNA
interface mutant proteins to form trimers. Using non-denaturing polyacrylamide gel
electrophoresis (PAGE), I showed that these mutant proteins have differing trimer
stabilities (Figure 4.2). Wild-type PCNA and the S179T and I181R mutant PCNA
proteins formed stable trimers under all concentrations tested (0.1 to 1.0 mg/ml). By
contrast, the G178S mutant PCNA protein was a monomer under all concentrations
tested. Interestingly, the S177G and V180A mutant proteins were less stable than wild-
type PCNA, but formed trimers more readily than the G178S mutant PCNA protein. At
all concentrations, the S177G mutant protein bands were mostly trimeric and streaking.
This suggests that, although this protein is generally a trimer, it is slightly less stable than
wild-type PCNA. In comparison, the oligomeric form of the V180A mutant PCNA
protein ranged from monomeric to trimeric as protein concentration increased. This
150
shows that the V180A mutant protein is capable of forming trimers at higher
concentrations, but is even less stable than the S177G mutant PCNA protein. From these
results, I suggest that the trimer stabilities of these PCNA interface mutant proteins span
a wide range as such: G178S < V180A < S177G < S179T, I181R, and wild-type PCNA,
where the G178S mutant PCNA protein is the least stable, and wild-type PCNA and the
S179T and I181R mutant PCNA proteins are the most stable.
In order to further assess PCNA trimer stability, Christine and Viana analyzed the
proteins by size exclusion chromatography. When loaded onto the size exclusion column
at a high concentration (10 mg/ml), the wild-type PCNA protein and the S179T and
I181R mutant proteins eluted in a narrow peak as expected for the 90-kDa trimer (Figure
4.3). When the S177G and V180A mutant proteins were loaded at a high concentration
(10 mg/ml), they eluted in a broad peak, likely corresponding to a mixture of trimeric and
non-trimeric species (Figure 4.3). The G178S mutant PCNA protein has yet to be
evaluated using size exclusion chromatography, but its elution profile is expected to be
similar to or even more broad than those of the S177G and V180A mutant PCNA
proteins. Taken together, both the native PAGE and size exclusion chromatography
results show that the S177G and V180A mutant PCNA proteins (and expectedly the
G178S mutant PCNA protein) are less stable than the S179T and I181R mutant proteins,
which form stable trimers like the wild-type PCNA protein.
Impact of the PCNA interface mutant proteins
on the activity of pol η.
In order to test the ability of the mutant PCNA proteins to stimulate the catalytic
activity of the non-classical polymerase pol η, I used running start experiments to
measure DNA synthesis by pol η on a non-damaged template in the presence of each
PCNA interface mutant protein (Figure 4.4A). Under these conditions, the presence of
wild-type PCNA stimulated DNA synthesis by pol η more than in its absence. Similar
151
stimulation was also observed in the presence of both the S179T and I181R mutant
PCNA proteins. In contrast, the G178S and V180A mutant PCNA proteins did not
stimulate the activity of pol η beyond that of the polymerase itself, whereas the S177G
mutant PCNA protein may show an intermediate stimulation of pol η that is above that of
pol η alone and less than that seen with wild-type PCNA. Therefore, these five PCNA
interface mutant proteins have varying degrees of stimulation of pol η on non-damaged
DNA.
To determine the ability of these mutant PCNA proteins to stimulate TLS by pol
η, I measured DNA synthesis by pol η on a DNA substrate containing an abasic site
located at the sixth position in the template following the primer terminus (Figure 4.4B).
In the absence of PCNA, pol η had little activity opposite the templating abasic site.
Similar to synthesis on non-damaged DNA, pol η’s activity to incorporate nucleotides
opposite the abasic site was stimulated in the presence of wild-type PCNA and the S179T
and V180A mutant PCNA proteins, with 1.3 fold, 1.7 fold, and 1.6 fold stimulation above
that seen with no PCNA, respectively. In the presence of the S177G, G178S, and V180A
mutant PCNA proteins, however, little to no stimulation was observed (0.9 to 1.1 fold for
each compared to pol η alone). Thus, the PCNA interface mutant proteins have differing
effects on the process of TLS by pol η. Altogether, these studies with pol η suggest that
the five mutations at the subunit interface of PCNA exhibit a range of abilities to
stimulate the activity of pol η, where the S177G, G178S, and V180A mutant proteins
show little to no stimulation and the S179T and I181R mutant proteins behave like wild-
type PCNA.
Impact of the PCNA interface mutant proteins
on the activity of pol δ.
Classical pol is responsible for lagging strand synthesis during normal DNA
replication and is also involved in base excision repair, nucleotide excision repair,
152
mismatch repair, and double strand break repair [244-247]. To examine the effects of the
five PCNA interface mutant proteins on the activity of pol δ, I carried out running start
experiments with pol δ on non-damaged DNA in the absence and presence of wild-type
PCNA and each of the mutant proteins (Figure 4.5). The PCNA interface mutant proteins
stimulated DNA synthesis by pol δ to differing extents. In the presence of either the
wild-type PCNA protein, the S179T mutant protein, or the I181R mutant protein,
approximately 4.7, 4.9, or 5.6 fold more full-length runoff products formed, respectively,
compared to in the absence of any PCNA. As observed with DNA synthesis by pol η,
neither of the G178S and V180A mutant PCNA proteins were able to stimulate pol δ
activity on non-damaged DNA past that of the polymerase alone. The amount of full
length product formed by pol δ in the presence of either of these mutant proteins was
~1.1 fold of the amount formed in their absence. Interestingly, the S177G did stimulate
pol δ’s activity (2 fold over the polymerase alone), but to a lesser extent than in the
presence of wild-type PCNA.
In addition to non-damaged DNA, pol δ is capable of bypassing abasic sites. To
determine if the PCNA interface mutant proteins affect TLS by pol δ, I examined DNA
synthesis by the classical polymerase on a DNA template containing an abasic site
(Figure 4.6). Approximately 2.4, 2.1, or 2.7 fold more extension products were observed
opposite the abasic site in the presence of the wild-type PCNA protein, the S179T mutant
PCNA protein, and the I181R mutant PCNA protein than in their absence, respectively.
Furthermore, ~4.8, 4.0, and 4.8 fold more full-length runoff products were formed in the
presence of the three PCNA proteins, respectively. In contrast, both of the G178S and
V180A mutant PCNA proteins failed to stimulate pol δ’s activity opposite the abasic site,
with ~0.9 fold pol δ activity seen in their presence compared to pol δ alone. The amount
of full-length runoff products formed in the presence of these two mutant PCNA proteins
was similar to this, as only ~0.8 and 0.9 fold extension was observed for the G178S and
V180A mutant PCNA proteins, respectively. In contrast to these results, in the presence
153
of the S177G mutant PCNA protein, there were ~1.3 fold more extension products
resulting from incorporation opposite the abasic site and ~1.4 fold more full-length runoff
products than in the absence of PCNA. Thus, the PCNA interface mutant proteins have
differing effects on DNA synthesis on an abasic-containing DNA template by pol δ.
Altogether, the running start experiments with pol δ suggest that five mutations at the
subunit interface of PCNA exhibit a range of abilities to stimulate the activity of pol δ,
where the G178S and V180A mutant proteins show no stimulation, the S179T and I181R
mutant proteins behave like wild-type PCNA, and the S177G mutant protein shows an
intermediate stimulation.
Structures of the PCNA interface mutant proteins.
Experiments to determine the X-ray crystal structures of each of the five PCNA
interface mutants have been underway by Christine Kondratick, Viana Nguyen, and Kyle
Powers. So far, the structures of the S177G, G178S, S179T, and V180A mutant PCNA
proteins have been solved and fully refined. The major changes observed between these
mutant proteins and the wild-type PCNA protein are generally located at the subunit
interface. As described in Chapter 2, the G178S substitution causes significant shearing
of the interface, in which only three of the original seven hydrogen bonds are maintained
between the β-strands that constitute the PCNA interface (Table 4.2). This same shearing
of the interface is also seen in the case of the S177G, S179T, and V180A mutant PCNA
proteins, however to lesser extents (Table 4.2). In the case of the S177G mutant protein,
three hydrogen bonds out of the original seven are broken. Similar to that observed with
the protein stability assays, the S179T mutation causes less distortion of the subunit
interface than either the G178S and S177G substitutions, i.e. only two of the original
seven hydrogen bonds are broken in this mutant PCNA protein. Therefore, the S177G
and G178S amino acid substitutions perturb the subunit interface of PCNA to varying
154
degrees and are likely the source of the differences in trimer stabilities and in vitro
functions of each mutant PCNA protein.
Contrary to what was expected, the V180A substitution only caused two hydrogen
bonds to be disrupted at the interface. However, upon further inspection, I noticed that
there were large perturbations of the V180A mutant PCNA protein outside of the subunit
interface. While the V180A substitution is positioned in domain B of PCNA, the largest
changes in structure are observed in domain A (Figure 4.7A). Figure 4.7B shows that the
β-strands in domain A (βA1, βG1, βH1, and βI1) are shifted significantly from the wild-
type PCNA structure. For instance, backbone residue shifts of 0.5 Å, 0.9 Å, and 0.8 Å
are observed between the mutant and wild-type PCNA for strands βI1, βH1, and βG1,
respectively. In the case of βA1, these shifts are as large as 3.0 Å. Together, the
propagated disruptions in these β-strands along with the small changes at the subunit
interface likely account for the reduced trimer stability seen with the V180A mutant
PCNA protein. Determination of the structures of the I181R mutant PCNA protein is still
underway, but because it behaves like wild-type PCNA in every assay performed, I
predict that the I181R mutant PCNA protein will have the least significant changes of all
of these mutant PCNA proteins.
Discussion
Studies with the five PCNA interface mutant proteins presented here suggest that
the subunit interface of PCNA is very dynamic and that small changes at the interface can
cause drastically different effects on TLS and MMR. Table 4.3 summarizes all of the
genetic and biochemical results characterizing the five mutant PCNA proteins. My
biochemical studies with the five mutant PCNA proteins suggest that there is a strong
correlation between PCNA trimer stability and TLS in vitro. Based on the structures of
these five mutant proteins, this trimer instability usually arises due to instability at the
155
interface, i.e. number of hydrogen bonds disrupted between the two β-strands that
constitute the interface. For instance, the S179T mutant PCNA protein behaves like
wild-type PCNA in stimulating both pol δ and pol η activity opposite damaged DNA and
has at least five of the original seven hydrogen bonds observed in wild-type PCNA. In
contrast, the G178S mutant PCNA protein is completely defective in stimulating TLS by
pol δ or pol η and only preserves three hydrogen bonds, resulting in a severely sheared
subunit interface. The S177G mutant PCNA protein, however, shows an intermediate
phenotype for stimulating DNA synthesis by pol δ and pol η on damaged DNA and
maintains four hydrogen bonds at the interface. The structure of the I181R mutant PCNA
protein has not yet been determined, but I expect that it will contain at least five hydrogen
bonds at the subunit interface due to its high trimer stability and its ability to stimulate
both pol δ and pol η on damaged and non-damaged DNA templates. These results are
consistent with the studies described in Chapters 2 and 3 in which the reduced trimer
stability of the E113G and C81R mutant PCNA proteins at the PCNA subunit interface
resulted in a decreased ability to stimulate DNA synthesis by polymerases.
The integrity of the PCNA subunit interface is clearly important for PCNA
function in vitro. However, the structure of the V180A mutant protein shows that amino
acid substitutions at the interface can also cause perturbations outside of the interface
itself. This substitution causes structural conformational shifts of up to 3.0 Å, which are
propagated up to the N-terminus of the mutant PCNA protein. These residues are
actually positioned in trans to the mutation site and together make up four β-strands in
domain A. Therefore, the defect in stimulating TLS by pol δ and pol η observed in the
presence of the V180A mutant PCNA protein is likely due to these large structural
changes. Together, my results suggest that, in addition to the PCNA subunit interface,
the first ten residues of PCNA are crucial for proper PCNA function in vitro.
The correlation between the crystal structures of the five PCNA interface mutant
proteins and their functions in vitro is quite evident. This correlation, however, is less
156
well-defined with their functions in vivo, as many more factors are likely involved in TLS
and MMR in the cell. Probably the clearest case is of the G178S mutant PCNA protein.
The subunit interface is significantly altered compared to the interface of the wild-type
protein, which is likely the cause of the defects seen in both the TLS and MMR in the
genetic assays. Since the structure of the I181R mutant PCNA protein has not been
determined, it is unclear how this mutation causes reduced UV-induced mutagenesis in
vivo despite the high trimer stability and ability to stimulate pol δ and pol η observed with
this mutant protein in vitro. It is plausible, though, that this structure is similar to the
structure of the S179T mutant PCNA protein, as their phenotypes are almost identical.
Based on conclusions drawn from Chapter 2, it is likely that the small disruption at the
interface seen in the S179T mutant PCNA protein causes a defect in TLS in vivo, but that
the disruption is not large enough to observe significant defects in vitro. The correlation
between the structure and in vivo phenotype of the S177G mutant PCNA protein is
unclear, as the structure of this mutant protein suggests that it would cause defect in TLS
or MMR. More in vivo studies will be necessary with this mutant protein to elucidate the
relationship between its structure and function.
The case of the V180A mutation PCNA protein is unique in that the largest
structural perturbations caused by this mutation is actually within domain A of the
protein instead of at the subunit interface. This causes defects in both growth rate and
MMR but not TLS in vivo. One possible explanation for the elevated mutation rate and
reduced growth rate is that the structural changes produced by the V180A mutation
interfere with the physical interaction between PCNA and pol δ. Although the integrity
of the interface of PCNA is not required for PCNA-polymerase interactions (see Chapter
2), the integrity of the β-strands of domain A near the inter-domain cleft may be required
for proper interactions with pol δ. Another possible explanation for this is that these
structural changes disrupt the functional interaction between PCNA and pol δ. The large
decrease in stability in PCNA may interfere with the activity of the exonuclease domain
157
or the active site in pol δ. In support of this, exonuclease-deficient S. cerevisiae pol δ
generated base substitution errors at rates 60 fold higher than those produced by wild type
pol δ in in vivo [87]. Therefore, disruption of the functional or physical interaction
between PCNA and pol δ caused by the introduction of the V180A substitution may
result in slow cell growth and defective MMR but not TLS in vivo. This may also be the
case with the S177G mutant PCNA protein since it also shows a slow growth phenotype.
More genetic and binding studies will be necessary to distinguish between these
possibilities.
158
Figure 4.1. The amino acid residues that comprise the PCNA subunit interface.
Residues located on β-strand βI1 are shown surrounded by green and residues located on
β-strand βD2 are shown surrounded by purple. Amino acid substitutions shown in both
grey and red were used in genetic studies, while the amino acid substitutions shown in
red were used for biochemical assays.
159
Table 4.1. Summary of genetic studies with the PCNA interface mutant proteins.
I II III IV
Cell growth UV survival UV-induced mutagenesis
Spontaneous mutagenesis
WT Normal Normal Normal Normal
K164R Normal Sensitive Reduced Normal
E113G Normal Sensitive Reduced Elevated
Y114F Normal Intermediate Reduced Normal
Y114A Normal Sensitive Intermediate Elevated
S115N Normal Intermediate Reduced Normal
S177G Slow Intermediate Normal Normal
G178S Normal Sensitive Reduced Elevated
G178M Slow Intermediate Normal Elevated
G178L Slow Sensitive Reduced Normal
S179R Normal Intermediate Reduced Normal
S179T Normal Intermediate Reduced Normal
V180A Slow Sensitive Normal Elevated
I181R Normal Sensitive Reduced Normal
Reduced UV-induced mutagenesis indicates a defect in TLS. Elevated spontaneous mutagenesis indicates a defect in MMR. (Data from Christine Kondratick and Viana Ngyuen.)
160
Figure 4.2. Analysis of the wild-type and mutant PCNA proteins by native gel
electrophoresis. Coomassie stained non-denaturing polyacrylamide gradient gel (4 to
20%) in which solutions of the wild-type and mutant PCNA proteins (1.0 to 1.0 mg/ml)
were run. The position of the PCNA trimers and monomers are shown.
161
Figure 4.3. Analysis of the PCNA interface mutant proteins by size exclusion
chromatography. The elution profiles of a size exclusion chromatography column in
which a solution of the wild-type PCNA protein or the S177G, S179T, V180A, or I181R
mutant PCNA protein (10 mg/ml) was run. (Data from Christine Kondratick and Viana
Ngyuen.)
162
Figure 4.4. DNA synthesis by pol η in the presence of the PCNA mutant proteins.
(A) An autoradiogram of the products of pol η-catalyzed DNA synthesis on a non-
damaged DNA substrate in the presence of no PCNA, the wild-type PCNA protein, and
the S177G, G178S, S179T, V180A, and I181R mutant PCNA proteins. (B) An
autoradiogram of the products of pol η-catalyzed DNA synthesis on an abasic site-
containing DNA substrate in the presence of no PCNA, the wild-type PCNA protein, and
the S177G, G178S, S179T, V180A, and I181R mutant PCNA proteins. The gel band
representing extension products 6 nt. in length, which corresponds to incorporation
opposite the abasic site, is indicated by an X.
163
Figure 4.5. DNA synthesis by pol on a non-damaged template in the presence of
the PCNA mutant proteins. (A) An autoradiogram of the products of pol -catalyzed
DNA synthesis on a non-damaged DNA substrate in the presence of no PCNA, the wild-
type PCNA protein, and the S177G, G178S, S179T, V180A, and I181R mutant PCNA
proteins after a 1 minute reaction time. The position of the fully extended, runoff product
is indicated with an arrow. (B) An autoradiogram of the products of pol -catalyzed DNA
synthesis on a non-damaged DNA substrate in the presence of no PCNA, the wild-type
PCNA protein, and the S177G, G178S, S179T, V180A, and I181R mutant PCNA
proteins after a 2 minute reaction time. The position of the fully extended, runoff product
is indicated with an arrow
164
Figure 4.6. DNA synthesis by pol on a template containing an abasic site in the
presence of the PCNA mutant proteins. (A) An autoradiogram of the products of pol -
catalyzed DNA synthesis on an abasic site-containing DNA substrate in the presence of
no PCNA, the wild-type PCNA protein, and the S177G, G178S, S179T, V180A, and
I181R mutant PCNA proteins after a 1 minute reaction time. The position of the abasic
site is indicated by an X, and the position of the fully extended, runoff product is
indicated with an arrow. (B) An autoradiogram of the products of pol -catalyzed DNA
synthesis on an abasic site-containing DNA substrate in the presence of no PCNA, the
wild-type PCNA protein, and the S177G, G178S, S179T, V180A, and I181R mutant
PCNA proteins after a 2 minute reaction time. The position of the abasic site is indicated
by an X, and the position of the fully extended, runoff product is indicated with an arrow.
165
Table 4.2. Distances between potential hydrogen bond donor and acceptor atoms at
Figure 4.7. Structure of the V180A mutant PCNA protein. (A) One subunit of the
V180A mutant PCNA protein (green) overlayed with wild-type PCNA (cyan). The
location of the V180A substitution and domains A and B are indicated. (B) Close up of
the region affected by the V180A substitution in domain A. β-strands disrupted by the
V180A substitution are indicated. The mutant PCNA protein is shown in green and the
wild-type PCNA protein is shown in cyan.
167
Table 4.3. Summary of in vivo and in vitro studies with the PCNA interface mutant
proteins.
Reduced UV-induced mutagenesis indicates a defect in TLS. Elevated spontaneous mutagenesis indicates a defect in MMR. *V180A contains structural changes in addition to at the subunit interface. ND – not determined.
control, sister chromatid cohesion [155, 199, 200, 257, 258]. Therefore, it is crucial that
PCNA functions properly and efficiently to regulate the position and function of proteins
involved in all of these pathways.
The means by which PCNA orchestrates control of so many proteins is currently
an active area of research. So far, several modes of regulation have been proposed that
are likely to help facilitate sequential action of PCNA-interacting proteins. The first and
perhaps the most obvious mode of regulation is competition by these proteins. For
205
instance, p21 has the highest affinity for PCNA than any other PIP motif-containing
protein [201]. Since p21 is a cell cycle regulator, it is essential that this protein have the
ability to displace any other protein from PCNA at any moment. Along these same lines
is the idea that post-translational modifications of PCNA regulate the binding and/or
removal of specific interacting proteins. Several studies have shown that the
monoubiquitylation of PCNA recruits non-classical polymerases that contain ubiquitin-
binding motifs, whereas sumoylation of PCNA recruits the anti-recombinogenic helicase
Srs2 to the DNA. Similarly, several PCNA-binding factors are known to be post-
translationally modified themselves. For example, FEN1 is phosphorylated by the
CDK2-PCNA complex, which leads its dissociation from PCNA [314]. In addition, the
observation that many non-classical polymerases are capable of ubiquitylation has led to
the notion that this event acts as a switch to prevent re-association of these enzymes after
TLS is completed.
PCNA is a homotrimer and therefore has the capability of binding up to three PIP
motif-containing proteins simultaneously. This would allow the ordered binding of
factors involved in multi-protein processes. Thus one protein could be recruited to one
subunit of PCNA, which would stimulate binding of a second factor to another subunit,
and so on. Indeed, studies show that this is likely the case with Sulfolobus solfataricus
PCNA, in which FEN1, DNA ligase, and a DNA polymerase can bind one PCNA
heterotrimer simultaneously [235].
After all of these factors bind, how does PCNA regulate the dissociation of these
proteins? As with loading on to DNA substrates, PCNA is also unloaded by RFC. This
would act to remove all PCNA-interacting partners from the DNA. It has also been
suggested that sumoylation of PCNA at K127 causes dissociation of proteins from
PCNA, although this pathway is not well understood [315]. Lastly, PCNA-protein
complexes can be permanently eliminated by PCNA degradation through ubiquitylation.
This ubiquitylation is separate from the RAD6 pathway and causes reduced DNA
206
synthesis and defects in mismatch repair, but specific details of this pathway remain to be
elucidated.
My work in Chapter 5 shows that the interaction between PCNA and pol η is
dependent on the unstructured CTR of pol η. With all of these complicated mechanisms
to carry out in a sequential order, it is likely that PCNA often interacts with its partners in
a transient and dynamic manner. Since unstructured regions of proteins provide a great
deal of flexibility for protein-protein interactions, it is conceivable that many of the
proteins that interact with PCNA contain one or more of these intrinsically disordered
regions. It has been estimated that nearly a third of eukaryotic proteins and a half of
mammalian proteins are partially or fully disordered [309, 310]. These disordered regions
are often involved in multiple interactions with several protein partners [309, 310, 316].
Another example of this is the interaction between sumoylated PCNA and the Srs2
helicase, which contains a large unstructured region that possesses ordered motifs that
bind SUMO and PCNA independently [308]. There are many ways in which PCNA
regulates cellular processes, and it is probable that many more exist. It is likely, though,
that intrinsically disordered proteins play a major role, and future studies will be focused
on the novel modes of interaction that exist through these unstructured gold mines.
207
Figure 6.1. Positions of all PCNA substitutions used for studies. (A) Front view and
(B) side view of the PCNA trimer with individual subunits colored in pink, green, and
blue. The location of each of the eight residues that were mutated in my studies are
indicated in relation to the front and back faces, the IDCL, and the domains of each
subunit of PCNA.
208
Figure 6.2. Close-up of positions of all PCNA substitutions used for studies. Close-
up view of a PCNA monomer with the eight individual residues that were mutated in my
studies indicated in relation to the front and back faces, the IDCL, and the domains of
PCNA.
209
Figure 6.3. Potential positions of the ubiquitin moiety on PCNA and possible roles
of these positions during TLS. PCNA is shown in purple, ubiquitin is shown in yellow,
and the polymerase is shown in green. The possible roles of each position during TLS
and the location of the E113G substitution are indicated.
210
Figure 6.4. SAXS analysis of the E113G mutant Ub-PCNA protein. (A) Comparison
of SAXS curves of the wild-type Ub-PCNA and the E113G mutant Ub-PCNA proteins.
(B) Close-up of the comparison of SAXS curves of the wild-type Ub-PCNA and the
E113G mutant Ub-PCNA proteins at low angles. (Data from Susan Tsutakawa.)
211
Figure 6.5. Non-denaturing gel electrophoresis of wild-type PCNA. Trimeric,
hexameric, and higher-order oligomeric states of wild-type PCNA are shown at
increasing protein concentrations.
212
Figure 6.6. B-factors of the PCNA interface. The dynamics of each backbone atom in
wild-type PCNA is represented according to size. The image was rendered using B-
factor putty as implemented in pymol. Domain A is shown in blue and domain B is
shown in red. The β-strands that constitute the interface and the position of loop J are
indicated.
213
REFERENCES
1. Devlin, T.M., ed. Textbook of Biochemistry: with Clinical Correlations. 6 ed. 2006, John Wiley & Sons, Inc.: Hoboken, NJ.
2. Matsunaga, T., K. Hieda, and O. Nikaido, WAVELENGTH DEPENDENT FORMATION OF THYMINE DIMERS AND (6-4)PHOTOPRODUCTS IN DNA BY MONOCHROMATIC ULTRAVIOLET-LIGHT RANGING FROM 150 TO 365 NM. Photochemistry and Photobiology, 1991. 54(3): p. 403-410.
3. Karran, P., DNA double strand break repair in mammalian cells. Current Opinion in Genetics & Development, 2000. 10(2): p. 144-150.
4. Beckman, K.B. and B.N. Ames, Oxidative decay of DNA. Journal of Biological Chemistry, 1997. 272(32): p. 19633-19636.
5. Helbock, H.J., et al., DNA oxidation matters: The HPLC-electrochemical detection assay of 8-oxo-deoxyguanosine and 8-oxo-guanine. Proceedings of the National Academy of Sciences of the United States of America, 1998. 95(1): p. 288-293.
6. Lindahl, T. and D.E. Barnes, Repair of endogenous DNA damage. Cold Spring Harbor Symposia on Quantitative Biology, 2000. 65: p. 127-133.
7. Lindahl, T., INSTABILITY AND DECAY OF THE PRIMARY STRUCTURE OF DNA. Nature, 1993. 362(6422): p. 709-715.
8. Lucas-Lledo, J.I. and M. Lynch, Evolution of Mutation Rates: Phylogenomic Analysis of the Photolyase/Cryptochrome Family. Molecular Biology and Evolution, 2009. 26(5): p. 1143-1153.
9. Parikh, S.S., et al., Base excision repair initiation revealed by crystal structures and binding kinetics of human uracil-DNA glycosylase with DNA. Embo Journal, 1998. 17(17): p. 5214-5226.
10. Guillet, M. and S. Boiteux, Origin of endogenous DNA abasic sites in Saccharomyces cerevisiae. Molecular and Cellular Biology, 2003. 23(22): p. 8386-8394.
11. Wilson, D.M. and D. Barsky, The major human abasic endonuclease: formation, consequences and repair of abasic lesions in DNA. Mutation Research-DNA Repair, 2001. 485(4): p. 283-307.
12. Wilson, S.H., Mammalian base excision repair and DNA polymerase beta. Mutation Research-DNA Repair, 1998. 407(3): p. 203-215.
13. Caldecott, K.W., XRCC1 and DNA strand break repair. DNA Repair, 2003. 2(9): p. 955-969.
14. Sancar, A., DNA excision repair. Annual Review of Biochemistry, 1996. 65: p. 43-81.
214
15. Fousteri, M. and L.H. Mullenders, Transcription-coupled nucleotide excision repair in mammalian cells: molecular mechanisms and biological effects. Cell Research, 2008. 18(1): p. 73-84.
16. Downs, J.A., Chromatin structure and DNA double-strand break responses in cancer progression and therapy. Oncogene, 2007. 26(56): p. 7765-7772.
17. Ward, J.F., THE YIELD OF DNA DOUBLE-STRAND BREAKS PRODUCED INTRACELLULARLY BY IONIZING-RADIATION - A REVIEW. International Journal of Radiation Biology, 1990. 57(6): p. 1141-1150.
18. Song, X.Y., et al., Phosphorylation of the SQ H2A.X motif is required for proper meiosis and mitosis in Tetrahymena thermophila. Molecular and Cellular Biology, 2007. 27(7): p. 2648-2660.
19. van Attikum, H. and S.M. Gasser, The histone code at DNA breaks: A guide to repair? Nature Reviews Molecular Cell Biology, 2005. 6(10): p. 757-765.
20. Symington, L.S., Role of RAD52 epistasis group genes in homologous recombination and double-strand break repair. Microbiology and Molecular Biology Reviews, 2002. 66(4): p. 630-+.
21. Espejel, S., et al., Mammalian Ku86 mediates chromosomal fusions and apoptosis caused by critically short telomeres. Embo Journal, 2002. 21(9): p. 2207-2219.
22. Iyer, R.R., et al., DNA mismatch repair: Functions and mechanisms. Chemical Reviews, 2006. 106(2): p. 302-323.
23. Hsieh, P. and K. Yamane, DNA mismatch repair: Molecular mechanism, cancer, and ageing. Mechanisms of Ageing and Development, 2008. 129(7-8): p. 391-407.
24. Kolodner, R.D. and G.T. Marsischky, Eukaryotic DNA mismatch repair. Current Opinion in Genetics & Development, 1999. 9(1): p. 89-96.
25. Modrich, P. and R. Lahue, Mismatch repair in replication fidelity, genetic recombination, and cancer biology. Annual Review of Biochemistry, 1996. 65: p. 101-133.
26. Lynch, H.T. and A. de la Chapelle, Genetic susceptibility to non-polyposis colorectal cancer. Journal of Medical Genetics, 1999. 36(11): p. 801-818.
27. Ban, C. and W. Yang, Crystal structure and ATPase activity of MutL: Implications for DNA repair and mutagenesis. Cell, 1998. 95(4): p. 541-552.
28. Junop, M.S., et al., Composite active site of an ABC ATPase: MutS uses ATP to verify mismatch recognition and authorize DNA repair. Molecular Cell, 2001. 7(1): p. 1-12.
29. Dao, V. and P. Modrich, Mismatch-, MutS-, MutL-, and helicase II-dependent unwinding from the single-strand break of an incised heteroduplex. Journal of Biological Chemistry, 1998. 273(15): p. 9202-9207.
215
30. Li, F., et al., Escherichia coli mismatch repair protein MutL interacts with the clamp loader subunits of DNA polymerase III. Mutation Research-Fundamental and Molecular Mechanisms of Mutagenesis, 2008. 637(1-2): p. 101-110.
31. Lopez de Saro, F.J., et al., The beta sliding clamp binds to multiple sites within MutL and MutS. The Journal of biological chemistry, 2006. 281(20): p. 14340-9.
32. Palombo, F., et al., hMutS beta, a heterodimer of hMSH2 and hMSH3, binds to insertion/deletion leaps in DNA. Current Biology, 1996. 6(9): p. 1181-1184.
33. Drummond, J.T., et al., ISOLATION OF AN HMSH2-P160 HETERODIMER THAT RESTORES DNA MISMATCH REPAIR TO TUMOR-CELLS. Science, 1995. 268(5219): p. 1909-1912.
34. Acharya, S., et al., hMSH2 forms specific mispair-binding complexes with hMSH3 and hMSH6. Proceedings of the National Academy of Sciences of the United States of America, 1996. 93(24): p. 13629-13634.
35. Flores-Rozas, H. and R.D. Kolodner, The Saccharomyces cerevisiae MLH3 gene functions in MSH3-dependent suppression of frameshift mutations. Proceedings of the National Academy of Sciences of the United States of America, 1998. 95(21): p. 12404-12409.
36. Harfe, B.D., B.K. Minesinger, and S. Jinks-Robertson, Discrete in vivo roles for the MutL homologs Mlh2p and Mlh3p in the removal of frameshift intermediates in budding yeast. Current Biology, 2000. 10(3): p. 145-148.
37. Habraken, Y., et al., Enhancement of MSH2-MSH3-mediated mismatch recognition by the yeast MLH1-PMS1 complex. Current Biology, 1997. 7(10): p. 790-793.
38. Prolla, T.A., D.M. Christie, and R.M. Liskay, DUAL REQUIREMENT IN YEAST DNA MISMATCH REPAIR FOR MLH1 AND PMS1, 2 HOMOLOGS OF THE BACTERIAL MUTL GENE. Molecular and Cellular Biology, 1994. 14(1): p. 407-415.
39. Genschel, J., L.R. Bazemore, and P. Modrich, Human exonuclease I is required for 5 ' and 3 ' mismatch repair. Journal of Biological Chemistry, 2002. 277(15): p. 13302-13311.
40. Schmutte, C., et al., Human exonuclease I interacts with the mismatch repair protein hMSH21. Cancer Research, 1998. 58(20): p. 4537-4542.
41. Tishkoff, D.X., et al., Identification of a human gene encoding a homologue of Saccharomyces cerevisiae EXO1, an exonuclease implicated in mismatch repair and recombination. Cancer Research, 1998. 58(22): p. 5027-5031.
42. Tishkoff, D.X., et al., Identification and characterization of Saccharomyces cerevisiae EXO1, a gene encoding an exonuclease that interacts with MSH2. Proceedings of the National Academy of Sciences of the United States of America, 1997. 94(14): p. 7487-7492.
216
43. Longley, M.J., A.J. Pierce, and P. Modrich, DNA polymerase delta is required for human mismatch repair in vitro. Journal of Biological Chemistry, 1997. 272(16): p. 10917-10921.
44. Lin, Y.L., et al., The evolutionarily conserved zinc finger motif in the largest subunit of human replication protein a is required for DNA replication and mismatch repair but not for nucleotide excision repair. Journal of Biological Chemistry, 1998. 273(3): p. 1453-1461.
45. Ramilo, C., et al., Partial reconstitution of human DNA mismatch repair in vitro: Characterization of the role of human replication protein A. Molecular and Cellular Biology, 2002. 22(7): p. 2037-2046.
46. Kadyrov, F.A., et al., Endonucleolytic function of MutL alpha in human mismatch repair. Cell, 2006. 126(2): p. 297-308.
47. Dzantiev, L., et al., Defined human system that supports bidirectional mismatch-provoked excision. Molecular Cell, 2004. 15(1): p. 31-41.
48. Yuan, F.H., et al., Evidence for involvement of HMGB1 protein in human DNA mismatch repair. Journal of Biological Chemistry, 2004. 279(20): p. 20935-20940.
49. Zhang, Y.B., et al., Reconstitution of 5 '-directed human mismatch repair in a purified system. Cell, 2005. 122(5): p. 693-705.
50. Pluciennik, A., et al., PCNA function in the activation and strand direction of MutL alpha endonuclease in mismatch repair. Proceedings of the National Academy of Sciences of the United States of America, 2010. 107(37): p. 16066-16071.
51. Umar, A., et al., Requirement for PCNA in DNA mismatch repair at a step preceding DNA resynthesis. Cell, 1996. 87(1): p. 65-73.
52. Gu, L.Y., et al., ATP-dependent interaction of human mismatch repair proteins and dual role of PCNA in mismatch repair. Nucleic Acids Research, 1998. 26(5): p. 1173-1178.
53. Clark, A.B., et al., Functional interaction of proliferating cell nuclear antigen with MSH2-MSH6 and MSH2-MSH3 complexes. Journal of Biological Chemistry, 2000. 275(47): p. 36498-36501.
54. Flores-Rozas, H., D. Clark, and R.D. Kolodner, Proliferating cell nuclear antigen and Msh2p-Msh6p interact to form an active mispair recognition complex. Nature Genetics, 2000. 26(3): p. 375-378.
55. Kleczkowska, H.E., et al., hMSH3 and hMSH6 interact with PCNA and colocalize with it to replication foci. Genes & Development, 2001. 15(6): p. 724-736.
56. Lau, P.J. and R.D. Kolodner, Transfer of the MSH2 center dot MSH6 complex from proliferating cell nuclear antigen to mispaired bases in DNA. Journal of Biological Chemistry, 2003. 278(1): p. 14-17.
217
57. Shell, S.S., C.D. Putnam, and R.D. Kolodner, The N terminus of Saccharomyces cerevisiale Msh6 is an unstructured tether to PCNA. Molecular Cell, 2007. 26(4): p. 565-578.
58. Loeb, K.R. and L.A. Loeb, Significance of multiple mutations in cancer. Carcinogenesis, 2000. 21(3): p. 379-385.
59. Loeb, L.A., K.R. Loeb, and J.P. Anderson, Multiple mutations and cancer. Proceedings of the National Academy of Sciences of the United States of America, 2003. 100(3): p. 776-781.
60. Prakash, S., R.E. Johnson, and L. Prakash, Eukaryotic translesion synthesis DNA polymerases: Specificity of structure and function. Annual Review of Biochemistry, 2005. 74: p. 317-353.
61. Guo, C., et al., Y-family DNA polymerases in mammalian cells. Cellular and Molecular Life Sciences, 2009. 66(14): p. 2363-2381.
62. Braithwaite, D.K. and J. Ito, COMPILATION, ALIGNMENT, AND PHYLOGENETIC-RELATIONSHIPS OF DNA-POLYMERASES. Nucleic Acids Research, 1993. 21(4): p. 787-802.
63. Ohmori, H., et al., The Y-family of DNA polymerases. Molecular Cell, 2001. 8(1): p. 7-8.
64. Steitz, T.A., DNA polymerases: Structural diversity and common mechanisms. Journal of Biological Chemistry, 1999. 274(25): p. 17395-17398.
65. Doublie, S., M.R. Sawaya, and T. Ellenberger, An open and closed case for all polymerases. Structure with Folding & Design, 1999. 7(2): p. R31-R35.
66. Rothwell, P.J. and G. Waksman, Structure and mechanism of DNA polymerases. Fibrous Proteins: Muscle and Molecular Motors, 2005. 71: p. 401-+.
67. Brautigam, C.A. and T.A. Steitz, Structural and functional insights provided by crystal structures of DNA polymerases and their substrate complexes. Current Opinion in Structural Biology, 1998. 8(1): p. 54-63.
68. Garg, P. and P.M.J. Burgers, DNA polymerases that propagate the eukaryotic DNA replication fork. Critical Reviews in Biochemistry and Molecular Biology, 2005. 40(2): p. 115-128.
69. Echols, H. and M.F. Goodman, FIDELITY MECHANISMS IN DNA-REPLICATION. Annual Review of Biochemistry, 1991. 60: p. 477-511.
70. Goodman, M.F. and D.K. Fygenson, DNA polymerase fidelity: From genetics toward a biochemical understanding. Genetics, 1998. 148(4): p. 1475-1482.
71. Kunkel, T.A. and R. Bebenek, DNA replication fidelity. Annual Review of Biochemistry, 2000. 69: p. 497-529.
72. Pursell, Z.F., et al., Yeast DNA polymerase epsilon participates in leading-strand DNA replication. Science, 2007. 317(5834): p. 127-130.
218
73. McCulloch, S.D. and T.A. Kunkel, The fidelity of DNA synthesis by eukaryotic replicative and translesion synthesis polymerases. Cell Research, 2008. 18(1): p. 148-161.
74. Swan, M.K., et al., Structural basis of high-fidelity DNA synthesis by yeast DNA polymerase delta. Nature Structural & Molecular Biology, 2009. 16(9): p. 979-U107.
75. Dieckman, L.M., et al., Pre-Steady State Kinetic Studies of the Fidelity of Nucleotide Incorporation by Yeast DNA Polymerase delta. Biochemistry, 2010. 49(34): p. 7344-7350.
76. Washington, M.T., et al., Fidelity and processivity of Saccharomyces cerevisiae DNA polymerase eta. Journal of Biological Chemistry, 1999. 274(52): p. 36835-36838.
77. Washington, M.T., et al., Accuracy of thymine-thymine dimer bypass by Saccharomyces cerevisiae DNA polymerase eta. Proceedings of the National Academy of Sciences of the United States of America, 2000. 97(7): p. 3094-3099.
78. Ng, L., et al., ENZYMOLOGIC MECHANISM OF CALF THYMUS DNA POLYMERASE-DELTA. Journal of Biological Chemistry, 1991. 266(18): p. 11699-11704.
79. Johnson, K.A., Rapid Quench Kinetic-Analysis of Polymerases, Adenosine-Triphosphatases, and Enzyme Intermediates. Enzyme Kinetics and Mechanism, Pt D, 1995. 249: p. 38-61.
80. Johnson, A. and M. O'Donnell, Cellular DNA replicases: Components and dynamics at the replication fork, in Annual Review of Biochemistry. 2005, Annual Reviews: Palo Alto. p. 283-315.
81. Podust, V.N., et al., Reconstitution of human DNA polymerase delta using recombinant baculoviruses - The p12 subunit potentiates DNA polymerizing activity of the four-subunit enzyme. Journal of Biological Chemistry, 2002. 277(6): p. 3894-3901.
82. Fortune, J.M., et al., RPA and PCNA suppress formation of large deletion errors by yeast DNA polymerase delta. Nucleic Acids Research, 2006. 34(16): p. 4335-4341.
83. Zhou, J.Q., et al., Purification and characterization of the catalytic subunit of human DNA polymerase delta expressed in baculovirus-infected insect cells. Journal of Biological Chemistry, 1996. 271(47): p. 29740-29745.
84. Hindges, R. and U. Hubscher, DNA polymerase delta, an essential enzyme for DNA transactions. Biological Chemistry, 1997. 378(5): p. 345-362.
85. Gerik, K.J., et al., Characterization of the two small subunits of Saccharomyces cerevisiae DNA polymerase delta. Journal of Biological Chemistry, 1998. 273(31): p. 19747-19755.
219
86. Hanna, M., et al., Pol32 is required for Pol zeta-dependent translesion synthesis and prevents double-strand breaks at the replication fork. Mutation Research-Fundamental and Molecular Mechanisms of Mutagenesis, 2007. 625(1-2): p. 164-176.
87. Fortune, J.M., et al., Saccharomyces cerevisiae DNA polymerase delta - High fidelity for base substitutions but lower fidelity for single- and multi-base deletions. Journal of Biological Chemistry, 2005. 280(33): p. 29980-29987.
88. Johnson, R.E., S. Prakash, and L. Prakash, Efficient bypass of a thymine-thymine dimer by yeast DNA polymerase, Pol eta. Science, 1999. 283(5404): p. 1001-1004.
89. Haracska, L., et al., Efficient and accurate replication in the presence of 7,8-dihydro-8-oxoguanine by DNA polymerase eta. Nature Genetics, 2000. 25(4): p. 458-461.
90. Washington, M.T., et al., Efficient and error-free replication past a minor-groove N-2-guanine adduct by the sequential action of yeast Rev1 and DNA polymerase zeta. Molecular and Cellular Biology, 2004. 24(16): p. 6900-6906.
91. Wolfle, W.T., et al., Human DNA polymerase iota promotes replication through a ring-closed minor-groove adduct that adopts a syn conformation in DNA. Molecular and Cellular Biology, 2005. 25(19): p. 8748-8754.
92. Pence, M.G., et al., Lesion Bypass of N-2-Ethylguanine by Human DNA Polymerase i. Journal of Biological Chemistry, 2009. 284(3): p. 1732-1740.
93. Ogi, T., et al., Pol kappa protects mammalian cells against the lethal and mutagenic effects of benzo a pyrene. Proceedings of the National Academy of Sciences of the United States of America, 2002. 99(24): p. 15548-15553.
94. Takenaka, K., et al., Involvement of vertebrate Pol kappa in translesion DNA synthesis across DNA monoalkylation damage. Journal of Biological Chemistry, 2006. 281(4): p. 2000-2004.
95. Avkin, S., et al., Quantitative analysis of translesion DNA synthesis across a benzo[a] pyrene-guanine adduct in mammalian cells - The role of DNA polymerase kappa. Journal of Biological Chemistry, 2004. 279(51): p. 53298-53305.
96. Pryor, J.M. and M.T. Washington, Pre-steady state kinetic studies show that an abasic site is a cognate lesion for the yeast Rev1 protein. DNA Repair, 2011. 10(11): p. 1138-1144.
97. Washington, M.T., et al., Efficient and error-free replication past a minor-groove DNA adduct by the sequential action of human DNA polymerases iota and kappa. Molecular and Cellular Biology, 2004. 24(13): p. 5687-5693.
98. Nair, D.T., et al., Protein-template-directed synthesis across an acrolein-derived DNA adduct by yeast rev1 DNA polymerase. Structure, 2008. 16(2): p. 239-245.
220
99. Choi, J.Y. and F.P. Guengerich, Kinetic analysis of translesion synthesis opposite bulky N-2- and O-6-alkylguanine DNA adducts by human DNA polymerase REV1. Journal of Biological Chemistry, 2008. 283(35): p. 23645-23655.
100. Zhang, Y.B., et al., Response of human REV1 to different DNA damage: preferential dCMP insertion opposite the lesion. Nucleic Acids Research, 2002. 30(7): p. 1630-1638.
101. Ohmori, H., et al., SEPARATE ROLES OF STRUCTURED AND UNSTRUCTURED REGIONS OF Y-FAMILY DNA POLYMERASES, in Advances in Protein Chemistry and Structural Biology, Vol 78, A. McPherson, Editor. 2009. p. 99-146.
102. Washington, M.T., et al., Mismatch extension ability of yeast and human DNA polymerase eta. Journal of Biological Chemistry, 2001. 276(3): p. 2263-2266.
103. Haracska, L., S. Prakash, and L. Prakash, Replication past O-6-methylguanine by yeast and human DNA polymerase eta. Molecular and Cellular Biology, 2000. 20(21): p. 8001-8007.
104. Johnson, R.E., et al., hRAD30 mutations in the variant form of xeroderma pigmentosum. Science, 1999. 285(5425): p. 263-265.
105. Masutani, C., et al., The XPV (xeroderma pigmentosum variant) gene encodes human DNA polymerase eta. Nature, 1999. 399(6737): p. 700-704.
106. Johnson, R.E., et al., Fidelity of human DNA polymerase eta. Journal of Biological Chemistry, 2000. 275(11): p. 7447-7450.
107. Washington, M.T., L. Prakash, and S. Prakash, Mechanism of nucleotide incorporation opposite a thymine-thymine dimer by yeast DNA polymerase eta. Proceedings of the National Academy of Sciences of the United States of America, 2003. 100(21): p. 12093-12098.
108. Carlson, K.D. and A.T. Washington, Mechanism of efficient and accurate nucleotide incorporation opposite 7,8-dihydro-8-oxoguanine by Saccharomyces cerevisiae DNA polymerase eta. Molecular and Cellular Biology, 2005. 25(6): p. 2169-2176.
109. Haracska, L., et al., Inefficient bypass of an abasic site by DNA polymerase eta. Journal of Biological Chemistry, 2001. 276(9): p. 6861-6866.
110. Silverstein, T.D., et al., Structural basis for the suppression of skin cancers by DNA polymerase eta. Nature, 2010. 465(7301): p. 1039-U96.
111. Ummat, A., et al., Human DNA Polymerase eta Is Pre-Aligned for dNTP Binding and Catalysis. Journal of Molecular Biology, 2012. 415(4): p. 627-634.
112. Biertuempfel, C., et al., Structure and mechanism of human DNA polymerase eta. Nature, 2010. 465(7301): p. 1044-U102.
113. Haracska, L., et al., Interaction with PCNA is essential for yeast DNA polymerase eta function. Molecular Cell, 2001. 8(2): p. 407-415.
221
114. Bomar, M.G., et al., Structure of the ubiquitin-binding zinc finger domain of human DNA Y-polymerase eta. Embo Reports, 2007. 8(3): p. 247-251.
115. Hishiki, A., et al., Structural Basis for Novel Interactions between Human Translesion Synthesis Polymerases and Proliferating Cell Nuclear Antigen. Journal of Biological Chemistry, 2009. 284(16): p. 10552-10560.
116. Li, G.M. and P. Modrich, RESTORATION OF MISMATCH REPAIR TO NUCLEAR EXTRACTS OF H6 COLORECTAL TUMOR-CELLS BY A HETERODIMER OF HUMAN MUTL HOMOLOGS. Proceedings of the National Academy of Sciences of the United States of America, 1995. 92(6): p. 1950-1954.
117. Modrich, P., Strand-specific mismatch repair in mammalian cells. Journal of Biological Chemistry, 1997. 272(40): p. 24727-24730.
118. Papadopoulos, N., et al., MUTATIONS OF GTBP IN GENETICALLY UNSTABLE CELLS. Science, 1995. 268(5219): p. 1915-1917.
119. Fishel, R. and T. Wilson, MutS homologs in mammalian cells. Current Opinion in Genetics & Development, 1997. 7(1): p. 105-113.
120. Alani, E., The Saccharomyces cerevisiae Msh2 and Msh6 proteins form a complex that specifically binds to duplex oligonucleotides containing mismatched DNA base pairs. Molecular and Cellular Biology, 1996. 16(10): p. 5604-5615.
121. Modrich, P., Mechanisms in eukaryotic mismatch repair. Journal of Biological Chemistry, 2006. 281(41): p. 30305-30309.
122. Warren, J.J., et al., Structure of the human MutS alpha DNA lesion recognition complex. Molecular Cell, 2007. 26(4): p. 579-592.
123. Obmolova, G., et al., Crystal structures of mismatch repair protein MutS and its complex with a substrate DNA. Nature, 2000. 407(6805): p. 703-710.
124. Lamers, M.H., et al., The crystal structure of DNA mismatch repair protein MutS binding to a G center dot T mismatch. Nature, 2000. 407(6805): p. 711-717.
125. Kunkel, T.A. and D.A. Erie, DNA mismatch repair, in Annual Review of Biochemistry. 2005. p. 681-710.
126. Lee, S.D., J.A. Surtees, and E. Alani, Saccharomyces cerevisiae MSH2-MSH3 and MSH2-MSH6 complexes display distinct requirements for DNA binding domain I in mismatch recognition. Journal of Molecular Biology, 2007. 366(1): p. 53-66.
127. Drotschmann, K., et al., Asymmetric recognition of DNA local distortion - Structure-based functional studies of eukaryotic Msh2-Msh6. Journal of Biological Chemistry, 2001. 276(49): p. 46225-46229.
128. Cristovao, M., et al., Single-molecule multiparameter fluorescence spectroscopy reveals directional MutS binding to mismatched bases in DNA. Nucleic Acids Research, 2012. 40(12): p. 5448-5464.
222
129. Masih, P.J., D. Kunnev, and T. Melendy, Mismatch repair proteins are recruited to replicating DNA through interaction with proliferating cell nuclear antigen (PCNA). Nucleic Acids Research, 2008. 36(1): p. 67-75.
130. Clark, A.B., et al., Multiple functions for the N-terminal region of Msh6. Nucleic Acids Research, 2007. 35(12): p. 4114-4123.
131. Matton, N., J. Simonetti, and K. Williams, Identification of mismatch repair protein complexes in HeLa nuclear extracts and their interaction with heteroduplex DNA. Journal of Biological Chemistry, 2000. 275(23): p. 17808-17813.
132. Edelbrock, M., et al., Recognition and binding of mismatch repair proteins at an oncogenic hot spot. Bmc Molecular Biology, 2005. 6.
133. Bellacosa, A., Functional interactions and signaling properties of mammalian DNA mismatch repair proteins. Cell Death and Differentiation, 2001. 8(11): p. 1076-1092.
134. Duckett, D.R., et al., Human MutS alpha recognizes damaged DNA base pairs containing O-6-methylguanine, O-4-methylthymine, or the cisplatin-d(GpG) adduct. Proceedings of the National Academy of Sciences of the United States of America, 1996. 93(13): p. 6443-6447.
135. Mu, D., et al., Recognition and repair of compound DNA lesions (base damage and mismatch) by human mismatch repair and excision repair systems. Molecular and Cellular Biology, 1997. 17(2): p. 760-769.
136. Colussi, C., et al., The mammalian mismatch repair pathway removes DNA 8-oxodGMP incorporated from the oxidized dNTP pool. Current Biology, 2002. 12(11): p. 912-918.
137. Russo, M.T., et al., Role of MUTYH and MSH2 in the Control of Oxidative DNA Damage, Genetic Instability, and Tumorigenesis. Cancer Research, 2009. 69(10): p. 4372-4379.
138. Mazurek, A., M. Berardini, and R. Fishel, Activation of human MutS homologs by 8-oxo-guanine DNA damage. Journal of Biological Chemistry, 2002. 277(10): p. 8260-8266.
139. Larson, E.D., K. Iams, and J.T. Drummond, Strand-specific processing of 8-oxoguanine by the human mismatch repair pathway: inefficient removal of 8-oxoguanine paired with adenine or cytosine. DNA Repair, 2003. 2(11): p. 1199-1210.
140. Arcangeli, L., et al., Site- and strand-specific mismatch repair of human H-ras genomic DNA in a mammalian cell line. Carcinogenesis, 1997. 18(7): p. 1311-1318.
141. Jiricny, J., The multifaceted mismatch-repair system. Nature Reviews Molecular Cell Biology, 2006. 7(5): p. 335-346.
223
142. Edelbrock, M., Kaliyaperumal, S., Williams, K., Structural, molecular, and cellular functions of MSH2 and MSH6 during DNA mismatch repair, damage signaling and other noncanonical activities. Mutation Research: Fundamental and molecular mechanisms of mutagenesis, 2013.
143. Chahwan, R., et al., AIDing antibody diversity by error-prone mismatch repair. Seminars in Immunology, 2012. 24(4): p. 293-300.
144. Rada, C., et al., Immunoglobulin isotype switching is inhibited and somatic hypermutation perturbed in UNG-deficient mice. Current Biology, 2002. 12(20): p. 1748-1755.
145. Wiesendanger, M., et al., Somatic hypermutation in MutS homologue (MSH)3-, MSH6-, and MSH3/MSH6-deficient mice reveals a role for the MSH2-MSH6 heterodimer in modulating the base substitution pattern. Journal of Experimental Medicine, 2000. 191(3): p. 579-584.
146. Martin, A., et al., Msh2 ATPase activity is essential for somatic hypermutation at A-T basepairs and for efficient class switch recombination. Journal of Experimental Medicine, 2003. 198(8): p. 1171-1178.
147. Zeng, X.M., et al., DNA polymerase eta is an A-T mutator in somatic hypermutation of immunoglobulin variable genes. Nature Immunology, 2001. 2(6): p. 537-541.
148. Faili, A., et al., DNA polymerase eta is involved in hypermutation occurring during immunoglobulin class switch recombination. Journal of Experimental Medicine, 2004. 199(2): p. 265-270.
149. Zeng, X.M., et al., Absence of DNA polymerase eta reveals targeting of C mutations on the nontranscribed strand in immunoglobulin switch regions. Journal of Experimental Medicine, 2004. 199(7): p. 917-924.
150. Wilson, T.M., et al., MSH2-MSH6 stimulates DNA polymerase eta, suggesting a role for A : T mutations in antibody genes. Journal of Experimental Medicine, 2005. 201(4): p. 637-645.
151. Miyachi, K., M.J. Fritzler, and E.M. Tan, AUTOANTIBODY TO A NUCLEAR ANTIGEN IN PROLIFERATING CELLS. Journal of Immunology, 1978. 121(6): p. 2228-2234.
152. Bravo, R. and J.E. Celis, SEARCH FOR DIFFERENTIAL POLYPEPTIDE-SYNTHESIS THROUGHOUT THE CELL-CYCLE OF HELA-CELLS. Journal of Cell Biology, 1980. 84(3): p. 795-802.
153. Bravo, R., et al., Identification of a nuclear polypeptide ("cyclin") whose relative proportion is sensitive to changes in the rate of cell proliferation and to transformation. Progress in clinical and biological research, 1982. 85 Pt A: p. 235-48.
154. Celis, J.E., et al., CYCLIN - A NUCLEAR-PROTEIN WHOSE LEVEL CORRELATES DIRECTLY WITH THE PROLIFERATIVE STATE OF NORMAL AS WELL AS TRANSFORMED-CELLS. Leukemia Research, 1984. 8(2): p. 143-157.
224
155. Moldovan, G.L., B. Pfander, and S. Jentsch, PCNA, the maestro of the replication fork. Cell, 2007. 129: p. 665-679.
156. Krishna, T.S.R., et al., Crystal structure of the eukaryotic DNA polymerase processivity factor PCNA. Cell, 1994. 79(7): p. 1233-1243.
157. Georgescu, R.E., et al., Structure of a sliding clamp on DNA. Cell, 2008. 132(1): p. 43-54.
158. McNally, R., et al., Analysis of the role of PCNA-DNA contacts during clamp loading. Bmc Structural Biology, 2010. 10.
159. Kochaniak, A.B., et al., Proliferating Cell Nuclear Antigen Uses Two Distinct Modes to Move along DNA. Journal of Biological Chemistry, 2009. 284(26): p. 17700-17710.
160. Freudenthal, B.D., et al., Structure of monoubiquitinated PCNA and implications for translesion synthesis and DNA polymerase exchange. Nature Structural & Molecular Biology, 2010. 17(4): p. 479-U123.
161. Freudenthal, B.D., et al., Crystal structure of SUMO-modified proliferating cell nuclear antigen. J Mol Biol, 2011. 406(1): p. 9-17.
162. Gomes, X.V. and P.M.J. Burgers, ATP utilization by yeast replication factor C I. ATP-mediated interaction with DNA and with proliferating cell nuclear antigen. Journal of Biological Chemistry, 2001. 276(37): p. 34768-34775.
163. Langston, L.D. and M. O'Donnell, DNA Polymerase delta Is Highly Processive with Proliferating Cell Nuclear Antigen and Undergoes Collision Release upon Completing DNA. Journal of Biological Chemistry, 2008. 283(43): p. 29522-29531.
164. de Saro, F.J.L., Regulation of Interactions with Sliding Clamps During DNA Replication and Repair. Current Genomics, 2009. 10(3): p. 206-215.
165. Chilkova, O., et al., The eukaryotic leading and lagging strand DNA polymerases are loaded onto primer-ends via separate mechanisms but have comparable processivity in the presence of PCNA. Nucleic Acids Research, 2007. 35(19): p. 6588-6597.
166. Einolf, H.J. and F.P. Guengerich, Kinetic analysis of nucleotide incorporation by mammalian DNA polymerase delta. Journal of Biological Chemistry, 2000. 275(21): p. 16316-16322.
167. Chapados, B.R., et al., Structural basis for FEN-1 substrate specificity and PCNA-mediated activation in DNA replication and repair. Cell, 2004. 116(1): p. 39-50.
168. Pascal, J.M., et al., Human DNA ligase I completely encircles and partially unwinds nicked DNA. Nature, 2004. 432(7016): p. 473-478.
169. Lee, S.D. and E. Alani, Analysis of interactions between mismatch repair initiation factors and the replication processivity factor PCNA. Journal of Molecular Biology, 2006. 355(2): p. 175-184.
225
170. Montecucco, A., et al., DNA ligase I is recruited to sites of DNA replication by an interaction with proliferating cell nuclear antigen: identification of a common targeting mechanism for the assembly of replication factories. Embo Journal, 1998. 17(13): p. 3786-3795.
171. Iyer, R.R., et al., The MutS alpha-proliferating cell nuclear antigen interaction in human DNA mismatch repair. Journal of Biological Chemistry, 2008. 283(19): p. 13310-13319.
172. Rousseau, D., et al., Growth inhibition by CDK-cyclin and PCNA binding domains of p21 occurs by distinct mechanisms and is regulated by ubiquitin-proteasome pathway. Oncogene, 1999. 18(30): p. 4313-4325.
173. Waga, S., et al., THE P21 INHIBITOR OF CYCLIN-DEPENDENT KINASES CONTROLS DNA-REPLICATION BY INTERACTION WITH PCNA. Nature, 1994. 369(6481): p. 574-578.
174. Ducoux, M., et al., Mediation of proliferating cell nuclear antigen (PCNA)-dependent DNA replication through a conserved p21(Cip1)-like PCNA-binding motif present in the third subunit of human DNA polymerase delta. Journal of Biological Chemistry, 2001. 276(52): p. 49258-49266.
175. Warbrick, E., et al., Homologous regions of Fen1 and p21(Cip1) compete for binding to the same site on PCNA: A potential mechanism to co-ordinate DNA replication and repair. Oncogene, 1997. 14(19): p. 2313-2321.
176. Oku, T., et al., Functional sites of human PCNA which interact with p21 (Cip1/Waf1), DNA polymerase delta and replication factor C. Genes to Cells, 1998. 3(6): p. 357-369.
177. Bergink, S. and S. Jentsch, Principles of ubiquitin and SUMO modifications in DNA repair. Nature, 2009. 458(7237): p. 461-467.
178. Shaheen, M., I. Shanmugam, and R. Hromas, The Role of PCNA Posttranslational Modifications in Translesion Synthesis. J Nucleic Acids, 2010. 2010.
179. Ulrich, H.D. and H. Walden, Ubiquitin signalling in DNA replication and repair. Nature Reviews Molecular Cell Biology, 2010. 11(7): p. 479-489.
180. Watts, F.Z., Sumoylation of PCNA: Wrestling with recombination at stalled replication forks. DNA Repair, 2006. 5(3): p. 399-403.
181. Naryzhny, S.N. and H. Lee, The post-translational modifications of proliferating cell nuclear antigen - Acetylation, not phosphorylation, plays an important role in the regulation of its function. Journal of Biological Chemistry, 2004. 279(19): p. 20194-20199.
182. Prosperi, E., et al., PROLIFERATING CELL NUCLEAR ANTIGEN BOUND TO DNA-SYNTHESIS SITES - PHOSPHORYLATION AND ASSOCIATION WITH CYCLIN-D1 AND CYCLIN-A. Experimental Cell Research, 1994. 215(2): p. 257-262.
183. Hoege, C., et al., RAD6-dependent DNA repair is linked to modification of PCNA by ubiquitin and SUMO. Nature, 2002. 419(6903): p. 135-141.
226
184. Stelter, P. and H.D. Ulrich, Control of spontaneous and damage-induced mutagenesis by SUMO and ubiquitin conjugation. Nature, 2003. 425(6954): p. 188-191.
185. Bienko, M., et al., Ubiquitin-binding domains in Y-family polymerases regulate translesion synthesis. Science, 2005. 310(5755): p. 1821-1824.
186. Guo, C.X., et al., Ubiquitin-binding motifs in REV1 protein are required for its role in the tolerance of DNA damage. Molecular and Cellular Biology, 2006. 26(23): p. 8892-8900.
187. Huttner, D. and H.D. Ulrich, Cooperation of replication protein A with the ubiquitin ligase Rad18 in DNA damage bypass. Cell Cycle, 2008. 7(23): p. 3629-3633.
188. Davies, A.A., et al., Activation of ubiquitin-dependent DNA damage bypass is mediated by replication protein A. Molecular Cell, 2008. 29(5): p. 625-636.
189. Kannouche, P.L., J. Wing, and A.R. Lehmann, Interaction of human DNA polymerase eta with monoubiquitinated PCNA: A possible mechanism for the polymerase switch in response to DNA damage. Molecular Cell, 2004. 14(4): p. 491-500.
190. Haracska, L., et al., Ubiquitylation of yeast proliferating cell nuclear antigen and its implications for translesion DNA synthesis. Proceedings of the National Academy of Sciences of the United States of America, 2006. 103(17): p. 6477-6482.
191. Garg, P. and P.M. Burgers, Ubiquitinated proliferating cell nuclear antigen activates translesion DNA polymerases eta and REV1. Proceedings of the National Academy of Sciences of the United States of America, 2005. 102(51): p. 18361-18366.
192. Tsutakawa, S.E., et al., Solution X-ray scattering combined with computational modeling reveals multiple conformations of covalently bound ubiquitin on PCNA. Proc Natl Acad Sci U S A, 2011. 108(43): p. 17672-7.
193. Ulrich, H.D. and S. Jentsch, Two RING finger proteins mediate cooperation between ubiquitin-conjugating enzymes in DNA repair. Embo Journal, 2000. 19(13): p. 3388-3397.
194. Plosky, B.S., et al., Controlling the subcellular localization of DNA polymerases iota and eta via interactions with ubiquitin. Embo Journal, 2006. 25(12): p. 2847-2855.
195. Papouli, E., et al., Crosstalk between SUMO and ubiquitin on PCNA is mediated by recruitment of the helicase Srs2p. Molecular Cell, 2005. 19(1): p. 123-133.
196. Parker, J.L., et al., SUMO modification of PCNA is controlled by DNA. Embo Journal, 2008. 27(18): p. 2422-2431.
197. Haracska, L., et al., Opposing effects of ubiquitin conjugation and SUMO modification of PCNA on replicational bypass of DNA lesions in Saccharomyces cerevisiae. Molecular and Cellular Biology, 2004. 24(10): p. 4267-4274.
227
198. Hingorani, M.M. and M. O'Donnell, Sliding clamps: A (tail)ored fit. Current Biology, 2000. 10(1): p. R25-R29.
199. Maga, G. and U. Hubscher, Proliferating cell nuclear antigen (PCNA): a dancer with many partners. Journal of Cell Science, 2003. 116(15): p. 3051-3060.
200. Tsurimoto, T., PCNA binding proteins. Frontiers in Bioscience, 1999. 4(CITED APRIL 7, 2000): p. d849-858.
201. Bruning, J.B. and Y. Shamoo, Structural and thermodynamic analysis of human PCNA with peptides derived from DNA polymerase-delta p66 subunit and flap endonuclease-1. Structure, 2004. 12(12): p. 2209-2219.
202. Prakash, S. and L. Prakash, Translesion DNA synthesis in eukaryotes: A one- or two-polymerase affair. Genes & Development, 2002. 16(15): p. 1872-1883.
203. Washington, M.T., et al., Variations on a theme: Eukaryotic Y-family DNA polymerases. Biochim Biophys Acta, 2009.
204. Vijayakumar, S., et al., The C-terminal domain of yeast PCNA is required for physical and functional interactions with Cdc9 DNA ligase. Nucleic Acids Research, 2007. 35(5): p. 1624-1637.
205. Floresrozas, H., et al., CDK-INTERACTING PROTEIN-1 DIRECTLY BINDS WITH PROLIFERATING CELL NUCLEAR ANTIGEN AND INHIBITS DNA-REPLICATION CATALYZED BY THE DNA-POLYMERASE-DELTA HOLOENZYME. Proceedings of the National Academy of Sciences of the United States of America, 1994. 91(18): p. 8655-8659.
206. Gibbs, E., et al., The influence of the proliferating cell nuclear antigen-interacting domain of p21(CIP1) on DNA synthesis catalyzed by the human and Saccharomyces cerevisiae polymerase delta holoenzymes. Journal of Biological Chemistry, 1997. 272(4): p. 2373-2381.
207. Gulbis, J.M., et al., Structure of the C-terminal region of p21(WAF1/CIP1) complexed with human PCNA. Cell, 1996. 87(2): p. 297-306.
208. Sakurai, S., et al., Structural basis for recruitment of human flap endonuclease 1 to PCNA. Embo Journal, 2005. 24(4): p. 683-693.
209. Bowman, G.D., M. O'Donnell, and J. Kuriyan, Structural analysis of a eukaryotic sliding DNA clamp-clamp loader complex. Nature, 2004. 429(6993): p. 724-730.
210. Pascal, J.M., et al., A flexible interface between DNA ligase and PCNA supports conformational switching and efficient ligation of DNA. Molecular Cell, 2006. 24(2): p. 279-291.
211. Williams, G.J., et al., Structure of the heterotrimeric PCNA from Sulfolobus solfataricus. Acta Crystallographica Section F-Structural Biology and Crystallization Communications, 2006. 62: p. 944-948.
228
212. Mayanagi, K., et al., Architecture of the DNA polymerase B-proliferating cell nuclear antigen (PCNA)-DNA ternary complex. Proceedings of the National Academy of Sciences of the United States of America, 2011. 108(5): p. 1845-1849.
213. Henneke, G., S. Koundrioukoff, and U. Hubscher, Phosphorylation of human Fen1 by cyclin-dependent kinase modulates its role in replication fork regulation. Oncogene, 2003. 22(28): p. 4301-4313.
214. Scott, M.T., N. Morrice, and K.L. Ball, Reversible phosphorylation at the C-terminal regulatory domain of p21(Waf1/Cip1) modulates proliferating cell nuclear antigen binding. Journal of Biological Chemistry, 2000. 275(15): p. 11529-11537.
215. Freudenthal, B.D., et al., Structure of monoubiquitinated PCNA and implications for translesion synthesis and DNA polymerase exchange. Nat Struct Mol Biol, 2010. 17(4): p. 479-84.
216. Haracska, L., et al., Physical and functional interactions of human DNA polymerase eta with PCNA. Molecular and Cellular Biology, 2001. 21(21): p. 7199-7206.
217. Haracska, L., et al., Targeting of human DNA polymerase iota to the replication machinery via interaction with PCNA. Proceedings of the National Academy of Sciences of the United States of America, 2001. 98(25): p. 14256-14261.
218. Haracska, L., et al., Stimulation of DNA synthesis activity of human DNA polymerase kappa by PCNA. Molecular and Cellular Biology, 2002. 22(3): p. 784-791.
219. Freudenthal, B.D., et al., Structure of a Mutant Form of Proliferating Cell Nuclear Antigen That Blocks Translesion DNA Synthesis. Biochemistry, 2008. 47(50): p. 13354-13361.
220. Acharya, N., et al., Roles of PCNA-binding and ubiquitin-binding domains in human DNA polymerase eta in translesion DNA synthesis. Proceedings of the National Academy of Sciences of the United States of America, 2008. 105(46): p. 17724-17729.
221. Vidal, A.E., et al., Proliferating cell nuclear antigen-dependent coordination of the biological functions of human DNA polymerase iota. Journal of Biological Chemistry, 2004. 279(46): p. 48360-48368.
222. Guo, C.X., et al., REV1 protein interacts with PCNA: Significance of the REV1 BRCT domain in vitro and in vivo. Molecular Cell, 2006. 23(2): p. 265-271.
223. Wood, A., P. Garg, and P.M.J. Burgers, A ubiquitin-binding motif in the translesion DNA polymerase Rev1 mediates its essential functional interaction with ubiquitinated proliferating cell nuclear antigen in response to DNA damage. Journal of Biological Chemistry, 2007. 282(28): p. 20256-20263.
224. Tissier, A., et al., Co-localization in replication foci and interaction of human Y-family members, DNA polymerase pol eta and REV1 protein. DNA Repair, 2004. 3(11): p. 1503-1514.
229
225. Murakumo, Y., et al., Analyses of ultraviolet-induced focus formation of hREV1 protein. Genes to Cells, 2006. 11(3): p. 193-205.
226. de Groote, F.H., et al., The Rev1 translesion synthesis polymerase has multiple distinct DNA binding modes. DNA Repair, 2011. 10(9): p. 915-925.
227. Bomar, M.G., et al., Unconventional Ubiquitin Recognition by the Ubiquitin-Binding Motif within the Y Family DNA Polymerases iota and Rev1. Molecular Cell, 2010. 37(3): p. 408-417.
228. Burschowsky, D., et al., Structural Analysis of the Conserved Ubiquitin-binding Motifs (UBMs) of the Translesion Polymerase iota in Complex with Ubiquitin. Journal of Biological Chemistry, 2011. 286(2): p. 1364-1373.
229. Tsutakawa, S.E., et al., Solution X-ray scattering combined with computational modeling reveals multiple conformations of covalently bound ubiquitin on PCNA. Proceedings of the National Academy of Sciences of the United States of America, 2011. 108(43): p. 17672-17677.
230. Indiani, C., et al., Translesion DNA polymerases remodel the replisome and alter the speed of the replicative helicase. Proceedings of the National Academy of Sciences of the United States of America, 2009. 106(15): p. 6031-6038.
231. Kuban, W., et al., Mutator phenotype resulting from DNA polymerase IV overproduction in Escherichia coli: Preferential mutagenesis on the lagging strand. Journal of Bacteriology, 2005. 187(19): p. 6862-6866.
232. Zhuang, Z.H., et al., Regulation of polymerase exchange between Pol eta and Pol delta by monoubiquitination of PCNA and the movement of DNA polymerase holoenzyme. Proceedings of the National Academy of Sciences of the United States of America, 2008. 105(14): p. 5361-5366.
233. Masuda, Y., J. Piao, and K. Kamiya, DNA Replication-Coupled PCNA Mono-Ubiquitination and Polymerase Switching in a Human In Vitro System. Journal of Molecular Biology, 2010. 396(3): p. 487-500.
234. Leach, C.A. and W.M. Michael, Ubiquitin/SUMO modification of PCNA promotes replication fork progression in Xenopus laevis egg extracts. Journal of Cell Biology, 2005. 171(6): p. 947-954.
235. Dionne, I., et al., A heterotrimeric PCNA in the hyperthermophilic archaeon Sulfolobus solfataricus. Molecular Cell, 2003. 11(1): p. 275-282.
236. van der Kemp, P.A., et al., PCNA monoubiquitylation and DNA polymerase ubiquitin-binding domain are required to prevent 8-oxoguanine-induced mutagenesis in Saccharomyces cerevisiae. Nucleic Acids Research, 2009. 37(8): p. 2549-2559.
237. Bunting, K.A., S.M. Roe, and L.H. Pearl, Structural basis for recruitment of translesion DNA polymerase Pol IV/DinB to the beta-clamp. Embo Journal, 2003. 22(21): p. 5883-5892.
230
238. Zhang, H.S., P.E.M. Gibbs, and C.W. Lawrence, The Saccharomyces cerevisiae rev6-1 mutation, which inhibits both the lesion bypass and the recombination mode of DNA damage tolerance, is an allele of POL30, encoding proliferating cell nuclear antigen. Genetics, 2006. 173(4): p. 1983-1989.
239. Amin, N.S. and C. Holm, In vivo analysis reveals that the interdomain region of the yeast proliferating cell nuclear antigen is important for DNA replication and DNA repair. Genetics, 1996. 144(2): p. 479-493.
240. Northam, M.R., et al., A novel function of DNA polymerase zeta regulated by PCNA. Embo Journal, 2006. 25(18): p. 4316-4325.
241. Freudenthal, B.D., et al., A charged residue at the subunit interface of PCNA promotes trimer formation by destabilizing alternate subunit interactions. Acta Crystallogr D Biol Crystallogr, 2009. 65(Pt 6): p. 560-6.
242. Lau, P.J., H. Flores-Rozas, and R.D. Kolodner, Isolation and characterization of new proliferating cell nuclear antigen (POL30) mutator mutants that are defective in DNA mismatch repair. Molecular and Cellular Biology, 2002. 22(19): p. 6669-6680.
243. Stone, J.E., et al., Role of proliferating cell nuclear antigen interactions in the mismatch repair-dependent processing of mitotic and meiotic recombination intermediates in yeast. Genetics, 2008. 178(3): p. 1221-1236.
244. Burgers, P.M.J., Polymerase Dynamics at the Eukaryotic DNA Replication Fork. Journal of Biological Chemistry, 2009. 284(7): p. 4041-4045.
245. Kunkel, T.A. and P.M. Burgers, Dividing the workload at a eukaryotic replication fork. Trends in Cell Biology, 2008. 18(11): p. 521-527.
246. McElhinny, S.A.N., et al., Division of labor at the eukaryotic replication fork. Molecular Cell, 2008. 30(2): p. 137-144.
247. Shcherbakova, P.V., K. Bebenek, and T.A. Kunkel, Functions of eukaryotic DNA polymerases. Sci Aging Knowledge Environ, 2003. 2003(8): p. RE3.
248. Johansson, E., J. Majka, and P.M.J. Burgers, Structure of DNA polymerase delta from Saccharomyces cerevisiae. Journal of Biological Chemistry, 2001. 276(47): p. 43824-43828.
249. Jain, R., et al., Structural Insights into Yeast DNA Polymerase delta by Small Angle X-ray Scattering. Journal of Molecular Biology, 2009. 394(3): p. 377-382.
250. Hashimoto, K., et al., Fidelity of DNA polymerase delta holoenzyme from Saccharomyces cerevisiae: The sliding clamp proliferating cell nuclear antigen decreases its fidelity. Biochemistry, 2003. 42(48): p. 14207-14213.
251. Lehmann, A.R., Replication of damaged DNA by translesion synthesis in human cells. Febs Letters, 2005. 579(4): p. 873-876.
252. Lehmann, A.R., Replication of damaged DNA. Cell Cycle, 2003. 2(4): p. 300-2.
231
253. Waters, L.S., et al., Eukaryotic Translesion Polymerases and Their Roles and Regulation in DNA Damage Tolerance. Microbiology and Molecular Biology Reviews, 2009. 73(1): p. 134-+.
254. Sale, J.E., A.R. Lehmann, and R. Woodgate, Y-family DNA polymerases and their role in tolerance of cellular DNA damage. Nature Reviews Molecular Cell Biology, 2012. 13(3): p. 141-152.
255. Trincao, J., et al., Structure of the catalytic core of S-cerevisiae DNA polymerase eta: Implications for translesion DNA synthesis. Molecular Cell, 2001. 8(2): p. 417-426.
256. Washington, M.T., L. Prakash, and S. Prakash, Yeast DNA polymerase eta utilizes an induced-fit mechanism of nucleotide incorporation. Cell, 2001. 107(7): p. 917-927.
257. Naryzhny, S.N., Proliferating cell nuclear antigen: a proteomics view. Cellular and Molecular Life Sciences, 2008. 65(23): p. 3789-3808.
258. Zhuang, Z.H. and Y.X. Ai, Processivity factor of DNA polymerase and its expanding role in normal and translesion DNA synthesis. Biochimica Et Biophysica Acta-Proteins and Proteomics, 2010. 1804(5): p. 1081-1093.
259. Dieckman, L.M., B.D. Freudenthal, and M.T. Washington, PCNA Structure and Function: Insights from Structures of PCNA Complexes and Post-translationally Modified PCNA. Subcell Biochem, 2012. 62: p. 281-99.
260. Freudenthal, B.D., et al., A charged residue at the subunit interface of PCNA promotes trimer formation by destabilizing alternate subunit interactions. Acta Crystallographica Section D-Biological Crystallography, 2009. 65: p. 560-566.
261. Finkelstein, J., et al., Overproduction and analysis of eukaryotic multiprotein complexes in Escherichia coli using a dual-vector strategy. Analytical Biochemistry, 2003. 319(1): p. 78-87.
262. Garg, P., et al., Proliferating cell nuclear antigen promotes translesion synthesis by DNA polymerase zeta. Journal of Biological Chemistry, 2005. 280(25): p. 23446-23450.
263. Hanahan, D. and R.A. Weinberg, Hallmarks of Cancer: The Next Generation. Cell, 2011. 144(5): p. 646-674.
264. O'Roak, B.J., et al., Exome sequencing in sporadic autism spectrum disorders identifies severe de novo mutations. Nature Genetics, 2011. 43(6): p. 585-U125.
265. Xu, B., et al., Strong association of de novo copy number mutations with sporadic schizophrenia. Nature Genetics, 2008. 40(7): p. 880-885.
266. Veltman, J.A. and H.G. Brunner, De novo mutations in human genetic disease. Nature reviews. Genetics, 2012. 13(8): p. 565-75.
232
267. Sharma, N.M., O.V. Kochenova, and P.V. Shcherbakova, The Non-canonical Protein Binding Site at the Monomer-Monomer Interface of Yeast Proliferating Cell Nuclear Antigen (PCNA) Regulates the Rev1-PCNA Interaction and Pol zeta/Rev1-dependent Translesion DNA Synthesis. Journal of Biological Chemistry, 2011. 286(38): p. 33557-33566.
268. Li, G.-M., Mechanisms and functions of DNA mismatch repair. Cell Research, 2008. 18(1): p. 85-98.
269. Kunz, C., Y. Saito, and P. Schaer, DNA Repair in Mammalian Cells. Cellular and Molecular Life Sciences, 2009. 66(6): p. 1021-1038.
270. Grilley, M., et al., ISOLATION AND CHARACTERIZATION OF THE ESCHERICHIA-COLI MUTL GENE-PRODUCT. Journal of Biological Chemistry, 1989. 264(2): p. 1000-1004.
271. Galio, L., C. Bouquet, and P. Brooks, ATP hydrolysis-dependent formation of a dynamic ternary nucleoprotein complex with MutS and MutL. Nucleic Acids Research, 1999. 27(11): p. 2325-2331.
272. Spampinato, C. and P. Modrich, The MutL ATPase is required for mismatch repair. Journal of Biological Chemistry, 2000. 275(13): p. 9863-9869.
273. Schofield, M.J., et al., Interaction of Escherichia coli MutS and MutL at a DNA mismatch. Journal of Biological Chemistry, 2001. 276(30): p. 28291-28299.
274. Acharya, S., et al., The coordinated functions of the E-coli MutS and MutL proteins in mismatch repair. Molecular Cell, 2003. 12(1): p. 233-246.
275. Au, K.G., K. Welsh, and P. Modrich, INITIATION OF METHYL-DIRECTED MISMATCH REPAIR. Journal of Biological Chemistry, 1992. 267(17): p. 12142-12148.
276. Palombo, F., et al., GTBP, A 160-KILODALTON PROTEIN ESSENTIAL FOR MISMATCH-BINDING ACTIVITY IN HUMAN-CELLS. Science, 1995. 268(5219): p. 1912-1914.
277. Iaccarino, I., et al., MSH6, a Saccharomyces cerevisiae protein that binds to mismatches as a heterodimer with MSH2. Current Biology, 1996. 6(4): p. 484-486.
278. Mazurek, A., et al., Sequence context effect for hMSH2-hMSH6 mismatch-dependent activation. Proceedings of the National Academy of Sciences of the United States of America, 2009. 106(11): p. 4177-4182.
279. Habraken, Y., et al., Binding of insertion/deletion DNA mismatches by the heterodimer of yeast mismatch repair proteins MSH2 and MSH3. Current Biology, 1996. 6(9): p. 1185-1187.
280. Genschel, J., et al., Isolation of MutS beta from human cells and comparison of the mismatch repair specificities of MutS beta and MutS alpha. Journal of Biological Chemistry, 1998. 273(31): p. 19895-19901.
233
281. Habraken, Y., et al., ATP-dependent assembly of a ternary complex consisting of a DNA mismatch and the yeast MSH2-MSH6 and MLH1-PMS1 protein complexes. Journal of Biological Chemistry, 1998. 273(16): p. 9837-9841.
282. Bowers, J., et al., MSH-MLH complexes formed at a DNA mismatch are disrupted by the PCNA sliding clamp. Journal of Molecular Biology, 2001. 306(5): p. 957-968.
283. Raschle, M., et al., Mutations within the hMLH1 and hPMS2 subunits of the human MutL alpha mismatch repair factor affect its ATPase activity, but not its ability to interact with hMutS alpha. Journal of Biological Chemistry, 2002. 277(24): p. 21810-21820.
284. Kolodner, R.D., MISMATCH REPAIR - MECHANISMS AND RELATIONSHIP TO CANCER SUSCEPTIBILITY. Trends in Biochemical Sciences, 1995. 20(10): p. 397-401.
285. de la Chapelle, A., Genetic predisposition to colorectal cancer. Nature Reviews Cancer, 2004. 4(10): p. 769-780.
286. Rowley, P.T., Inherited susceptibility to colorectal cancer, in Annual Review of Medicine. 2005. p. 539-554.
287. Bertrand, P., et al., Physical interaction between components of DNA mismatch repair and nucleotide excision repair. Proceedings of the National Academy of Sciences of the United States of America, 1998. 95(24): p. 14278-14283.
288. Johnson, R.E., et al., Evidence for involvement of yeast proliferating cell nuclear antigen in DNA mismatch repair. Journal of Biological Chemistry, 1996. 271(45): p. 27987-27990.
289. Dieckman, L.M., B.D. Freudenthal, and M.T. Washington, PCNA Structure and Function: Insights from Structures of PCNA Complexes and Post-translationally Modified PCNA. Sub-cellular biochemistry, 2012. 62: p. 281-99.
290. Chen, C., et al., Saccharomyces carevisiae pol30 (proliferating cell nuclear antigen) mutations impair replication fidelity and mismatch repair. Molecular and Cellular Biology, 1999. 19(11): p. 7801-7815.
291. Kokoska, R.J., et al., A mutation of the yeast gene encoding PCNA destabilizes both microsatellite and minisatellite DNA sequences. Genetics, 1999. 151(2): p. 511-519.
292. Antony, E. and M.M. Hingorani, Mismatch recognitioh-coupled stabilization of Msh2-Msh6 in an ATP-bound state at the initiation of DNA repair. Biochemistry, 2003. 42(25): p. 7682-7693.
293. Pflugrath, J.W., The finer things in X-ray diffraction data collection. Acta Crystallographica Section D-Biological Crystallography, 1999. 55: p. 1718-1725.
294. Read, R.J., Pushing the boundaries of molecular replacement with maximum likelihood. Acta Crystallographica Section D-Biological Crystallography, 2001. 57: p. 1373-1382.
234
295. Bailey, S., The Ccp4 Suite - Programs for Protein Crystallography. Acta Crystallographica Section D-Biological Crystallography, 1994. 50: p. 760-763.
296. Emsley, P. and K. Cowtan, Coot: model-building tools for molecular graphics. Acta Crystallographica Section D-Biological Crystallography, 2004. 60: p. 2126-2132.
297. TorresRamos, C.A., et al., Requirement of proliferating cell nuclear antigen in RAD6-dependent postreplicational DNA repair. Proceedings of the National Academy of Sciences of the United States of America, 1996. 93(18): p. 9676-9681.
298. Zhang, Z.G., K. Shibahara, and B. Stillman, PCNA connects DNA replication to epigenetic inheritance in yeast. Nature, 2000. 408(6809): p. 221-225.
299. Sharp, J.A., et al., Yeast histone deposition protein Asf1p requires Hir proteins and PCNA for heterochromatic silencing. Current Biology, 2001. 11(7): p. 463-473.
300. Goldsby, R.E., et al., High incidence of epithelial cancers in mice deficient for DNA polymerase delta proofreading. Proceedings of the National Academy of Sciences of the United States of America, 2002. 99(24): p. 15560-15565.
301. Goldsby, R.E., et al., Defective DNA polymerase-delta proofreading causes cancer susceptibility in mice. Nature Medicine, 2001. 7(6): p. 638-639.
302. Venkatesan, R.N., et al., Mutation at the polymerase active site of mouse DNA polymerase delta increases genomic instability and accelerates tumorigenesis. Molecular and Cellular Biology, 2007. 27: p. 7669-7682.
303. Flohr, T., et al., Detection of mutations in the DNA polymerase delta gene of human sporadic colorectal cancers and colon cancer cell lines. International Journal of Cancer, 1999. 80(6): p. 919-929.
304. Dacosta, L.T., et al., Polymerase-Delta Variants in Rer Colorectal Tumors. Nature Genetics, 1995. 9(1): p. 10-11.
305. Dieckman, L.M. and M.T. Washington, PCNA trimer instability inhibits translesion synthesis by DNA polymerase eta and by DNA polymerase delta. DNA Repair, 2013.
306. Masutani, C., et al., Xeroderma pigmentosum variant (XP-V) correcting protein from HeLa cells has a thymine dimer bypass DNA polymerase activity. Embo Journal, 1999. 18(12): p. 3491-3501.
307. Ishida, T. and K. Kinoshita, Prediction of disordered regions in proteins based on the meta approach. Bioinformatics, 2008. 24(11): p. 1344-1348.
308. Armstrong, A.A., F. Mohideen, and C.D. Lima, Recognition of SUMO-modified PCNA requires tandem receptor motifs in Srs2. Nature, 2012. 483(7387): p. 59-U99.
309. Fink, A.L., Natively unfolded proteins. Current Opinion in Structural Biology, 2005. 15(1): p. 35-41.
235
310. Dunker, A.K., et al., Function and structure of inherently disordered proteins. Current Opinion in Structural Biology, 2008. 18(6): p. 756-764.
311. Cortese, M.S., V.N. Uversky, and A.K. Dunker, Intrinsic disorder in scaffold proteins: Getting more from less. Progress in Biophysics & Molecular Biology, 2008. 98(1): p. 85-106.
312. Mayanagi, K., et al., Three-dimensional electron microscopy of the clamp loader small subunit from Pyrococcus furiosus. Journal of Structural Biology, 2001. 134(1): p. 35-45.
313. Kumar, R., et al., Stepwise loading of yeast clamp revealed by ensemble and single-molecule studies. Proceedings of the National Academy of Sciences of the United States of America, 2010. 107(46): p. 19736-19741.
314. Koundrioukoff, S., et al., A direct interaction between proliferating cell nuclear antigen (PCNA) and Cdk2 targets PCNA-interacting proteins for phosphorylation. Journal of Biological Chemistry, 2000. 275(30): p. 22882-22887.
315. Moldovan, G.L., B. Pfander, and S. Jentsch, PCNA controls establishment of sister chromatid cohesion during S phase. Molecular Cell, 2006. 23(5): p. 723-732.
316. Cortese, M.S., V.N. Uversky, and A. Keith Dunker, Intrinsic disorder in scaffold proteins: Getting more from less. Progress in Biophysics and Molecular Biology, 2008. 98(1): p. 85-106.
