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DNA replication by Escherichia coli DNA polymerase III is
regulated by the umuD gene products
A dissertation presented
by
Michelle C. Silva
to
The Department of Chemistry and Chemical Biology
In partial fulfillment of the requirements for the degree of
Doctor of Philosophy
In the field of
Chemistry
Northeastern University
Boston, Massachusetts
July 19, 2013
2
DNA replication by Escherichia coli DNA polymerase III is
regulated by the umuD gene products
by
Michelle C. Silva
ABSTRACT OF DISSERTATION
Submitted in partial fulfillment of the requirements for the degree of
Doctor of Philosophy in Chemistry and Chemical Biology in the
College of Science at Northeastern University
July 19, 2013
3
ABSTRACT
DNA polymerase III (DNA pol III) efficiently replicates the Escherichia coli genome, but it
cannot bypass DNA damage. Instead, translesion synthesis (TLS) DNA polymerases are
employed to replicate past damaged DNA; however, the exchange of replicative for TLS
polymerases is not understood. The umuD gene products, which are up-regulated during the SOS
response, were previously shown to bind to the α, β, ε subunits of DNA pol III. Full-length
UmuD inhibits DNA replication and prevents mutagenic TLS, while the cleaved form UmuD'
facilitates mutagenesis. In this work, we investigate the interactions between the α subunit of
DNA pol III, the single-stranded DNA binding protein SSB, and UmuD and we find that these
interactions regulate DNA replication in E. coli.
We show that the α subunit possesses two UmuD binding sites: at the N-terminus (residues 1-
280) and the C-terminal domain (residues 956-975) of α. The C-terminal site favors UmuD over
UmuD′. We also find that UmuD, but not UmuD′, disrupts the α-β complex. The C-terminal
binding site is also adjacent to the single-stranded DNA (ssDNA) binding site of α. We have
used single molecule DNA stretching experiments to demonstrate that UmuD specifically
inhibits binding of α to ssDNA. We predict using molecular modeling that UmuD residue D91 is
involved in the interaction between UmuD and α, and demonstrate that mutation of these
residues decreases the affinity of α for UmuD. We propose that the interaction between α and
UmuD contributes to the transition between replicative and TLS polymerases by removing α
from the β clamp and from ssDNA.
4
E. coli SSB binds to and protects ssDNA present during various DNA processing mechanisms
such as DNA repair, recombination, and replication. Here, we investigate the role of SSB during
DNA replication by DNA pol III. It has been previously shown that when SSB is bound to
ssDNA at the replication fork, replication by the polymerase core, which consists of the α, ε, and
θ subunits, is inhibited. In order for replication to occur in the presence of SSB the χ and ψ
subunits of DNA polymerase III are required. To date, the mechanism of this inhibition is poorly
understood. We demonstrate that SSB inhibits DNA polymerase III by directly binding to the C-
terminal domain of the α polymerase subunit. Interestingly, instead of binding the disordered tail
of SSB like other proteins including the χ subunit, the α subunit binds the globular N-terminal
domain of SSB that is responsible for binding ssDNA. Using single molecule force spectroscopy,
we also show that the α subunit stabilizes the SSB/ssDNA interaction. In the absence of the α
subunit, SSB stabilizes ssDNA below 20 pN and fully dissociates above 20 pN. However, in the
presence of the α subunit, SSB does not dissociate above 20 pN and the energy required to
dissociate SSB from ssDNA is increased by a factor of two. We show here that SSB inhibits
replication by α both through direct interactions between the two proteins as well as through SSB
binding to DNA; the overall effect of these interactions is that α stabilizes SSB binding to DNA,
which contributes to the inhibition.
We also show that SSB binds both full-length UmuD and the cleavage product UmuD′, with a
slightly higher affinity for UmuD′ versus full-length UmuD. Like with other SSB binding
partners, binding was localized to the C-terminal tail, as a decrease in binding to UmuD was
observed with the SSB-113 variant, which harbors the C-terminal tail mutation P177S. But
unlike other SSB protein binding partners, UmuD and especially UmuD′ also bind the globular
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N-terminal domain due to our observation that the isolated SSB-OB domain binds both UmuD
and UmuD′. In vivo, elevated levels of UmuD and UmuC but not UmuD′ complemented the
temperature sensitive phenotype of the ssb-113 allele, suggesting that the interactions between
UmuD and SSB and between UmuD′ and SSB have different physiological consequences for the
cell. Overall, we show that a network of protein-protein and protein-DNA interactions modulates
DNA replication under normal cellular conditions and in response to DNA replication stress.
6
ACKNOWLEDGEMENTS
I would like to thank my advisor Dr. Penny J. Beuning for allowing me to work in her lab all
these years. I am especially grateful to her for giving me the opportunity to manage the chemical
biology lab class. This opportunity inspired me to maybe consider a career in academia one day.
I would like to thank my thesis committee members: Dr. Mary Jo Ondrechen, Dr. David Budil,
and Dr. Carla Mattos. Thank you for accepting my invitation to be on my committee and for
advising me through the many thesis committee meetings we have had over the past couple of
years.
I would like to thank the Williams lab here in the physics department whom I collaborated with
on numerous projects, especially Dr. Kathy Chaurasiya, Dr. Kiran Pant, and Dr. Mark C.
Williams. Kathy, thank you for hearing me vent about my research all these years.
I would also like to thank all past and present members of the Beuning lab for supporting me
throughout this process. Especially, Dr. Jana Sefcikova, Dr. Ingrid DeVries Salgado, Dr. Jaylene
Ollivierre, Dr. Jason Walsh, Dr. Lisa Hawver, Ramya Parasuram, Philip Nevin, Nicholas
DeLateur, David Murison, Nicole Antczak, Lisa Ngu, and Caitlyn Mills. I would also like to
thank the undergrads and high school students I’ve worked closely with: Erin Ronayne, Lukas
Voortman, Arianna DiBenedetto, Monyrath Chan, Rachel Yao, Celeste Dang, and Christine
Alves.
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Finally, I would like to thank my family, especially my parents. Thank you for supporting me
and allowing me the opportunity to have the best education possible. Without you, I would not
have been who I am today.
Thank you all.
8
TABLE OF CONTENTS
ABSTRACT .................................................................................................................................... 2
ACKNOWLEDGEMENTS ............................................................................................................ 6
TABLE OF CONTENTS ................................................................................................................ 8
LIST OF FIGURES ...................................................................................................................... 13
LIST OF TABLES ........................................................................................................................ 16
LIST OF ABBREVIATIONS ....................................................................................................... 17
CHAPTER 1: Overview of DNA Replication in Escherichia coli ............................................... 19
1.1 E. coli DNA Polymerase III .............................................................................................................. 20
1.2 The Polymerase Core ........................................................................................................................ 24
1.2.1 The Polymerase Subunit, ........................................................................................................ 24
1.2.2 The Subunit ............................................................................................................................. 27
1.2.3 The Subunit ............................................................................................................................. 28
1.3 The Clamp Loader Complex and the Clamp Subunit .................................................................... 29
1.3.1 The Complex: ′:1:2:3:; Loading the Clamp ...................................................................... 29
1.3.2 The and Subunits ................................................................................................................. 34
1.3.3 The Clamp Subunit ................................................................................................................. 34
1.3.4 The Subunit ............................................................................................................................. 38
9
1.4 The Single-Stranded DNA Binding Protein, SSB............................................................................. 40
1.5 DNA Damage disrupts DNA Replication ......................................................................................... 42
1.6 Specialized DNA Polymerases facilitate DNA Damage Tolerance.................................................. 42
1.6.1 Regulation of Y Family DNA Polymerases ............................................................................... 45
1.6.2 Structural dynamics of UmuD and UmuD′ ................................................................................ 49
1.6.3 UmuD-Beta clamp interactions .................................................................................................. 51
1.7 DNA Polymerase Switching ............................................................................................................. 53
1.7.1 Replisome Dynamics ................................................................................................................. 54
1.7.2 Polymerase Switching: Tool-Belt and Active Exchange Models .............................................. 56
1.7.3 Gap-filling Model ...................................................................................................................... 59
1.8 REFERENCES ................................................................................................................................. 60
CHAPTER 2: Selective disruption of the DNA polymerase III α-β complex by the umuD gene
products ......................................................................................................................................... 76
2.1 INTRODUCTION ............................................................................................................................ 77
2.2 MATERIALS AND METHODS ...................................................................................................... 81
2.2.1 Proteins and Plasmids ................................................................................................................ 81
2.2.2 Tryptophan Fluorescence Assay ................................................................................................ 83
2.2.3 Thermal-Shift Assays ................................................................................................................. 84
2.2.4 UmuD in vitro Cleavage Assays ................................................................................................ 84
2.2.5 Cross-Linking of UmuD using Bis-maleimidohexane (BMH) .................................................. 85
2.2.6 Fluorescence Resonance Energy Transfer ................................................................................. 85
10
2.3 RESULTS ......................................................................................................................................... 87
2.3.1 α binds UmuD via two UmuD Binding Sites on α ..................................................................... 87
2.3.2 α Inhibits RecA/ssDNA-Facilitated Cleavage in vitro ............................................................... 91
2.3.3 The Interaction between α and UmuD is Conformation Dependent .......................................... 95
2.3.4 The C-terminal Binding Site Favors Full-length UmuD ............................................................ 97
2.3.5 UmuD Disrupts the DNA Pol III α-β Complex ......................................................................... 99
2.4 DISCUSSION ................................................................................................................................. 101
2.5 REFERENCES ............................................................................................................................... 104
Chapter 3: Polymerase manager protein UmuD directly regulates E. coli DNA polymerase III α
binding to ssDNA ....................................................................................................................... 110
3.1 INTRODUCTION .......................................................................................................................... 111
3.2 MATERIALS AND METHODS .................................................................................................... 114
3.2.1 Proteins and plasmids............................................................................................................... 114
3.2.2 Single molecule DNA stretching ............................................................................................. 115
3.2.3 Protein-protein docking............................................................................................................ 118
3.2.4 Thermal stability assay............................................................................................................. 119
3.2.5 RecA/ssDNA facilitated cleavage assay .................................................................................. 119
3.2.6 Tryptophan fluorescence assay ................................................................................................ 120
3.3 RESULTS ....................................................................................................................................... 120
3.3.1 UmuD inhibits α binding to ssDNA ......................................................................................... 120
3.3.2 Specific UmuD variants disrupt the UmuD-α interaction ........................................................ 122
11
3.3.3 Specific variants disrupt UmuD inhibition of α binding to ssDNA ......................................... 127
3.4 DISCUSSION ................................................................................................................................. 129
3.5 REFERENCES ............................................................................................................................... 132
Chapter 4: Replication by E. coli DNA pol III α is inhibited by direct binding to the OB domain
of single stranded DNA binding protein ..................................................................................... 136
4.1 INTRODUCTION .......................................................................................................................... 136
4.2 MATERIALS AND METHODS .................................................................................................... 139
4.2.1 Proteins and Plasmids .............................................................................................................. 139
4.2.2 Primer-Extension Assay ........................................................................................................... 141
4.2.3 Tryptophan Fluorescence Quenching Assay ............................................................................ 142
4.2.4 Single Molecule Force Spectroscopy using Optical Tweezers ................................................ 143
4.3 RESULTS ....................................................................................................................................... 146
4.3.1 DNA pol III α interacts with the OB fold of SSB .................................................................... 146
4.3.2 SSB interacts with the C-terminal domain of DNA pol III α ................................................... 150
4.3.3 SSB inhibits DNA pol III α when bound to ssDNA ................................................................ 152
4.3.4 DNA pol III α stabilizes SSB on ssDNA ................................................................................. 155
4.4 DISCUSSION ................................................................................................................................. 157
4.5 REFERENCES ............................................................................................................................... 159
Chapter 5: The E. coli single stranded DNA binding protein SSB binds the UmuD2 polymerase
manager protein .......................................................................................................................... 163
5.1 INTRODUCTION .......................................................................................................................... 163
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5.2 MATERIALS AND METHODS .................................................................................................... 166
5.2.1 Proteins, Strains and Plasmids ................................................................................................. 166
5.2.2 Tryptophan Fluorescence Assay .............................................................................................. 167
5.2.3 Quantitative Transformation Assay ......................................................................................... 168
5.2.4 Bis-maleimidohexane (BMH) Cross-linking ........................................................................... 168
5.3 RESULTS ....................................................................................................................................... 169
5.3.1 SSB binds the umuD gene products ......................................................................................... 169
5.3.2 The variant SSB-113 disrupts the interaction with UmuD ...................................................... 171
5.3.3 Full-length UmuD but not UmuD′ complements the phenotype of the ssb-113 allele ............ 173
5.3.4 SSB does not disrupt the dynamics of the N-terminal arms of UmuD .................................... 174
5.4 DISCUSSION ................................................................................................................................. 176
5.5 REFERENCES ............................................................................................................................... 178
Chapter 6: Future Considerations ............................................................................................... 182
6.1 Does UmuD affect the processivity of DNA pol III? ..................................................................... 184
6.2 Do these proteins bind as a complex? ............................................................................................. 185
6.3 Does UmuD also have a role in regulating DNA repair? ................................................................ 186
6.3 REFERENCES ............................................................................................................................... 187
13
LIST OF FIGURES
Figure 1.1 DNA polymerase III holoenzyme (dimer form) at a replication fork. ........................ 22
Figure 1.2 The τ and γ subunits of DNA pol III. .......................................................................... 30
Figure 1.3 The process of loading the β clamp onto the primer-template DNA duplex. ............. 31
Figure 1.4 Residue substitutions in the β clamp that are implicated in interactions with UmuD
(left), UmuD′ (middle) and α subunit of Pol III (right) (105). .......................................... 35
Figure 1.5 Regulation of SOS induced genes after DNA damage. ............................................... 43
Figure 1.6 Model of full-length UmuD (172) and crystal (169) and NMR (168) structures of
UmuD′. .............................................................................................................................. 46
Figure 1.7 Polymerase switching in response to DNA damage.................................................... 54
Figure 2.1 Models of α and UmuD. .............................................................................................. 78
Figure 2.2 UmuD binds wild-type α with a Kd = 1.1 μM. ............................................................ 88
Figure 2.3 UmuD binds α at two distinct binding sites. ............................................................... 89
Figure 2.4 The presence of α inhibits UmuD cleavage. ................................................................ 92
Figure 2.5 The α truncation 1-280 inhibits UmuD cleavage. ....................................................... 94
Figure 2.6 Binding to α alters the conformation of UmuD. .......................................................... 96
Figure 2.7 Fluorescence resonance energy transfer analysis shows that UmuD disrupts the
interaction between the α polymerase, labeled with Alexa Fluor 647 C2-maleimide, and
the β processivity clamp, labeled with Alexa Fluor 488 C5-maleimide. ........................ 100
14
Figure 3.1 Diagram of DNA pol III α, with domain labels within the boxes and known
interaction sites above the boxes (sequence numbering shown below). ......................... 113
Figure 3.2 DNA pol III α binding to ssDNA characterized with single molecule force
measurements. ................................................................................................................. 116
Figure 3.3 The fraction of ssDNA bound by α decreases with increasing UmuD concentration.
......................................................................................................................................... 122
Figure 3.4 Docking model predicts residues involved in the interaction between DNA pol III α
and UmuD. ...................................................................................................................... 123
Figure 3.5 UmuD variants are structurally stable and enzymatically active. ............................. 125
Figure 3.6 Binding curves between pol III α and UmuD variants measured by tryptophan
fluorescence quenching. .................................................................................................. 126
Figure 3.7 UmuD variants have compromised ability to disrupt α binding to ssDNA............... 128
Figure 4.1 5′ and 3′ biotin-labeled ssDNA was constructed using exonucleolysis by T7 DNA
polymerase. ..................................................................................................................... 145
Figure 4.2 SSB inhibits primer extension by DNA pol III α. ..................................................... 146
Figure 4.3 SSB binds DNA pol III α with a dissociation binding constant, KD, of 1.8 ± 0.4 µM.
......................................................................................................................................... 147
Figure 4.4 The structure (9) of the SSB homo-tetramer with the SSB variants used in this work
indicated. ......................................................................................................................... 149
Figure 4.5 SSB D91N disrupts the interaction with DNA pol III α but not with ssDNA. ......... 150
Figure 4.6 SSB binds the C-terminal domain of DNA pol III α subunit. ................................... 152
15
Figure 4.7 SSB F61A and SSB D91N only minimally inhibit replication by DNA pol III α. ... 154
Figure 4.8 DNA pol III α stabilizes the ssDNA/SSB interaction. .............................................. 156
Figure 5.1 Wild-type SSB binds the umuD gene products. ........................................................ 170
Figure 5.2 Equilibrium dissociation constant KD between UmuD or UmuD′ and SSB variants
measured by tryptophan fluorescence quenching. .......................................................... 172
Figure 5.3 Overproduction of UmuD but not UmuD′ suppresses the temperature sensitive
conditional lethality phenotype of the ssb-113 allele. .................................................... 174
Figure 5.4 The interaction between SSB and UmuD does not affect the dynamic nature of the N-
terminal arms of UmuD. ................................................................................................. 176
Figure 6.1 The umuD gene products regulate E. coli DNA replication in the presence of DNA
damage. ........................................................................................................................... 183
16
LIST OF TABLES
Table 1.1. Components of DNA polymerase III. .......................................................................... 24
Table 2.1 Equilibrium dissociation constants Kd (µM) for binding of α truncations to UmuD
variants. ............................................................................................................................. 98
Table 5.1 Strains and plasmids used. .......................................................................................... 167
17
LIST OF ABBREVIATIONS
~ Approximately
° Degrees
µL Microliter
µM Micromolar
A Alanine
A Adenosine
Å Angstrom
Ala Alanine
Asn Asparagine
Asp Aspartic Acid
ATP Adenosine triphosphate
BMH Bis-maleimidohexane
BSA Bovine serum albumin
C Cytidine
CaCl2 Calcium Chloride
CD Circular Dichroism
CV Column volume
Cys Cystiene
D Aspartic Acid
DNA Deoxyribonucleic acid
DNA pol I DNA polymerase I
DNA pol III DNA polymerase III
DNA pol IV DNA polymerase IV
DNA pol V DNA polymerase V
DNA pol β DNA polymerase β
dsDNA Double-stranded DNA
DTT Dithiothreitol
E. coli Escherichia coli
EDTA Ethylenediaminetetraacetic acid
EPR Electron paramagnetic resonance
F Phenylalanine
FRET Fluorescence resonance energy transfer
G Glycine
G Guanosine
Gly Glycine
HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
hr Hour
I Isoleucine
18
Ka Equilibrium association constant
kb Kilobase
KD Equilibrium dissociation constant
kDa Kilodaltons
L Leucine
Leu Leucine
Lys Lysine
Met Methionine
min Minute
mL Milliliter
mM Millimolar
mRNA Messenger RNA
N Asparagine
NaCl Sodium Chloride
nM Nanomolar
nm Nanometer
NMR Nuclear Magnetic Resonance
nt/s Nucleotides per second
OB Oligonucleotide/oligosaccharide binding
P Proline
PAGE Polyacrylamide Gel Electrophoresis
PHP Polymerase and histidinol phosphatase
pM Picomolar
pN Piconewtons
Pro Proline
Q Glutamine
RNA Ribonucleic Acid
S Serine
SDS Sodium dodecyl sulfate
Ser Serine
SSB Single-standed DNA binding protein
ssDNA Single-stranded DNA
T Tyrosine
T Thymidine
TLS Trans-lesion Synthesis
Tm Melting temperature
Ts Temperature Sensitive
UV Ultraviolet
V Valine
Val Valine
W Tryptophan
Y Tyrosine
19
CHAPTER 1: Overview of DNA Replication in Escherichia coli
Portions of this chapter were originally published by Springer as:
"Polymerase switching in response to DNA damage" Jaylene N. Ollivierre, Michelle C. Silva,
Jana Sefcikova, and Penny J. Beuning in Biophysics of DNA-Protein Interactions: from single
molecules to biological systems M.C. Williams and L. J. Maher, III, Eds., Springer, New York,
NY (2010).
It is reproduced here with permission from Springer:
DATE: June 28, 2013
SPRINGER REFERENCE
Biophysics of DNA-Protein Interactions
ISBN 978-0-387-92807-4
Biological and Medical Physics, Biomedical Engineering 2011,
pp 241-292 “Polymerase Switching in Response to DNA Damage” by Jaylene N. Ollivierre,
Michelle C. Silva, Jana Sefcikova, Penny J. Beuning
YOUR PhD Thesis:
University: Northeastern University, Boston MA
Title: "DNA replication by Escherichia coli DNA polymerase III is regulated by the
umuD gene products"
Dear Ms. Silva,
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1.1 E. coli DNA Polymerase III
DNA replication requires the coordination of many different proteins to accomplish the goal of
simultaneous replication of the two antiparallel stands of DNA. This process is tightly regulated
so that DNA is replicated in a timely and accurate manner. Moreover, DNA replication is highly
21
processive, which allows efficient replication of over three million base pairs in every E. coli cell
cycle.
In E. coli, DNA polymerase III (DNA pol III) is responsible for the majority of DNA replication
(Figure 1.1). Each cell expresses approximately ten copies of DNA pol III core (see below) (1).
Along with auxiliary proteins, DNA pol III semi-discontinuously replicates DNA at a speed of
approximately 1 kilobase (kb) per second (2), making less than one error in approximately 105
nucleotide additions (1, 3). In the presence of proofreading and mismatch repair, the error
frequency is approximately 10-10
per base pair (1, 4). At each replication fork, the DNA pol III
core acts as an asymmetric dimer (5-9): one monomer acts in the continuous replication of the
leading strand and another acts in the discontinuous replication of the lagging strand (Figure 1.1).
Even though only two polymerase cores are needed to replicate DNA, it has been shown that the
replisome may contain three DNA polymerase cores. The third polymerase core may function on
the lagging strand or serve as a spare polymerase, able to replace either polymerase when needed
(10).
22
Figure 1.1 DNA polymerase III holoenzyme (dimer form) at a replication fork.
The two polymerase cores (green), which are tethered to the β clamps (light blue), contain the
three subunits α, , and . The γ complex (blue), assembled with two τ subunits, couples the
polymerization of both the leading strand and the lagging strand. The single-stranded lagging
strand is threaded through DnaB helicase (orange) and is coated with SSB (gray). Also shown is
the primase (light green) that synthesizes the RNA primers (red) on the lagging strand. The ψ
and χ subunits are not shown but would connect the γ complex to SSB.
E. coli DNA pol III consists of ten subunits (2, 6, 9, 11) that can be classified into three
subassemblies: the core, the clamp and the clamp loader complex (9, 12) (Table 1.1). The core is
composed of three subunits: , , and . The subunit contains the polymerase activity and is
responsible for synthesizing DNA. The processivity of the isolated subunit, defined as the
23
number of nucleotides incorporated into the nascent DNA per association event, is low (1-10
nucleotides) compared to >50 kb for the holoenzyme (1, 13-17). The subunit is a 3′ to 5′
exonuclease that is responsible for the proofreading capability of the core. In the absence of ,
the frequency of mutations due to misincorporations during replication increases by
approximately forty-fold (18). When coupled, the exonuclease and polymerase activities of the
and subunits increase significantly (18). The third component of the core, , is not required for
high processivity (19).
The clamp, also known as the processivity clamp, is the major contributor to the processivity
of DNA pol III (1). The β clamp encircles DNA and tethers the subunit to its DNA substrate
(Figure 1.1). The β clamp is loaded onto DNA by a complex known as the clamp loader, which
is composed of six subunits: , , , ′, , (Table 1.1) (20). The subunits coordinate
replication on both strands by coupling the polymerase cores to the clamp loader complex (9,
21). When is coupled to the core, processivity increases approximately six-fold (1). The
subunit is an ATPase. Along with and ′, is responsible for loading the clamp onto DNA (2,
11). The two other subunits, and , bind single stranded DNA binding protein (SSB) and help
regulate replication on the lagging strand (22, 23). The subunit also has a role in clamp loading
(24).
24
Table 1.1. Components of DNA polymerase III.
Mass
(kDa)
Gene
Function
References
Polymerase Core
130 dnaE polymerase (25-28)
27.5 dnaQ (mutD) 3′-5′ exonuclease (18, 29-36)
10 holE stabilizes the core (33, 36-39)
Clamp Loader (20, 40, 41)
71 dnaX coordinates replication (9, 21, 42-51)
47.5 dnaX ATPase (47, 52-54)
35 holA “wrench”; opens clamp (52, 55)
′ 33 holB mediator between & (52, 56)
15 holC binds SSB (22, 23, 57, 58)
12 holD bridges γ complex & (22, 57)
Clamp
40.6 dnaN clamp (11, 28, 40, 59-63)
1.2 The Polymerase Core
The DNA pol III core includes the polymerase and proofreading exonuclease activity of the
replisome. Alone, the core can replicate DNA at a rate of approximately 20 nucleotides per
second (nt/s) with a processivity of 11 nucleotides, values much lower than the entire replisome
(2, 11). The following is an in-depth description of each of the subunits of the core.
1.2.1 The Polymerase Subunit,
25
The subunit, encoded by the dnaE gene and a member of the polymerase C family, is a protein
of approximately 130 kDa, containing several distinct domains. Although this polymerase has
been studied for decades, the crystal structure of the subunit of E. coli was only solved recently
(25). Like other polymerases, the structure resembles a right hand with three characteristic
domains: the palm, the fingers, and the thumb domains (64-66). The palm domain contains the
active site of the polymerase consisting of three aspartic acid residues: Asp401, Asp403, and
Asp555 (67). This domain is similar to the palm domains of polymerases in the X family,
especially to that of DNA pol (25). Modeling DNA onto the E. coli DNA pol III α structure
using the human DNA pol co-crystal structure with DNA (68) as the modeling template,
showed that the DNA strand collides with a short -helix in the palm domain in a sterically
unfavorable interaction (25). This suggests that although DNA pol III and pol have similar
active sites, there may be differences in how they bind DNA.
The finger domain includes four sub-domains: the index finger, the middle finger, the ring
finger, and the little finger. These sub-domains are responsible for binding the incoming
nucleotide. The thumb domain guides the newly formed DNA duplex as it leaves the active site.
The Polymerase and Histidinol Phosphatase (PHP) domain is located in the “wrist” position,
relative to the hand of the polymerase domain. The exact role of this domain is unknown but
because of the domain’s sequence similarity to histidinol phosphatases, it was proposed to
possess pyrophosphatase activity for the pyrophosphate produced during replication (69). Based
on the crystal structure of DNA pol III , this domain is unlikely to harbor such activity (25).
However, it has been demonstrated that the Thermus thermophilus DNA pol III subunit
contains a Zn2+
-dependent 3′ to 5′ exonuclease activity (70).
26
The C-terminal domain is not present in the crystal structure of E. coli DNA pol III . This
domain, which is located C-terminal to the tip of the little finger domain, includes an
oligonucleotide/oligosaccharide binding (OB) fold (25, 26) and the binding sites for both the τ
subunit (49, 71) and the β clamp (28, 50). The recently-solved structure of DNA pol III from
Thermus aquaticus (26) includes this C-terminal domain, consisting of an OB fold. The OB fold
domain consists of five -strands arranged in a barrel, similar to that of other OB folds (72).
The OB fold domain of α has been shown to bind single-stranded DNA (ssDNA) specifically
(73). Compared to other ssDNA binding proteins, the OB fold domain of the α subunit is
somewhat unusual in that it does not actively melt DNA, but rather binds to ssDNA that is pre-
formed, in this case by force-induced melting (73). Along with the rest of the polymerase, this
function provides insight into the regulation of DNA pol III , discussed below.
A co-crystal structure of T. aquaticus DNA pol III bound to primer-template DNA and an
incoming deoxynucleoside 5′-triphosphate has been determined with a resolution of 4.6 Å (27).
When compared to the structure of T. aquaticus DNA pol III without DNA, it is possible to see
significant movements of the thumb, finger, and -binding domains. These movements position
the protein on the DNA, allowing for the interaction with the DNA backbone at the minor
groove. This structure also indicates that the DNA and incoming nucleotide bind in a similar
fashion to that of DNA pol . The C-terminal domain undergoes an approximately 30o rotation
putting the OB fold in position to bind the single stranded template DNA. The internal -binding
motif also seems to be correctly positioned in the structure with DNA in order to bind to the
hydrophobic pocket on the clamp (27, 28).
27
The structure of a ternary complex of PolC, the replicative polymerase of the gram-positive
bacterium Geobacillus kaustophilus, with primed DNA and an incoming dideoxynucleoside
substrate has been solved to 2.4 Å resolution (74). Unlike the DNA pol III polymerase, PolC
contains an intrinsic 3′ to 5′ exonuclease domain as an insertion within the PHP domain. Instead
of being located in the C-terminal domain, the OB fold of PolC is located N-terminal to the palm
domain and is positioned so it could bind the single-stranded template strand approximately 15-
20 nucleotides away from the polymerase active site. The thumb domain contains -strands that
bind DNA in the minor groove, possibly allowing for the detection of mismatched base pairs
after incorporation (74). Flexibility in the palm domain suggests a large conformational change
upon DNA binding (74, 75).
1.2.2 The Subunit
The subunit, a 27.5-kDa protein encoded by the dnaQ (also known as mutD) gene, is
responsible for the 3′ to 5′ exonuclease activity of the polymerase core (1) and forms a tight
complex with DNA pol III α. Whereas in E. coli the polymerase and exonuclease reside on two
different polypeptides, in other cases, the exonuclease activity and the polymerase activity are
part of the same polypeptide, as in gram positive PolC and in DNA pol I (64, 66, 74), another
eubacterial polymerase. Such an interaction between and enhances the overall activity of
each protein. In fact, it has been shown that has greater polymerase activity in complex with
than alone (18). The exonuclease activity of the subunit is also substantially stimulated within
the complex (18).
28
Although the structure of the N-terminal domain of has been determined by X-ray
crystallography (29) and by NMR (76), no structures for the full-length subunit have been
determined due to the difficulty in obtaining large amounts of pure protein. The structures
include the 186 N-terminal residues of responsible for its exonuclease activity. The 57 residues
that are not present in the structure include the C-terminal domain, which contains a flexible
linker that has been shown to bind to the subunit (30-33). The subunit binds in the PHP
domain located in the N-terminal domain (77).
1.2.3 The Subunit
The third subunit of the polymerase core is , the 10-kDa product of the holE gene (1). The
subunit binds to close to the active site (37, 38), although it is unlikely to play a direct role in
exonuclease activity (36). Extensive hydrophobic surfaces define the interactions between θ and
ε (36, 39, 78). The θ subunit has not been shown to bind directly to (38, 79), but it may have a
stabilizing effect on the : complex. Such a function is supported by the results of a series of
yeast two-hybrid experiments indicating that the interaction between and is strengthened in
the presence of (80). The subunit also seems to enhance the exonuclease activity of the
subunit by stabilizing ε (38, 81). Deletion of θ results in a slight increase in the spontaneous
mutation frequency of E. coli (80). In biochemical experiments, the exonuclease activity of
I170T/V215A double mutant was not substantially stimulated in the presence of either α or
(82). Upon addition of both α and , however, activity of the I170T/V215A variant was
stimulated (82).
29
1.3 The Clamp Loader Complex and the Clamp Subunit
The clamp loader complex consists of at least two subunits, up to three subunits, and one
each of the , ′, , and subunits (2, 9, 11). As mentioned above, DNA pol III can be
assembled as a trimer (10). In this case, three subunits are present in the absence of subunits.
Although not required for the clamp loading process, the subunits play a critical role in
managing replication at the fork (9, 11). The subunits, closely related to , bind ATP, and
facilitate loading the clamp onto DNA (11, 40). The and ′ subunits are directly involved
with the loading of the clamp, as the “wrench” and ′ as the mediator between and (40, 55).
1.3.1 The Complex: ′:1:2:3:; Loading the Clamp
The dnaX gene encodes both and . The subunit is the full-length product of the gene and is
produced due to a -l frameshift, which causes a stop codon to be inserted prematurely, forming
the shorter product, (47, 83, 84). This frameshift is caused by two factors: a heptanucleotide
sequence that induces frameshifts and a downstream RNA stem-loop structure (Figure 1.2A).
Therefore, consists of only the first three of the five domains (Figure 1.2B). These three
domains contain the ATPase site and so both and are ATPases. In the crystal structure of the
complex, containing γ3′ (52), it is possible to distinguish these domains. Domains I and II
30
contain the nucleotide binding site in which ATP binds at the interface between the subunits and
Domain III forms a circular collar with the other subunits of the γ complex.
Figure 1.2 The τ and γ subunits of DNA pol III.
(A) The segment of the dnaX mRNA, responsible for coding the γ and τ subunits. The two
factors that cause the -1 frameshift are a heptanucleotide sequence (bold) and a downstream
stem-loop. As a result, the γ subunit consists of the first 430 residues of the dnaX gene product
shared with the τ subunit, followed by a glutamic acid residue and a stop codon. (B) A side-by-
side comparison of the domains of the τ and γ subunits. The γ subunit, the shorter protein,
contains only the first three domains of the entire gene product. These domains include the
ATPase active site and the collar domain. Domains IV and V of the τ subunit contain the binding
sites for both the pol III α subunit and DnaB.
A crystal structure of the first 243 residues of γ (Domains I and II) solved with and without
nucleotides (53), shows a conformational change upon nucleotide binding, suggesting a
mechanism for binding to the clamp and ATP hydrolysis (Figure 1.3). In the crystal structure of
the complex (52), the subunits, together with and ′, are arranged in a heptameric complex
31
resembling an opened ring in the order ′:1:2:3: (Figure 1.3). At each interface there is an
ATP binding site (52). Such a configuration suggests that upon ATP binding, the complex
undergoes a conformational change from a closed state (without ATP) to an open state (with
ATP) (53). The subunit is then free to interact with the clamp.
Figure 1.3 The process of loading the β clamp onto the primer-template DNA duplex.
ATP binds the γ complex, causing a conformational change from a closed state to an open state,
which allows the N-terminal domain of the δ subunit (dark blue) to bind to the β clamp. This
interaction disrupts the dimer interface of the β clamp, creating an opening for the primer-
template DNA duplex to enter. The bound ATP is then hydrolyzed, causing the γ complex to
relax back to its closed state. The δ subunit then releases the β clamp, re-establishing the dimer
interface.
The ′ subunit, the 33-kDa member of the complex encoded by the holB gene, acts as a
mediator between and (1). Although sequence and structure alignments of ′ and suggest
that these two subunits are homologous, ′ does not have a functional nucleotide binding domain,
32
as shown in the crystal structure of the ′ subunit (56). This crystal structure shows that ′, like ,
consists of three consecutive domains organized in a C-shaped architecture. The first domain
consists of a sheet with five parallel strands surrounded by six -helices similar to the
nucleotide binding domain of RecA (56, 85). The ′ subunit contains a zinc-binding module
whose function is unknown but because it is found on what resembles a phosphate binding loop,
it may help couple DNA binding with ATP hydrolysis by the clamp loader (56).
The subunit, a 35-kDa product of the holA gene, is considered the “wrench” of the clamp
loader complex because its binding to the clamp causes a spring-like conformational change
(55). This allows the dimeric ring of the clamp to transition from its default closed state, where
both dimeric interfaces are intact, to an open state, where only one dimeric interface exists
(Figure 1.3) (55, 86-88). This spring-like mechanism facilitates loading of the clamp onto the
primer-template DNA duplex (2, 9, 11, 55). The “wrench”-like interaction between δ and the
clamp is shown in a crystal structure involving the N-terminal 140 residues of and a variant
form of the clamp that cannot dimerize (55). Attempts to crystallize with the full-length
clamp dimer have been unsuccessful (55). In fact, binds to the monomer form approximately
fifty-fold more tightly than to the dimer. The binding interface between these two subunits is
contained in a hydrophobic tip of and a hydrophobic pocket on the surface of the clamp that
is also known for binding other components of the replisome (55, 63, 89) and DNA polymerases
(90-93). When the structure of the monomer form of the clamp without was compared to
that with , a distortion of the curvature of the clamp that results in a ~15 Å opening can be
seen (55). Such an opening is enough to allow ssDNA into the center of the clamp (40, 55).
33
The crystal structures of the clamp loader subunits suggest a multistep process for loading the
clamp onto a primed DNA strand (9, 11, 40), powered by the binding of two ATP molecules (54)
(Figure 1.3). When ATP binds to the closed ring-shaped complex, a conformational change
takes place disrupting the interaction between and ′. The subunit is then free to bind to the
hydrophobic pocket of the clamp dimer. This binding event distorts the dimer interface
allowing DNA to enter the clamp, creating an opened ring-like structure consisting of the opened
γ complex and clamp. Site-directed mutagenesis (94), electron microscopy (95), and X-ray
crystallography (41) experiments have shown that DNA can bind to the center chamber of the
complex. Once ATP is hydrolyzed, the dimer interface of the clamp is restored, allowing the γ
complex to relax back to its closed state. This causes the clamp loader to release its hold on the
clamp, the rate-limiting step of the clamp loading process (96). The clamp and primer-template
DNA duplex are then competent for polymerase loading (97).
The recent crystal structure that shows the clamp loader complex coupled to DNA (41) supports
the notched screw-cap model for clamp loading as described above. The structure shows that the
complex forms a right-handed spiral-like structure and is loaded onto dsDNA like a cap,
allowing ssDNA to exit through a slit formed by the complex. In this structure, the complex
does not recognize DNA via both strands of the primer:template complex as previously thought
(95, 98). Rather, recognition occurs on the phosphate backbone of the template strand alone, and
at just the 3′ nucleotide of the primer strand (41). This allows for both DNA and RNA primers to
be recognized, the mechanism of which was previously unclear (41, 99).
34
1.3.2 The and Subunits
The and subunits, products of the holC and holD genes, respectively, are subunits of the
clamp loader complex (2, 9, 11, 100). These two subunits function during replication on the
lagging strand (22, 23). Because DNA polymerases replicate DNA only in the 5′ to 3′ direction,
the lagging strand must be replicated in a direction opposite to the movement of the replication
fork. This is accomplished by replicating DNA in ~1 kb fragments, called Okazaki fragments
(101). As a result, an abundance of ssDNA is present and coated with SSB (Figure 1.1). The
and subunits help to coordinate replication on the lagging strand (22, 23). The subunit acts
as a mediator between the complex and by binding the collar domains (Domain III) of the
and subunits (41, 57). The subunit binds to the C-terminal domain of SSB (23), thereby
coupling it to the replisome and allowing the clamp loader complex to be in close proximity to
the primer-template DNA on the lagging strand (58). Together, the and subunits constitute a
tightly held complex that increases the affinity of and for and ′ (57). The subunit also
serves to increase affinity of the clamp loader for the β clamp in the presence of ATPγS (24).
1.3.3 The Clamp Subunit
Once it is loaded onto the primer-template DNA duplex, the clamp (dnaN) has two specific
roles. It tethers the polymerase to the DNA and contributes to the mobility of the polymerase on
the DNA strand. The clamp, a ring-shaped homodimer (Figure 1.4) (61, 62), is the major
contributor to processivity (1), by allowing the polymerase to maintain close contact with the
DNA. In order to facilitate processive DNA synthesis, the clamp must remain bound to the DNA
35
with or without the polymerase present, which has been shown (59, 102). Other studies show that
the clamp can remain on a circular plasmid two to three times longer than the time it takes for
cells to divide (60). With the use of single molecule fluorescence spectroscopy, it was found that
the diffusion constant for the clamp is at least three orders of magnitude lower than for
diffusion through water. The clamp seems to be held at the 3′ end of the primer in the presence
of SSB (103). The relatively slow motion of the clamp may be due to the attractive interactions
of positively charged residues on the inside of the clamp that come in contact with the negatively
charged phosphate backbone of the DNA (103, 104).
Figure 1.4 Residue substitutions in the β clamp that are implicated in interactions with
UmuD (left), UmuD′ (middle) and α subunit of Pol III (right) (105).
Positions in green are important to the interaction between the β clamp and all three proteins
listed above. Positions in purple exhibit only a modest effect. Substitutions that result in an
increase or decrease in the affinity of UmuD and UmuD′ for the β clamp by formaldehyde or
glutaraldehyde cross-linking are shown in red. Residue Lys74 shown in grey (left) cross-links to
UmuD using formaldehyde. The hydrophobic channel is shown in brown (residues Leu177,
Pro242, Val247, Val360, Met362) (55) (106), while the rim interaction residue Leu98 is shown
in black. Structures were generated using VMD (107) and coordinates for β (2POL) from the
PDB (61).
36
Binding experiments of the clamp with DNA pol III and other components suggest that the
same hydrophobic region on the surface of to which δ binds is also responsible for binding
DNA pol III and other DNA polymerases (55, 63, 90-93, 105, 108, 109). It has also been
suggested that the DNA pol III C-terminus binds to the clamp (see below) and binds . These
observations suggest that the polymerases, , and compete with one another for binding to the β
clamp, creating a mechanism for polymerase loading and switching on the β clamp (46, 50, 63,
110).
An internal binding site ( residues 920-924), rather than the 20 C-terminal residues, was shown
to be responsible for the interaction between the clamp and (28). Replacement of all residues
in this internal binding site eliminated binding to the clamp, but binding between the subunit
and was not affected. When an analogous set of mutations was made in the C-terminal binding
site of , the β clamp still showed affinity for the α subunit (28). In fact, participated in
processive replication even when the entire C-terminal binding site was removed. The absence of
the C-terminal peptide, however, specifically effected the interaction of with . These findings
suggest that the clamp does not bind to the C-terminus of , but instead to an internal site. This
discrepancy might be due to the use of an variant with a relatively large C-terminal truncation
in the previous study (28, 50), rather than site-directed mutant variants or more modest deletions
(28).
There is also evidence that two different polymerases can simultaneously bind to the clamp. The
subunit, a homodimer, has two hydrophobic pockets per functional protein, thus allowing it to
37
bind two DNA polymerases (11, 105, 111, 112). Such a situation allows replication to alternate
between the two DNA polymerases without the need for dissociation from the β clamp. This
“toolbelt” hypothesis allows for high processivity, even under conditions where multiple
polymerases are used. The ability of the clamp to bind both DNA pol III and pol IV (a Y family
polymerase) was investigated using fluorescence resonance energy transfer (FRET). Proximity
of the pol III α subunit and pol IV was detected in a β-clamp-dependent manner, providing
experimental evidence that two different DNA polymerase molecules can simultaneously bind to
the β clamp (112). The crystal structure of with the C-terminal little finger domain of pol IV
(106) showed that the pol IV polymerase domain is angled off to the side providing enough room
for another polymerase, such as DNA pol III, to bind. Modeling the full length structure of Dpo4
(113) (a Pol IV homolog from Sulfolobus solfataricus), containing a primer-template duplex and
incoming nucleotide, onto that of the little finger structure of E. coli pol IV with the clamp,
showed that when bound in this position, the polymerase likely does not have access to the DNA
strand (106). The little finger domain can likely undergo a conformational change (106),
positioning the polymerase onto the DNA substrate.
Another model for polymerase switching is “dynamic processivity” (114). This model involves
polymerase replacement without affecting apparent overall processivity. Such a scheme was
observed during bacteriophage T4 DNA replication (114). The addition of a catalytically inactive
variant D408N of gp43, the T4 DNA polymerase, to an active replication fork, arrested
replication while still retaining wild-type-like affinity for DNA and the clamp (gp45). This
observation suggested that the active polymerase is quickly (<1 min) replaced by the inactive
38
variant (114). This dynamic processivity of polymerases implies that multiple replicative
polymerases may be required to replicate normal, undamaged DNA.
1.3.4 The Subunit
Although similar to the subunit, the role of the subunit is distinct from the remainder of the
complex. As a central component of the DNA pol III replisome, coordinates replication on both
strands by connecting the subassemblies of the core and clamp as τ binds to the polymerase
(48) and the helicase DnaB (45, 48), and is part of the clamp loader complex (57). Numerous
distinct roles have been assigned to , as summarized in a review (9). The roles of the subunit
in replication are to: 1) coordinate replication on both strands; 2) bind to DnaB; 3) prevent
premature removal of the clamp; and 4) function in the processivity switch.
When DNA polymerase III′ (core + τ) was first isolated (21), it was observed that two
polymerases were coupled by two subunits, suggesting that may be a key factor in
coordinating replication (Figure 1.1). This hypothesis was tested by varying the concentration of
(42). At low concentrations of , shorter DNA fragments were observed upon agarose gel
electrophoresis analysis of replication reaction samples (42). These results suggest that the two
polymerase cores must be coupled through in order to effectively replicate both the lagging and
leading strands.
DNA pol III binds through the C-terminal domain (28, 50) with an equilibrium
dissociation constant (KD) of 4 nM (48). The binding site on was determined to be at the C-
39
terminal domain (45), which is not part of (Figure 1.2B). Along with determining the NMR
solution structure for this domain (51), combinatorial binding studies were conducted to
determine which residues bind to (49). It was concluded that the 18 C-terminal residues of τ
are required for binding to α (49).
Four different reconstituted clamp loader complexes, γ3', 1γ2', 2γ1', 3',
have similar rates of loading the clamp onto DNA (10). The complex containing 3 may
coordinate three DNA polymerases at the replication fork. A “triple-polymerase” model has been
proposed in which two of the pol III cores function on the lagging strand to synthesize Okazaki
fragments. Alternatively, one pol III core is utilized on the lagging strand with the third pol III
core held “in reserve” off of the DNA (10). The latter model suggests a possible switching
mechanism between high and low fidelity DNA polymerases (10).
The affinity of the subunit for DnaB (45) also seems to affect the rate at which the replication
fork proceeds (8, 43). DnaB, a hexameric helicase, unwinds DNA ahead of the fork while
encircling only the lagging strand (Figure 1.1) (8). Without the replisome, the helicase unwinds
DNA at a rate of approximately 35 nt/s, similar to that in the presence of the complex without .
When is added, the rate increases to at least 400 nt/s, approaching that of DNA pol III (43, 45).
The subunit may also indirectly prevent the premature removal of the clamp by the
complex, thereby maintaining processivity. The length of DNA produced in the absence of τ is
directly proportional to the concentration of and inversely proportional to the concentration of
the clamp loader complex (44). This suggests that the clamp removal function is inhibited
40
during normal replication in the presence of the γ complex, allowing the clamp to stay on the
DNA.
Discontinuous replication of the lagging strand requires that DNA pol III must constantly
dissociate from and re-associate with different β clamps. Such a cycle is known as the
processivity switch and is thought to involve (46). It has been shown that the polymerase core/τ
complex must complete the newly formed strand, leaving only a nick, before the switch can be
activated (46). In such a scenario, loses affinity for when primed DNA is present. Then when
replication is completed, gains affinity for , releasing from the clamp (46). The ssDNA
binding function of α also appears to play a role in this process (110).
1.4 The Single-Stranded DNA Binding Protein, SSB
E. coli SSB plays a significant role in DNA replication, recombination, and repair by binding
ssDNA present during these DNA processing events, preventing ssDNA from forming secondary
structures and protecting the ssDNA from chemical and nucleolyitc attacks (115-117). SSB is a
homo-tetramer with a D2 axis of symmetry (118). Each monomer contains an N-terminal
globular domain and a disordered C-terminal tail (119).
The N-terminal globular domain of SSB contains an oligonucleotide/oligosaccharide-binding
(OB) fold that binds ssDNA via stacking interactions with one phenylalanine (F61) and two
41
tryptophan (W41 and W55) residues of SSB (120-125). ssDNA binds SSB by wrapping around
the homo-tetramer in two binding modes: (SSB)35 and (SSB)65 (126). In the (SSB)35 binding
mode, ssDNA only wraps around two monomers of the homo-tetramer, favoring high
cooperativity. On the other hand, the (SSB)65 binding mode wraps around all four monomers and
favors limited cooperativity (126).
In addition to binding and protecting ssDNA, SSB also serves as a hub for at least 15 proteins,
such as proteins in homologous recombination, DNA pol II and pol V in damage tolerance, and
the χ subunit of DNA pol III in replication, involved in numerous DNA processing pathways
(127). Most of these proteins have been found to bind the C-terminal tail suggesting that SSB
uses this tail to facilitate interactions with other proteins involved in DNA maintenance (128).
The temperature sensitive allele ssb-113 is associated with the mutation P177S (129) and impairs
DNA replication at temperatures higher than 30°C (130, 131). The P177S mutation has been
shown to disrupt the interaction with proteins that bind this C-terminal tail (127).
SSB, as well as protecting ssDNA present at the replication fork, also has been observed to
directly impact replication activity of DNA pol III. SSB increases processivity of the polymerase
when all the subunits of DNA pol III are present. In contrast, when only the core subunits are
present, SSB inhibits replication (13). This inhibition is alleviated when the χ subunit of the
clamp loader complex is present due to a direct interaction between χ and the C-terminal tail of
SSB (22, 132-134).
42
1.5 DNA Damage disrupts DNA Replication
DNA pol III function requires finely tuned interactions of multiple proteins for efficient and
accurate DNA replication. When this complex encounters non-canonical DNA structures,
including DNA damage, it is not equipped to replicate them. DNA damage is ubiquitous, arising
from numerous exogenous and endogenous sources. For example, it is estimated that 10,000
abasic sites are formed per human cell per day (135). The outcome of the replisome encountering
DNA damage in the template may depend on whether the damage is encountered during leading
strand or lagging strand synthesis, but typically the rate of progress of replication is decreased
(136, 137). A lesion in the leading strand has been observed to slow progression of the
replication fork, but was not observed to interfere with lagging strand replication (138). On the
other hand, a lesion in the lagging strand does not block overall progression of the replication
fork. Instead, the lagging strand DNA polymerase appears to re-initiate downstream of the lesion
at the next Okazaki fragment, leaving a gap (138-141). Indeed, it has been shown that replication
can restart downstream of obstacles, even in the case of leading strand synthesis (142).
1.6 Specialized DNA Polymerases facilitate DNA Damage Tolerance
In E. coli and some other bacteria, DNA damage and other stresses lead to induction of the SOS
response (135). Stalling of DNA replication at damaged sites results in the accumulation of
single stranded DNA (ssDNA). RecA polymerizes on the ssDNA, forming a nucleoprotein
43
filament that serves as the inducing signal for the SOS response. As the result, the expression of
at least 57 genes is induced (143, 144). SOS-regulated genes code for proteins involved in
regulation of cell division, nonmutagenic repair of chemically modified DNA or in damage
tolerance mechanisms, which can be mutagenic or error prone (Figure 1.5) (135, 143, 145, 146).
Figure 1.5 Regulation of SOS induced genes after DNA damage.
The SOS response is induced by the formation of a RecA nucleoprotein filament on single
stranded DNA (RecA*). This stimulates the auto-proteolysis of the LexA repressor which leads
to the induction of at least 57 genes. Among these genes are Y family polymerases pol IV (DinB)
and pol V (UmuD′2C). UmuD2 undergoes RecA* facilitated cleavage of its N-terminal 24 amino
acids to yield UmuD′2, the form that is active in translesion synthesis. Lon and ClpXP proteases
play a role in regulating the levels of UmuD2, UmuD′2, and UmuC in the cell. ClpXP also
specifically targets UmuD′ in UmuDD′ heterodimers (not shown).
44
Three of the five known E. coli DNA polymerases are under SOS inducible regulation (145,
147). These SOS inducible DNA polymerases are pol II (polB), pol IV (dinB), and pol V
(umuDC; UmuD′2C). The latter two belong to the Y family of DNA polymerases, which are
characterized by their ability to perform translesion synthesis (TLS) on damaged DNA
templates, as well as their relatively low fidelity on undamaged DNA (148). Y family DNA
polymerases also lack intrinsic 3'-to-5' exonucleolytic proofreading and exhibit low processivity
(145, 148, 149).
Although high-resolution structures of E. coli Y family polymerases have not yet been
experimentally determined, the structures of Y family polymerases from Sulfolobus solfataricus
and Sulfolobus acidocaldaricus homologs provide insights into their function (113, 150, 151).
While there is no obvious sequence homology between replicative and Y family DNA
polymerases, the crystal structures of the latter reveal a similar right-hand structure of the
catalytic domain consisting of thumb, palm, and finger domains, common to other DNA
polymerases. Another domain, the little finger domain, is present in the Y family polymerases,
providing additional DNA binding contacts in the major groove (113, 148). This domain may be
responsible for both substrate specificity and processivity (152). Moreover, the O helix that is
responsible for high fidelity in the replicative polymerases (153-155) is not present in Y family
polymerases, suggesting a structural basis for the low fidelity of Y family DNA polymerases
while replicating undamaged DNA. The specialized ability of Y family polymerases to replicate
damaged DNA has been attributed to their loose, flexible active sites that accommodate aberrant
DNA structures (113, 148, 151). In addition, Y family polymerases have fewer contacts with
their DNA substrates than replicative DNA polymerases (113, 151). Although the crystal
45
structures show that the catalytic domains of Y family polymerases have similar overall folds,
these polymerases exert different efficiencies and fidelity in bypassing various DNA lesions
(148, 156).
1.6.1 Regulation of Y Family DNA Polymerases
Expression of the umuDC and dinB gene products is negatively regulated by the LexA repressor
as part of the SOS transcriptional response. LexA binds to a sequence in the operator region of
the genes (135, 157, 158). Derepression of the umuDC and dinB operons occurs when the RecA
protein binds to single-stranded regions of DNA that develop at replication forks that are stalled
by DNA damage (159). The RecA/ssDNA nucleoprotein filament serves as a coprotease to
facilitate cleavage of the LexA repressor (Figure 1.5). As the cellular concentration of LexA
diminishes, the genes whose expression is normally repressed by LexA are transcribed (135).
The umuDC genes are among the most tightly regulated SOS genes; the equilibrium dissociation
constant (KD) is 0.2 nM for LexA binding to the “SOS-box” in the promoter region (157). In
comparison, the KD values for LexA binding to the “SOS-boxes” of the recA and lexA genes are
estimated to be 2 nM and 20 nM, respectively (160). Immunoblotting assays have shown the
cellular steady-state levels of UmuD to be ~180 copies per uninduced cell and ~2400 copies per
cell under SOS induction (161). A single protein in a compartment with the volume of a typical
E. coli cell is present at a concentration of ~1 nM. The level of UmuC is approximately 12-fold
lower than UmuD with about 15 molecules per cell in the absence of induction and ~200
molecules of UmuC per cell under SOS induced conditions (161). There are ~250 molecules of
46
DinB per cell in the absence of induction and ~2500 molecules of DinB per cell after treatment
with the DNA damaging agent mitomycin C (162). It should be noted that the dinB gene is also
present on the E. coli F′ episome, and expression levels of DinB from the episome under
uninduced and induced conditions are approximately three-fold higher than from the
chromosome (162). Expression of the umuDC genes initially produces UmuD (139 amino acids)
which undergoes a RecA/ssDNA-stimulated autodigestion reaction after induction resulting in
UmuD' (115 amino acids) (Figure 1.5 and 1.6) (163, 164). UmuD is the predominant species for
the first approximately 20-40 min after SOS induction, after which UmuD′ is the predominant
species (165). UmuD proteins exist in solution as UmuD2 and UmuD'2 homodimers as well as
the UmuD-UmuD' heterodimer, which is more stable than either of the homodimers (166-169).
The Kd for UmuD2 dimerization is estimated to be in the low-pM range, so UmuD is likely to be
present in the cell as a dimer under most conditions (170, 171).
Figure 1.6 Model of full-length UmuD (172) and crystal (169) and NMR (168) structures of
UmuD′.
UmuD model shown in the trans, elbows down conformation (left). Crystal structure (middle)
and NMR structure (right) of UmuD′ in trans conformation. The N-terminal arms of UmuD′ are
cleaved between Cys24 and Gly25. Residues 1-24 are shown in magenta; residues 25 to 40 are
shown in blue. Active site Ser60 and Lys97 are highlighted in red and green, respectively.
47
SOS-induced mutagenesis is also regulated at the post-translational level. UmuD is functionally
inactive for facilitating TLS until it undergoes RecA/ssDNA-mediated cleavage to generate
UmuD' (164, 173, 174). Full-length UmuD also inhibits -1 frameshift mutagenesis by DinB
(175). Efficient cleavage of UmuD in vitro and in vivo was observed at elevated levels of
activated RecA, suggesting that TLS likely occurs when cells are under more severe
environmental stress (161, 173). The removal of the UmuD N-terminal 24 amino acids through
the cleavage of the Cys24-Gly25 bond occurs via an intermolecular pathway, that is, one
protomer of the dimer acts as an enzyme, while the other is the substrate (176). The crystal
structure of UmuD'2 revealed that the active site, consisting of conserved serine (Ser60) and
lysine (Lys97) residues, is found at the end of a cleft within the C-terminal globular domain of
the protein and these residues are poised for cleavage (Figure 1.6) (169). In the NMR structure
of UmuD'2 Ser60 and Lys97 are further apart and not correctly oriented for catalysis. It has been
suggested that the crystal structure of UmuD2 mimics the effect of the RecA/ssDNA
nucleoprotein filament as it serves to realign these residues, thereby activating UmuD2 for self-
cleavage (168, 177).
The two different forms of UmuD provide a temporal switch between accurate and mutagenic
phases of the cellular response to DNA damage (165, 178, 179). The combination of uncleaved
UmuD and UmuC specifically decreases the rate of DNA replication and increases resistance of
cells to killing by UV radiation (165, 180). Uncleaved UmuD2C improves DNA damage survival
by allowing time for error-free repair mechanisms to act before the combination of cleaved
UmuD'2 and UmuC (UmuD'2C, pol V) initiates potentially error-prone TLS (165). The
combination of UmuC and noncleavable UmuD (S60A) significantly delayed recovery of cell
48
growth after UV radiation (165, 179). Therefore, a model for a umuDC-dependent DNA damage
checkpoint in E. coli was proposed wherein a delay in DNA synthesis provides time for error-
free nucleotide excision repair to remove DNA lesions. TLS is then enabled by the presence of
UmuD' (181-183). This model suggests that the different umuD gene products, in combination
with UmuC, are involved in distinct survival pathways after UV damage.
Increasing UmuD'2C protein complex concentration was found to antagonize RecA-mediated
recombination of a UV-damaged gene (184); this effect was also observed in vitro (185). In the
proposed model, high concentrations of the UmuD'2C proteins induce replisome switching from
recombination to SOS mutagenesis. Indeed, the UmuD′2C complex has been shown to bind
directly to the RecA/ssDNA filament and could disrupt the DNA pairing activity of RecA (186,
187). Notably, in the presence of homologous DNA sequences, homologous recombination
repair is more prevalent than TLS in responding to DNA damage (188).
The mutagenic potential of Y family polymerases may be further regulated by preferential
formation of heterodimers between UmuD and UmuD', thereby depleting the cell of
mutagenically active UmuD' homodimers (166). UmuD' is degraded by the ATP-dependent
protease ClpXP while in a heterodimeric complex with UmuD (189, 190). Formation of
UmuDD' heterodimers in preference to mutagenically active UmuD' homodimers therefore
specifically targets UmuD' for proteolysis. UmuD also targets its UmuD homodimer partner for
proteolytic degradation by ClpXP (191). The ATP-dependent serine protease Lon is responsible
for the degradation of both UmuD and UmuC proteins in vivo (192). Targeted proteolysis of the
49
umuD and umuC gene products is one mechanism for returning protein levels to their uninduced
state.
1.6.2 Structural dynamics of UmuD and UmuD′
The umuD gene products interact with multiple replication factors such as polymerases UmuC
(as UmuD′2C, pol V), DinB (pol IV) and components of the pol III holoenzyme. The latter
include the polymerase subunit α, proofreading subunit ε, and the processivity clamp β (161,
170, 172, 175, 193, 194). These interactions are due in part to the relative flexibility of full-
length UmuD and its UmuD′ cleavage product as shown biochemically and by X-ray
crystallography, nuclear magnetic resonance spectroscopy (NMR) and circular dichroism (CD)
(168-170). The cleaved form UmuD′ contains disordered N-terminal arms that expose the C-
terminal globular domain to solvent upon cleavage, while in full-length UmuD, the arms are
more stably bound to the globular domain (168, 169). Therefore, UmuD and UmuD′ make
specific contacts that facilitate a variety of protein-protein interactions (168, 170, 193, 195).
The crystal structure of UmuD′ reveals extended N-terminal arms (residues 25-39) and a
globular C-terminal body (residues 40 to 139) that contains the catalytic dyad Ser60 and Lys97
(169). Although two dimer interfaces (designated as molecular and filament) were observed in
the crystal structure, NMR and cross-linking experiments support the conclusion that the so-
called filament dimer interface is the form present in solution (Figure 1.6) (168, 169, 195-200).
However, there is evidence that the filament structure may be biologically relevant (196). To
date, there is no high resolution structure of the UmuD2 homodimer. Cross-linking studies of a
50
series of single-cysteine derivatives of UmuD are consistent with the UmuD2 homodimer
interface resembling the interface of the UmuD'2 homodimer, involving contacts between the C-
termini of the monomers and intermolecular interactions between Asn41 and Leu44 of α helix 1
(168, 171, 200). NMR experiments also suggest that the UmuDD′ heterodimer most closely
resembles the UmuD2 homodimer (168). Dimerization appears to be important for biological
activity, as UmuD' variants that resulted in decreased UV-induced mutagenesis also have severe
deficiencies in their abilities to form homodimers in vivo (197, 201, 202).
Four models of the UmuD homodimer have been generated from NMR, Electron Paramagnetic
Resonance (EPR), and cross-linking studies, and by homology to LexA (168, 172, 203, 204).
One model shows UmuD with the N-terminal arms in trans with the elbows down, where the N-
terminal arm of one monomer folds down across the C-terminal body of the adjacent monomer
and crosses the catalytic site (Figure 1.6). Each UmuD monomer cleaves the N-terminal arm of
its partner at Cys24-Gly25 (172). A trans, elbows up model positions the arms along the outer
edge of the globular domains. Two cis versions with elbows up and elbows down suggest that
each N-terminal arm could bind over its own globular domain (172). The N-terminal region
(residues 1-14) is likely to be in a random extended conformation. Cross-linking and chemical
modification experiments suggest that the trans, elbows down conformation of the N-terminal
domain is the most prevalent in solution (172, 203). However, given the dynamic nature of
UmuD, all four conformations may be physiologically relevant (170, 172).
Circular dichroism (CD) spectroscopy simulating physiological conditions detected random coil
conformations for both UmuD dimers (170), rather than the -sheet-rich structure determined by
51
X-ray crystallography and NMR spectroscopy (168, 169, 195). At higher salt concentration both
UmuD and UmuD′ dimers have more typical -sheet appearance. Thus, the umuD gene products
belong to the group of intrinsically disordered proteins (IDPs) (205). Like their IDP counterparts,
UmuD dimers are capable of making a remarkable number of specific protein-protein contacts.
It has been shown that UmuD and UmuD′ interact with the , , and ε subunits of DNA
polymerase III (193). UmuD interacts less strongly with the subunit in vitro than cleaved
UmuD' (193). Moreover, uncleaved UmuD interacts more strongly with the clamp than
cleaved UmuD' as observed by affinity chromatography (193). Different interactions of UmuD
and UmuD' with the clamp suggest that these interactions regulate how the umuD gene
products access the replication fork. Overexpression of the subunit or the clamp inhibits
cleavage of UmuD to UmuD' in vivo, further supporting the specific interactions between the
pairs of proteins (193). It has also been found that overexpression of the clamp reduces UV-
induced mutagenesis (206).
1.6.3 UmuD-Beta clamp interactions
To date, interactions between the umuD gene products and the β clamp have been studied in
much more detail than other interactions involving the umuD gene products. Proteins that
interact with the β clamp, with the exception of UmuD and UmuD′, contain the eubacterial
clamp-binding motif (QL[S/D]LF) (207). UmuD contains a 14
TFPLF18
sequence within its N-
terminal arm (207). Although the motif lies in a region of UmuD that is important for its
interaction with the β clamp (194), the interaction does not depend on the sequence identity of
52
the motif (172). That is, a UmuD variant containing mutations in this motif binds to the β clamp
with similar affinity as that of wild-type UmuD (see below) (172).
UmuD and UmuD′ affinity chromatography and in vitro cross-linking studies confirm that the β
clamp has a higher affinity for UmuD than UmuD′ (194). However, it has been shown that both
the N-terminal arms and C-terminal globular domains of UmuD are important for interaction
with the β clamp (194). UmuD lacking its N-terminal nine residues is proficient for interactions
with the β clamp, while UmuD lacking the N-terminal 19 residues results in reduced
formaldehyde cross-linking to β (194). The UmuD variant UmuD-3A (T14A L17A F18A), a
non-cleavable variant with mutations of the most conserved residues of the TFPLF motif,
possesses some of the biological functions of the cleaved form UmuD′. Although the Kd values
for interaction of the β clamp with UmuD (5.5 +/- 0.8 μM) and UmuD-3A (6.1 +/- 0.5 μM) are
similar, their interactions with the β clamp may be different, as observed by intrinsic tryptophan
fluorescence of the β clamp (172). Tryptophan fluorescence is a relatively sensitive probe of the
microenvironment. The single tryptophan of the β clamp is located on a flexible loop between
domains I and II of β, and thus is a sensitive reporter of conformational changes (61, 172). The β
clamp tryptophan fluorescence differed dramatically upon UmuD vs. UmuD-3A binding,
suggesting they have different binding modes (172).
UmuD, UmuD′, the α catalytic subunit, UmuC, DinB and clamp loader all interact with the β
clamp around the β clamp hydrophobic pocket, approximately defined by residues Leu177,
Pro242, Val247, Val360, Met362 (Figure 1.4) (52, 90, 93, 105, 106). UmuD, UmuD′ and the α
subunit interact with overlapping regions of β, suggesting that there may be competition for
53
binding (Figure 1.4) (193). This implies that UmuD plays a regulatory role following the SOS
response, interacting with components of Pol III, interfering with α binding to β, slowing
replication and allowing time for error-free repair mechanisms to act (165, 193). It is also
possible that α and UmuD or UmuD′ bind the homodimeric β clamp simultaneously. A model
has been proposed in which cleavage of UmuD to form UmuD′ reduces binding to the β clamp,
thereby releasing the DNA damage checkpoint and enabling translesion synthesis (208).
1.7 DNA Polymerase Switching
Replicative DNA polymerases are unable to copy damaged DNA under most circumstances
(209, 210). In the polymerase-switching model (111, 112, 211, 212), a lesion blocks the progress
of the replicative DNA polymerase at the replication fork, and translesion polymerases act to
allow replication to proceed (Figure 1.7). Since the SOS inducible polymerases are characterized
by low processivity and low-fidelity replication, specialized polymerases must subsequently be
replaced by replicative polymerases to restore efficient and accurate DNA replication.
54
Figure 1.7 Polymerase switching in response to DNA damage.
Replicative DNA polymerases (green) are generally unable to copy damaged DNA. A
polymerase switch occurs allowing a TLS polymerase (orange) access to DNA. The TLS
polymerase synthesizes DNA opposite the lesion and far enough beyond it that the replicative
polymerase can resume synthesis without disruption due to the lesion. In eukaryotes, there is
evidence that two different TLS polymerases act in a stepwise fashion to insert nucleotides
opposite the lesion and then extend the primer beyond the lesion (213).
1.7.1 Replisome Dynamics
Elevated levels of UmuD and UmuC inhibit DNA replication, a phenomenon suggested to
represent a primitive DNA damage checkpoint (165, 180). DinB (pol IV) also interferes with
replication fork progression both in vitro and in vivo, and its overexpression inhibits cell growth
(212, 214). Neither DinB catalytic activity nor its canonical β-clamp binding motif are required
55
for inhibition of replication (214). Both pol II and DinB inhibit replication fork progression by
altering the speed of the DnaB helicase (215). In this case, the β-clamp binding motif is required
for DinB to occupy the replisome and slow the helicase (215). This decrease in helicase velocity
allows time for accurate DNA repair processes to take place while maintaining the overall
structure of the replication fork (215). Taken together, these observations suggest a high degree
of plasticity in the replisome as well as parallels between the physiological effects of pol IV and
pol V on the replication fork.
The clamp binds to, and increases the processivity of, all five known E. coli DNA polymerases
(1, 108, 216) and is important for the lesion-bypass activity of UmuD′2C (90, 91, 93). Both
UmuD and UmuD′ interact with the β clamp and the α and ε subunits of pol III (193). In general,
the clamp-binding proteins possess a canonical peptide motif that binds to the hydrophobic
channel on the clamp (109, 207), suggesting that they compete for the same binding site.
Extensive analysis of DNA polymerase binding to site-directed mutants of the clamp show that
the different polymerases have partly, but not completely, overlapping sites of interaction on the
clamp (217-219). Based on the co-crystal structure of the little finger domain of DinB with the
clamp, a physical basis for recruitment of Y family polymerases to the replisome was proposed
(106). Two interfaces between the DinB little finger domain and the clamp were identified.
One is a peptide-protein interaction involving the extended C-terminal tail of the DinB little
finger domain and hydrophobic channel, or “cleft”, on the surface of the clamp (106). A
second interaction was observed between surface loops of the DinB little finger (residues 303-
305) and the outer rim of the clamp at the dimer interface (residue 98) (106). Modeling of the
entire S. solfataricus DinB-ortholog Dpo4 protein onto the structure of the DinB little finger
56
domain showed that, when bound in this position, DinB would likely not have access to a linear
DNA duplex (106). Therefore, DinB could have two modes of binding to the clamp: a ‘locked-
down’ conformation with no access to DNA, and a ‘tethered’ complex attached to the clamp
via the C-terminal canonical clamp binding motif, which would allow interactions with DNA
(106). By analogy, mutations that disrupt the proposed clamp outer rim interaction of UmuC
result in increased UV-induced mutagenesis, presumably because the ‘lock’ is released, giving
UmuC more access to DNA (90). Recent crystal structures of Dpo4 show that it also adopts
multiple conformations whether in the apo form, bound to DNA or bound to the sliding clamp
(220). Specifically, the little finger rotates around the linker region and adopts different
conformations depending on the presence of different binding partners of Dpo4. While the Dpo4
little finger domain makes additional contacts with the clamp beyond the canonical interaction
motif (PIP-box), unlike DinB the Dpo4 little finger domain lacks interactions with the outer rim
of the clamp.
1.7.2 Polymerase Switching: Tool-Belt and Active Exchange Models
The multiple conformations and flexibility of polymerase binding to the clamp support the
‘tool-belt’ model of polymerase switching (90, 93, 106, 112, 211), in which several polymerases
simultaneously attach to the clamp. Depending on the DNA substrate, either a replicative or a
translesion polymerase is given access to the DNA. The proposed triple polymerase replisome
provides another possible mechanism for efficient switching between high- and low-fidelity
polymerases (10). The observed tilt angle of the clamp on DNA may also facilitate polymerase
switching (104). It was recently reported that only one of the two possible hydrophobic channels
57
that are sites of polymerase interaction on the β clamp is required for switching between pol III
and DinB (221). This result challenges the “tool-belt” model (221). Other studies suggest an
active exchange model in which one polymerase readily displaces another at the replication fork
(114).
DNA replication assays reveal the exchange of the replicative pol III and TLS pol V on DNA in
the presence of DNA damage (111, 222). The pol III holoenzyme replicates up to the nucleotide
position preceding the damaged base, but not beyond it. Through contacts between their minor
groove recognition domain and the DNA, replicative DNA polymerases sense distortions in the
DNA within the last four to five nucleotides replicated (223-225). In agreement with these
structural studies of other replicative polymerases, it has been experimentally shown that pol III
requires the primer to be four to five nucleotides beyond the lesion for resumption of efficient
replication (111). If the primer is not extended sufficiently beyond the lesion and pol III has
access to the DNA, the proofreading subunit ε will degrade the primer (111). In the presence of
DNA damage, the SOS response is induced and RecA coats ssDNA. Pol V binds to RecA and
the clamp and inserts nucleotides opposite the lesion. The less constrained active site of pol V
permits insertion and extension of the 3' terminus of the primer beyond the lesion. Pol V
synthesizes a ‘TLS patch’ ranging from 1 to 60 nucleotides, averaging 20 nucleotides in a single
binding event (111, 222). Once pol V synthesizes a TLS patch of at least four nucleotides, the
pol III holoenzyme is able to efficiently resume DNA replication even in the presence of a
RecA/ssDNA nucleoprotein filament (111).
58
There is growing genetic evidence that multiple polymerases act during replication and in lesion
bypass (226-230). It was hypothesized that SOS inducible polymerases II, IV, and V alter the
fidelity of DNA replication by interfering with the cellular functions of pol I and III. This was
tested by exploiting the observation that each polymerase leaves a unique mutagenic ‘fingerprint’
when copying DNA (226). The spectrum of mutations in the rpoB gene arising in various ΔmutL
strains was measured (226). Deletion of mutL provides a genetic background deficient in
postreplicative mismatch repair to allow determination of actual polymerase-specific
misincorporation events, rather than simply assaying mutations that escape repair. Furthermore,
polymerase expression levels were modulated using strains with mutations in recA and lexA
(226). Analysis of the rpoB genes in strains with deletions of different combinations of
polymerases revealed changes in the spectrum of mutations. This result was interpreted as
evidence that E. coli DNA polymerases compete for access to genomic DNA, thereby promoting
or suppressing mutagenesis (226). Specifically, multiple spontaneous transversion mutations are
due to pol V or to the combined action of pol IV and pol V (226). Moreover, the chromosomally
encoded levels of pol V are the limiting factor for the production of the mutations and even
modest low-level expression causes an increase in transversion at specific hot spots. Facile
switching between the five DNA polymerases of E. coli during genome replication suggests that
spontaneous mutations arise due to the interactions of multiple polymerases (226).
Additional evidence for switching between pol III and pol V on UV-damaged DNA came from a
study revealing that strains harboring variants of E. coli pol V with mutations of F10 or Y11 (the
steric gate residue) display hypersensitivity to UV light and dominant negative genetics (230).
The steric gate residue has been shown to prevent incorporation of ribonucleotides by sterically
59
occluding the 2'-OH moiety in the Y family polymerases (231, 232). The dominant negative
character of umuDC Y11A is suppressed by disruption of dnaQ, the gene encoding the ε
proofreading subunit (Table 1.1). It seems likely that pol V Y11A is recruited to damaged DNA
and prevents other DNA polymerases from accessing the replication fork. The steric gate
variants of pol V (Y11A) and pol IV (F13V) conferred on a wild-type E. coli strain extreme
sensitivity to both nitrofurazone and UV radiation (230, 233). Considering that pol IV copies N2-
furfuryl-dG lesions efficiently and accurately (233), this finding suggest that the pol V steric gate
variant prevents another Y family polymerase from accessing damaged DNA.
1.7.3 Gap-filling Model
Several studies have suggested that lesion bypass does not occur at the replication fork, but at
gaps that are left by the replication machinery (141). This model is supported by the observation
that the rate of replication recovery after DNA damage is independent of the presence of pol IV
or pol V (234, 235). Moreover, as discussed above, lesions in the lagging strand do not slow the
overall rate of replication, but lead to re-initiation downstream of the lesion, leaving a gap (138-
140). Finally, it has been observed that β clamps can be left behind when the polymerase
dissociates from DNA (102), possibly as an aid to recruitment of other replication factors,
including Y family polymerases. The observation that Rev1, a Y family polymerase in yeast, is
upregulated in the G2-M transition of the cell cycle, rather than in S phase when most DNA
replication occurs, is also consistent with this model (236). Ultimately, the two models of
coordinated polymerase switching and gap filling are not mutually exclusive. The specific
60
mechanism a particular cell utilizes likely depends on the type of lesion encountered and the
conditions in the cell at that time.
In this work, we investigate the interactions between the α subunit of DNA pol III, the single-
stranded DNA binding protein SSB, and UmuD. Overall, we show that a network of protein-
protein and protein-DNA interactions modulate DNA replication under normal cellular
conditions and in response to DNA replication stress.
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CHAPTER 2: Selective disruption of the DNA polymerase III α-β
complex by the umuD gene products
This chapter was originally published by Oxford University Press in Nucleic Acids Research as:
"Selective disruption of the DNA polymerase III α-β complex by the umuD gene products"
Michelle C. Silva, Philip Nevin, Erin A. Ronayne, and Penny J. Beuning, Nucleic Acids
Research 40 5511-5522 (2012).
It is reproduced with permission from Oxford University Press:
http://www.oxfordjournals.org/access_purchase/publication_rights.html
Rights retained by ALL Oxford Journal Authors The right, after publication by Oxford Journals, to use all or part of the Article and abstract, for their own
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The right, after publication by Oxford Journals, to use all or part of the Article and abstract, in the
preparation of derivative works, extension of the article into book-length or in other works, provided that a full acknowledgement is made to the original publication in the journal;
The right to include the article in full or in part in a thesis or dissertation, provided that this not published commercially;
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Oxford University Press in advance. Please go ahead with the use ensuring that a full acknowledgment is made to the original source of the material including the journal name, volume, issue, page numbers, year of publication, title of article and to Oxford University Press and/or the learned society.
Michelle C. Silva designed bulk biochemical experiments, purified proteins, performed
experiments, analyzed the data, and wrote the manuscript; Philip Nevin performed FRET studies;
77
Erin A. Ronayne purified proteins and performed bulk experiments; Penny J. Beuning designed
experiments and wrote the manuscript.
2.1 INTRODUCTION
DNA polymerase III (DNA pol III) is the main DNA polymerase in Escherichia coli and is
responsible for replicating the entire genome. It contains ten subunits that allow for accurate and
processive replication to occur (1-4). These ten subunits are organized into three subassemblies:
the polymerase core; the β processivity clamp; and the clamp loader complex (5). The
polymerase core consists of the α polymerase subunit, the ε proofreading subunit and the θ
subunit whereas the clamp loader complex loads the processivity clamp onto primer:template
DNA (6) and coordinates continuous replication on both leading and lagging strands of DNA (7-
9). Once the β clamp is positioned, the clamp loader complex regulates the loading of the
polymerase core onto the β clamp (9). This ensemble of proteins allows for continuous
replication to occur on the leading strand and discontinuous replication to occur on the lagging
strand (10), allowing for efficient replication of DNA.
A crystal structure has been solved of a truncated form of the E. coli α subunit (residues 1-917)
(Figure 2.1A) (11). With a resolution of 2.3 Å, it is possible to distinguish the characteristic
domains of DNA polymerases including the palm domain containing the conserved aspartic acid
residues of the active site, and the fingers and thumb domains (12, 13). The N-terminal domain,
78
which was thought to possess pyrophosphatase activity (14), is also present in the crystal
structure. This domain contains the binding site of the ε proofreading subunit (15). The C-
terminal domain, not included in the crystal structure, contains the
oligonucleotide/oligosaccharide binding (OB) fold (11, 16) responsible for binding single-
stranded DNA (17), the binding site for the τ subunit of the clamp loader complex (9, 18-20) and
the binding site for the β processivity clamp (Figure 2.1A) (21, 22). It was originally suggested
through the use of truncations of α that the C-terminus is responsible for binding to the β clamp
(22); however, site-directed mutagenesis experiments revealed that residues 920-924 are
primarily responsible for binding to the β clamp and that the C-terminus plays a minor role (21).
Because of these findings, we refer to residues 920-924 as the β binding site on α.
Figure 2.1 Models of α and UmuD.
(A) The x-ray crystal structure of α 1-917 (PDB code 2hnh (11)) (left) shows the major domains
of the polymerase, emphasizing the active site residues and the binding site for ε (15). A
homology model of full-length α (23) (right) depicts the C-terminal domain that is not present in
the crystal structure containing the β (21) and τ (9) binding sites as well as the OB domain (11,
17). (B) The homology model of UmuD (24) and the x-ray crystal structure of UmuD′ (25)
79
showing one monomer in green and the other in purple. The first 24 N-terminal residues missing
from the UmuD′ structure (yellow) are cleaved as part of a primitive DNA damage checkpoint
by the active site (orange). The single cysteine residues at C24 is shown (red) in full-length
UmuD.
The α subunit, together with the other subunits of DNA pol III, is capable of efficiently
replicating undamaged DNA. But when it encounters DNA damage, replication by the
polymerase is disrupted (26). DNA pol III is capable of inserting nucleotides opposite some
DNA lesions, but usually cannot extend beyond the unusual primer terminus that it generates.
Thus, it can become trapped in a futile cycle of insertion and exonucleolytic proofreading at sites
of DNA damage (27). On the leading strand, the blockage of DNA synthesis is disruptive,
causing the inhibition of replication fork progression (28, 29). On the lagging strand however,
progression of the replication fork is less affected because the polymerase can reassemble on the
next primer terminus downstream (28). In order to withstand DNA damage, the SOS response is
employed by the cell (30).
The initial molecular trigger for the SOS response is an abundance of ssDNA caused by the
stalled DNA polymerase (31). RecA coats the ssDNA forming a nucleoprotein filament that
facilitates the cleavage of the LexA repressor and up-regulates the expression of at least 57 SOS
genes (32). The products of these genes are involved in DNA damage tolerance mechanisms,
such as recombination, DNA repair and translesion synthesis (TLS), which is the process
involved in replicating damaged DNA. TLS is carried out by potentially mutagenic Y family
DNA polymerases that can replace a stalled replicative DNA polymerase (33-37).
80
Y family DNA polymerases have the specialized ability to bypass DNA damage by inserting
nucleotides opposite the lesion (33-40). Of the five DNA polymerases present in E. coli, DNA
pol IV (DinB) and DNA pol V (UmuD′2C) are members of this family (41-43). DNA pol V is
composed of the UmuD′2 dimer, which is the RecA/ssDNA-facilitated cleavage product of
UmuD2, and UmuC, which possesses polymerase activity (44, 45). During the first 20-40 min
after the activation of the SOS response, UmuD2 is the predominant species (46, 47). UmuD2
together with UmuC serves to affect a primitive DNA damage checkpoint, inhibiting replication
to allow time for accurate DNA repair processes to act (46, 48, 49). After approximately 40 min,
UmuD′2 becomes the predominant species, releasing the primitive DNA damage checkpoint (46,
50, 51). UmuD′2 and UmuC form DNA pol V (44, 45, 52), which is the form active in TLS.
UmuD2 is a very tight dimer (Kd <10 pM) (53) and is expected to be dimeric in all experiments
reported here; therefore we use UmuD to refer to homodimeric UmuD and variants. Each UmuD
monomer consists of two domains: the N-terminal arms and a C-terminal globular domain. One
model of UmuD places the N-terminal arms folded down onto the globular domain, positioning
the cleavage site (between C24 and G25) at the active site (S60 and K97) (an “arms down”
conformation) (Figure 2.1B) (24, 54). But because of the dynamic nature of arms, UmuD can
exist in a conformation where the arms are unbound (an “arms up” conformation) exposing the
globular domain (24, 53, 55).
The umuD gene products have also been shown to interact with other proteins such as DNA pol
IV (Kd 0.62 µM) (56), and components of DNA pol III including the α polymerase, the ε
proofreading subunit and the β processivity clamp (57). UmuD and UmuD′ differentially interact
with components of DNA pol III (57). Specifically, affinity chromatography experiments
81
suggested that UmuD binds more strongly to the β processivity clamp than to α (57), supporting
the role of UmuD in inhibiting DNA synthesis as part of a primitive DNA damage checkpoint
(50), however the α-UmuD interaction has not been characterized at the molecular level. It has
also been suggested that upon binding to the β clamp or to DinB, the structure of UmuD
becomes less disordered (53). These interactions suggest that both UmuD and UmuD′ play a role
in replication fork management.
The process by which exchange of the α polymerase subunit and a TLS polymerase occurs is not
fully understood. There are likely a number of factors that facilitate such exchange; for example,
it has been shown that the presence of DinB inhibits DNA pol III replication (58). In this work,
we characterized the interactions of α with the umuD gene products. We determined the binding
affinity between UmuD and α. We also determined that there are two UmuD binding sites on α,
one of which favors UmuD over UmuD′. The UmuD-specific site overlaps with the β binding
site on α and consequently, we found that UmuD, but not UmuD′, specifically disrupts the α-β
interaction. Therefore, we find that UmuD plays a key role in regulating polymerase access to
the β clamp.
2.2 MATERIALS AND METHODS
2.2.1 Proteins and Plasmids
82
UmuD and UmuD′ were expressed from pSG5 and pSG4 plasmids, respectively, as previously
described (59). UmuD3A (UmuD containing the triple mutation T14A, L17A, and F18A),
UmuD-S60A, and UmuD′-S60A variants were previously described (24). Plasmids that express
His-tagged wild-type α and the truncations α1-280, α1-917, α1-956 and α1-975 were provided by
Dr. Meindert Lamers and Prof. John Kuriyan, UC Berkeley (11). The plasmids encoding the
truncations α1-835 and α917-1160 were cloned as described previously (17). Wild-type α and
truncations were expressed in Tuner(DE3) (Novagen) competent cells and purified using the
established protocol and stored at -20 °C in a buffer containing 50 mM Hepes (pH 7.5), 100 mM
NaCl, and 50% glycerol (protein storage buffer) (11). The β processivity clamp was purified
according to published protocols (60). LexA was purified from pJWL228 (provided by Dr.
Ronaldo Mohana-Borges, Instituto de Biofísica Carlos Chagas Filho-UFRJ, Brazil) as described
(61).
To improve the stability of the α protein, various conditions were tested including different
buffer conditions and varying temperatures. The most stable proteins were those stored at -20 °C
in a buffer containing 50% glycerol, at 25 µM or lower concentrations. The high concentration of
glycerol was necessary to keep the proteins from freezing. It was noted that the stability of α was
reduced when the protein was frozen and thawed before use. In solutions with lower glycerol
concentrations, aggregation was observed. We found that the high concentration of glycerol
interfered with many of the experiments described here; therefore, before each experiment, the
buffer was exchanged to the relevant buffer for each assay. This was accomplished by using
Zeba Desalt Spin Columns (Thermo Scientific) with the buffer exchange protocol provided.
83
2.2.2 Tryptophan Fluorescence Assay
The intrinsic tryptophan fluorescence of α in the presence of increasing amounts of UmuD was
used to determine the equilibrium constant for α binding to UmuD. UmuD has no tryptophan
residues and α has eight tryptophans. To determine the fluorescence of α alone, 60 µL of 5 µM α
in 50 mM Hepes (pH 7.5), 100 mM NaCl was excited at 278 nm, and emission was monitored
from 300 to 500 nm. To determine the binding constant between α and UmuD, the sample of α
was then titrated with UmuD. After each addition, the sample was excited at 278 nm and
emission was monitored from 300 to 500 nm, with both excitation and emission slits set to 5 nm.
To analyze the change in fluorescence of α, the maximum of each peak was determined and
corrected for the dilution caused by adding UmuD. To correct for any fluorescence contribution
from UmuD alone, fluorescence emission was monitored from UmuD in reaction buffer at the
same concentrations used to titrate α. To produce a binding curve, the fraction of fluorescence
from α quenched by the addition of UmuD (Q) was plotted versus the ratio of UmuD to α. The
curve was fit to the following equation using GraphPad Prism (La Jolla, CA):
[
( [ ]
[ ]
) √ ( [ ]
[ ]
)
[ ]
[ ]
]
(Equation 2.1)
84
where P is the initial concentration of α and T is the concentration of the UmuD stock used in the
titration. The binding constant at pH 10 for α and UmuD was obtained in alkaline cleavage
buffer (see below).
2.2.3 Thermal-Shift Assays
Reactions were assembled in 96-well PCR plates (Applied Biosystems) in which each sample
(16 μL total volume) consisted of thermal-shift assay buffer (50 mM Hepes (pH 7.5), 100 mM
NaCl), and 25x Sypro Orange (Invitrogen) excited at 490 nm. In experiments in which the
melting transitions of UmuD were observed in the presence of α, each sample contained 45 µM
UmuD (monomer concentration) and 1 μM α. Samples were incubated for 2 h at room
temperature before detection. In experiments in which the stability of the truncations of α was
compared with that of wild-type α, each construct of α was added to a final concentration of 5
µM without incubation before detection. In order to increase the thermal stability of the α
proteins for ease of detection, 30% glycerol was added to each sample. Once the plate was sealed
with optical adhesive film (Applied Biosystems), an iCycler iQ5 Real-Time PCR (Bio-Rad) was
used to increase the temperature from 25 °C to 75 °C with an increment of 0.1 °C and a 10 sec
dwell time per temperature increment. The fluorescence intensities emitted at 575 nm and
detected by the built-in CCD camera were plotted versus temperature. The melting temperature
(Tm) was calculated as described (62, 63). The assays were repeated several times with similar
observations and consistent melting temperatures.
2.2.4 UmuD in vitro Cleavage Assays
85
RecA/ssDNA nucleoprotein filament-facilitated UmuD cleavage reactions were carried out as
described in LG Buffer (59). Samples containing α, UmuD and buffer were incubated for 2 h at
room temperature before the addition of RecA/ssDNA, after which reactions were carried out at
either 37 °C or 30 °C for 45 min. The reaction temperature was adjusted to 30 °C wherever
possible because of the relatively low melting temperature of α (approximately 38 °C). To rule
out any non-specific interactions, BSA was used instead of α, or, alternatively, LexA cleavage
was assayed in the presence of α. Alkaline cleavage of UmuD was also carried out as previously
described (59). The alkaline cleavage buffer (100 mM Glycine (pH 10), 10 mM CaCl2, 50 mM
NaCl, 10 mM dithiothreitol (DTT), 0.25 mg/mL BSA) was added after a 30 min incubation at
room temperature, and the reactions were carried out at either 37 °C or 30 °C for 48 hours.
Reactions of cleavage assays were analyzed by using 18% SDS-PAGE.
2.2.5 Cross-Linking of UmuD using Bis-maleimidohexane (BMH)
Bis-maleimidohexane (BMH, Pierce) cross-linking (24, 64) was carried out as described by
incubating 10 µM UmuD-S60A in the presence of 10 µM wild-type α, α917-1160, or α1-280 in
10 mM Sodium Phosphate (pH 6.8), 100 mM NaCl and 1 mM BMH. The reactions were
incubated for 5 min at room temperature, after which the reactions were quenched with 50 mM
DTT. Cross-linked dimers of UmuD-S60A were resolved from monomers by 4-20% SDS-PAGE
(Bio Rad). An immunoblot was used to identify those bands containing UmuD-S60A using
rabbit anti-UmuD as the primary antibody as described (24, 59).
2.2.6 Fluorescence Resonance Energy Transfer
86
Protein labeling was carried out using Alexa Fluor 488 C5-maleimide (Invitrogen) for the β
clamp and Alexa Fluor 647 C2-maleimide (Invitrogen) for the α subunit, in a reaction buffer
containing 50 mM Hepes (pH 7.5) and 200 mM NaCl. The β clamp (100-150 μM) and α (20-50
μM) were each labeled by adding 5-10 molar excess of the respective Alexa Fluor maleimide at
room temperature, followed by overnight incubation at 4 °C in the dark. Unreacted reagent was
separated from labeled protein on a Sephadex G-50 (GE Healthcare) column at 4 °C using a
buffer containing 25 mM Hepes (pH 7.5) and 200 mM NaCl. Fractions were analyzed by SDS-
PAGE and the gels were analyzed on a Storm 860 phosphorimager (Molecular Dynamics) by
monitoring fluorescence after irradiating the gels at 450 nm. The degree of labeling was
determined as described by the manufacturer and was on average 3-4 fluorophores per α subunit
and 1-2 fluorophores per β monomer.
The interaction between the α subunit and the β clamp was monitored by FRET. A reaction
mixture containing 350 nM donor-labeled β subunit (βA488
) and 1 μM acceptor-labeled α subunit
(αA647
) in 70 μL FRET buffer (25 mM Hepes (pH 7.5), 25 mM NaCl, 1 mM DTT, and 0.5 mM
EDTA) was incubated for 15 min on ice, followed by analysis at room temperature with a Cary
Eclipse spectrofluorimeter (Varian). Samples were excited at 495 nm and emission spectra were
recorded from 500 to 700 nm, with slits for both excitation and emission set to 5 nm. The effects
of UmuD, UmuD′ and UmuD-S60A were investigated by adding up to 40 μM UmuD or variants
to each sample prior to analysis. FRET efficiency (E) was calculated by finding the ratio between
the emission intensity of the donor in the presence of the acceptor (Fd′) to the emission intensity
of the donor alone (Fd), which is then subtracted from one (Equation 2.2).
87
′
(Equation 2.2)
2.3 RESULTS
2.3.1 α binds UmuD via two UmuD Binding Sites on α
In order to develop a more complete picture of the interactions of UmuD with subunits of the
replisome, we probed the interaction between α and the umuD gene products. Because α has
eight tryptophans and UmuD has none, we were able to use the intrinsic tryptophan fluorescence
of α to determine the equilibrium binding constant for the interaction between α and UmuD. By
analyzing the extent of α fluorescence quenching observed with the addition of UmuD, the Kd
was determined to be 1.1 ± 0.6 µM (Figure 2.2). This binding constant is similar to that for other
protein interactions with UmuD (UmuD and the β clamp: 5.5 ± 0.8 μM (24); UmuD and DinB:
0.62 μM (56)). To localize the binding site of UmuD on α, a number of truncations of α were
constructed and dissociation constants for their interactions with UmuD were determined (Figure
2.3A). The truncation α1-975 has a binding constant (3.1 ± 1.0 µM) similar to that of full-length
α indicating that the binding site is located within this region. When the truncation α1-956 was
analyzed, a significant decrease in affinity was observed (Kd = 14.7 ± 1.9 µM), suggesting that
88
there is a UmuD binding site between residues 956-975 positioned in the C-terminal domain
(Figure 2.1A). This region is also near the β-binding site (residues 920-924) (21).
Figure 2.2 UmuD binds wild-type α with a Kd = 1.1 μM.
The curve representing the fraction of α tryptophan fluorescence quenched by various
concentrations of UmuD was fit to Equation 2.1, which produced a Kd value of 1.1 ± 0.6 µM at
pH 7.5. The error bars represent the standard deviation from five independent experiments.
89
Figure 2.3 UmuD binds α at two distinct binding sites.
(A) Kd values for UmuD binding to various α truncations. The large difference between the Kd
values for the truncations α1-975 and α1-956 indicates that one binding site is between residues
956-975. The reduced but still significant binding constants for the truncations α1-956, α1-917,
α1-835, and α1-280 suggests another UmuD binding site is present in the N-terminal domain α1-
280. The domains of α are labeled above the schematic of the protein; ε, β, and τ indicate the
location of the respective binding site for each protein, HhH indicates the helix-hairpin-helix
domain involved in double-stranded DNA binding, OB indicates the OB-fold domain involved in
single-stranded DNA binding. (B) Melting curves for the various α truncations used show
comparable stability to that of full-length α.
90
The Kd values for truncations α1-917, α1-835, α1-280 are all similar to that of truncation α1-956.
That binding was still detected with these truncations suggests that there is a second UmuD
binding site within α. N-terminal truncations smaller than 280 residues were expressed poorly
and so could not be analyzed. As a result, this second binding site could not be localized any
further than to residues 1-280, the PHP domain (Figure 2.1A). The Kd value for UmuD binding
to the truncation α917-1160, which contains the proposed C-terminal UmuD binding site (α
residues 956-975), was determined to be 13.9 ± 5.1 µM which is somewhat similar to that for
truncation α1-280 (7.3 µM) but greater than that of full-length α (1.1 µM). The increased affinity
for UmuD seen with full-length α as opposed to that of the truncations containing only one
binding site may be due to the simultaneous binding of two UmuD dimers to α, as the two UmuD
binding regions are spatially somewhat distant (Figure 2.1A).
A thermal-shift assay was used to determine if any of the truncations assayed here were unstable.
This assay involves the use of an environmentally-sensitive fluorescent dye where the
fluorescence intensity is proportional to the hydrophobicity of its environment. When a folded
protein is present, the dye is in an aqueous environment and so the fluorescence of the dye is
quenched. As the temperature reaches the melting point of the protein, the dye binds to the
exposed hydrophobic interior of the unfolding protein, increasing the fluorescence intensity of
the dye. The temperature at the midpoint of the transition between the folded and unfolded states
is known as the melting temperature, or Tm. Because the Tm is related to the stability of the
protein, a protein with a higher Tm is more stable than a protein with a lower Tm. All α
truncations were found to be about as stable as full-length α (Figure 2.3B); while the truncations
α1-280 and α917-1160 were even more stable than full-length α.
91
2.3.2 α Inhibits RecA/ssDNA-Facilitated Cleavage in vitro
It was previously shown that overexpression of α inhibited UmuD cleavage in vivo (57). We
wanted to determine whether this phenomenon could be observed in vitro and therefore could be
used to further characterize the UmuD binding site on α. Assays of RecA/ssDNA-facilitated
UmuD cleavage were conducted in the presence of 10 to 25 µM α, which resulted in
approximately 50% less cleavage of UmuD at 37 °C than in assays without α (Figure 2.4A).
Because the melting point of α is approximately 38 °C at pH 7.5 (Figure 2.4B), the assay was
also conducted at 30 °C to minimize denaturation of α present in the reaction. This resulted in
even less UmuD cleavage, approximately 60% less cleavage than with UmuD alone. In order to
determine whether the inhibition observed was specific for both UmuD and α, two controls were
employed: (1) UmuD cleavage in the presence of BSA; and (2) LexA cleavage in the presence of
α, as LexA also undergoes RecA/ssDNA-facilitated self-cleavage (61). In both cases, there was
minimal change in cleavage efficiency (Figure 2.4D), indicating that the inhibition of UmuD
cleavage by α is specific.
92
Figure 2.4 The presence of α inhibits UmuD cleavage.
After each cleavage reaction, UmuD and UmuD′ were resolved by 18% SDS-PAGE.
Representative data are shown in A and C. (A) RecA/ssDNA nucleoprotein filament-facilitated
UmuD cleavage assays were performed in the presence of 10 and 25 µM α wild type and at 37
°C and 30 °C. (B) To determine stability, the melting temperature was obtained for full-length α
at pH 7.5 (25 µM α, 50 mM Hepes (pH 7.5)) and at pH 10 (25 µM α in alkaline cleavage buffer).
(C) Cleavage assays were conducted under alkaline conditions in the absence of RecA/ssDNA.
(D) A comparison of the cleavage assay performed under different conditions: RecA/ssDNA
facilitated UmuD cleavage in the presence of α (red); UmuD cleavage under alkaline conditions
in the presence of α (purple); RecA/ssDNA facilitated UmuD cleavage in the presence of BSA
(blue); RecA/ssDNA facilitated LexA cleavage in the presence of α (green). Percentage of
cleavage was calculated by comparing the density of cleaved product to the total amount of
protein present. Each point represents the average of at least three experiments and the error bars
93
represent the standard deviation from three independent experiments. (E) The Kd at pH 10 was
determined to be 21.7 ± 11.7 µM.
Inhibition of UmuD cleavage in the RecA/ssDNA-facilitated cleavage assay may be due to the
competitive binding of RecA/ssDNA and α for UmuD. When bound to α, UmuD cannot bind to
the RecA/ssDNA filament and so cleavage cannot occur under these conditions. It is also
possible that in addition to competitive binding, α may directly affect the ability of UmuD to
cleave itself. In order to assess the possibility that α inhibited UmuD cleavage directly, rather
than via competition with the RecA/ssDNA filament, cleavage assays were conducted under
alkaline conditions (pH 10) (59) without the RecA/ssDNA nucleoprotein filament (Figure 2.4C).
It was previously shown that UmuD can undergo autocleavage, albeit slowly, in the absence of
RecA/ssDNA under alkaline conditions, as the elevated pH apparently facilitates deprotonation
of the S60 nucleophile on UmuD (59, 65). Under these conditions, no change in cleavage
efficiency was observed, except at the highest concentrations of α and at 30 °C. At pH 10, α was
determined to be stable with a melting temperature of approximately 36 °C (Figure 2.4B).
However, we found that at pH 10, the Kd was higher than at pH 7.5, indicating a decrease in
affinity of α for UmuD (Figure 2.4E). Consequently, the observation that α did not substantially
change the cleavage efficiency of UmuD under alkaline conditions may be attributed to the
decreased affinity at pH 10. As a result, we conclude that in the case of RecA/ssDNA-facilitated
cleavage, cleavage is inhibited likely due to competition for binding to UmuD between α and the
RecA/ssDNA filament.
94
To confirm the binding sites between α and UmuD, the RecA/ssDNA-facilitated cleavage assay
was conducted in the presence of two α truncations: the α1-280 truncation containing the N-
terminal binding site (Figure 2.5A) and the α917-1160 truncation containing the C-teriminal
binding site (Figure 2.5B). As the concentration of α1-280 was increased from 0 µM to 40 µM,
inhibition of cleavage was observed (Figure 2.5A). Approximately four times more α1-280 than
wild-type α is required to observe the same extent of cleavage inhibition, consistent with the
decreased affinity of α1-280 for UmuD. In contrast, the presence of 0 to 5 µM α917-1160
modestly increased UmuD cleavage efficiency (Figure 2.5B), which suggests that UmuD binds
the C-terminal binding site in a conformation that may facilitate cleavage.
Figure 2.5 The α truncation 1-280 inhibits UmuD cleavage.
95
RecA/ssDNA nucleoprotein filament-facilitated UmuD cleavage was carried out at 30 °C in the
presence of (A) the α1-280 truncation and (B) the α917-1160 truncation. SDS-PAGE analysis
showed that approximately four times as much α1-280 is needed to produce the same extent of
cleavage inhibition as wild-type α. On the other hand, the presence of α917-1160 increased
cleavage efficiency. Percentage of cleavage, shown below each lane, was calculated by
comparing the density of cleaved product to the total amount of UmuD present.
2.3.3 The Interaction between α and UmuD is Conformation Dependent
A thermal-shift assay was used to determine whether the presence of α had an effect on the
conformational dynamics of UmuD. Previously, it had been shown that UmuD exhibits two
distinct melting transitions: one at approximately 28 °C and another at approximately 60 °C (63).
The absence of the first transition in the case of UmuD′ suggests that the lower-temperature
transition represents the dissociation of the N-terminal arms of UmuD from the globular domain
(55, 63). In order to observe the effect of α on these two melting transitions, the thermal-shift
assay was conducted in the presence of α (Figure 2.6A). By comparing the melting curve of
UmuD alone with the melting curve of UmuD with α, it is clear that upon addition of α, the first
melting transition is diminished. This suggests that α binds UmuD in a conformation-dependent
manner. Two scenarios are possible (Figure 2.6C) when α binds to UmuD: (1) the arms are
confined in such a way that they cannot dissociate from the body (“arms down”) or (2) the arms
are not bound to the body and therefore do not undergo the transition to the unbound
conformation (“arms up”). The second melting transition is essentially unchanged in the
presence of α.
96
Figure 2.6 Binding to α alters the conformation of UmuD.
(A) Melting curves of 45 μM UmuD (solid line), 45 μM UmuD with 1 μM α (dotted line), and 45
μM UmuD′ (dashed line). The solid line shows the melting transitions for UmuD at 32 °C and 60
°C. The disappearance of the first melting transition suggests that the interaction between α and
UmuD influences the conformation of the arms. (B) Percent UmuD-S60A cross-linked by using
BMH in the presence of either full-length α, α1-280 or α917-1160. (C) Cartoon showing two
possible conformations of the UmuD N-terminal arms. The star indicates the position of the C24
residues, which are cross-linked by BMH.
To determine whether the C-terminal binding site binds UmuD in an “arms up” or “arms down”
conformation (Figure 2.6C), UmuD was cross-linked using the homobifunctional thiol-specific
reagent bis-maleimidohexane (BMH) in the presence or absence of α (Figure 2.6B). The UmuD
dimer has two cysteine residues, one on each arm, which are represented by a star in Figure
2.6C. In order to cross-link with BMH, the two cysteine residues need to be in close proximity
(within ~13 Å) to each other. If the arms are bound to the body in the “arms down”
97
conformation, the two Cys residues are too far apart to be cross-linked; the arms will only be
efficiently cross-linked if they are unbound from the body. As a result, this cross-linking assay is
ideal for looking at the conformational dynamics of the arms. In order to prevent contamination
with the UmuD′ cleavage product, the UmuD-S60A non-cleavable variant was used. It was
observed that in the presence of the α truncation α917-1160, the amount of cross-linked UmuD-
S60A dimer compared to the total UmuD-S60A present in the reaction was significantly reduced
compared to when UmuD was cross-linked alone (Figure 2.6B). This suggests that the C-
terminus of α binds UmuD in an “arms down” conformation. The presence of the α truncation
α1-280 showed a similar but less dramatic reduction of cross-linking efficiency (Figure 2.6B).
2.3.4 The C-terminal Binding Site Favors Full-length UmuD
To determine if the interaction between α and UmuD is selective for UmuD or UmuD′, binding
constants were determined for the interaction of UmuD variants with full-length α, and the
truncations α1-280 and α917-1160 (Table 2.1). When wild-type UmuD is purified, there is
typically some amount of contaminating UmuD′. Moreover, cleavage can occur upon incubation
or interaction with other proteins, so it is nearly always a complicating factor with wild-type
UmuD. Furthermore, mixtures of UmuD and UmuD′ form UmuDD′ heterodimers (66);
therefore, UmuD with some UmuD′ present will be a mixture of several dimeric species.
Consequently, to determine the binding constant between full-length α and UmuD
uncontaminated with UmuD′, the non-cleavable UmuD variant UmuD-S60A was used. The
binding constant between UmuD-S60A and full-length α was determined to be 10.6 ± 2.9 µM.
98
With UmuD′, the binding constant was 10.9 ± 1.6 µM, suggesting that full-length α does not
favor UmuD′ or UmuD.
Table 2.1 Equilibrium dissociation constants Kd (µM) for binding of α truncations to UmuD
variants.
UmuD variants Full-length α α1-280 α917-1160
UmuD 1.1 ± 0.6 7.3 ± 1.4 13.9 ± 5.1
UmuD-S60A 10.6 ± 2.9 10.3 ± 4.3 0.7 ± 0.3
UmuD′ 10.9 ± 1.6 8.6 ± 1.0 3.8 ± 0.9
UmuD3A 8.7 ± 1.5 9.3 ± 3.2 3.4 ± 1.0
UmuD′-S60A/UmuD-S60A 4.6 ± 0.9 ---- ----
Binding affinity was also analyzed between the α truncation α917-1160, containing the proposed
C-terminal binding site (residues 956-975), and the UmuD variants UmuD-S60A and UmuD′. As
shown in Table 1, α917-1160 binds significantly more strongly to UmuD-S60A (0.7 ± 0.3 µM)
than to UmuD′ (3.8 ± 0.9 µM), suggesting that the C-terminal binding site selectively favors full-
length UmuD over the cleavage product, UmuD′. On the other hand, the α truncation α1-280,
containing the proposed N-terminal binding site, does not distinguish between full-length UmuD
and UmuD′, exhibiting similar binding constants with UmuD-S60A (10.3 ± 4.3 µM) and UmuD′
(8.6 ± 1.0 µM) (Table 2.1).
UmuD3A is a full-length UmuD variant that exhibits properties of UmuD′ (24). UmuD3A
contains three mutations (T14A, L17A, F18A) located in the N-terminal arms of the protein,
which disrupt packing of the arms against the C-terminal globular domain and cause UmuD3A to
adopt a more UmuD′-like conformation (24, 55, 63). As expected, UmuD3A has similar affinity
for the α variants as UmuD′ (Table 2.1). Because both UmuD′ and UmuD3A have a more
99
exposed globular domain than wild-type UmuD, it seems that the α C-terminus preferentially
binds UmuD when the arms are bound to the C-terminal globular domain of UmuD, which is
consistent with our findings in the cross-linking experiments with BMH (Figure 2.6B).
The distinguishably different binding constants for wild-type UmuD compared to those of
UmuD-S60A and UmuD′ are consistent with our observation that preparations of wild-type
UmuD may contain some UmuD'. To probe this further, we assembled a mixture of 1:1 UmuD-
S60A:UmuD′-S60A (Table 2.1) to somewhat resemble the conditions of wild-type UmuD.
Because it is known that a monomer of UmuD can cleave the arm of its partner (67), we used the
active site mutation S60A in both UmuD and UmuD′ to prevent any further cleavage. Under
these conditions, the Kd was determined to be 4.6 ± 0.9 µM, which is intermediate between the
Kds determined with UmuD compared to with UmuD′ or UmuD-S60A.
2.3.5 UmuD Disrupts the DNA Pol III α-β Complex
Our observation that UmuD binds α at a site (residues 956-975) that is near the β-binding site on
α (residues 920-924) prompted us to investigate the effect of the umuD gene products on the
DNA pol III α-β complex. The β clamp and the α subunit were labeled with donor and acceptor
fluorophores, respectively, as shown in Figure 2.7C, and FRET was monitored in the presence or
absence of UmuD, UmuD′ or UmuD-S60A (Figure 2.7A and Figure 2.7B). As expected, FRET
was observed when the donor on the β clamp was excited in the presence of acceptor-labeled α
subunit (Figure 2.7A), indicating that the two proteins interact with each other. Addition of
UmuD to the reaction resulted in higher emission intensity from the donor, indicating a decrease
100
in FRET efficiency and consistent with disruption of the α-β complex. The effect was more
apparent with the addition of UmuD-S60A, the full-length non-cleavable variant of UmuD. This
suggests that UmuD is able to disrupt the interaction between the α subunit and the β clamp. In
comparison, no difference in FRET efficiency was observed in the presence of UmuD′ (Figure
2.7A and 2.7B), suggesting that UmuD′ is not able to disrupt the α-β complex. These findings are
consistent with our observation that the binding site for UmuD on the C-terminal domain of α,
near where β also binds to α, binds full-length UmuD (UmuD-S60A) with higher affinity than it
binds UmuD′. Taken together, these observations indicate that specifically full-length UmuD is
capable of disrupting the α-β complex (Figure 2.7C).
Figure 2.7 Fluorescence resonance energy transfer analysis shows that UmuD disrupts the
interaction between the α polymerase, labeled with Alexa Fluor 647 C2-maleimide, and the
β processivity clamp, labeled with Alexa Fluor 488 C5-maleimide.
101
(A) FRET was reduced in the presence of UmuD and UmuD-S60A but not in the presence of
UmuD′. FRET between β and α was reduced in the presence of UmuD-S60A in a concentration-
dependent manner, from 1-40 μM UmuD-S60A (not shown). (B) Bar graph showing FRET
efficiency at each condition; representative of five trials. The error bars represent the standard
deviation of at least three trials. (C) Schematic of FRET experiment showing that the FRET
observed upon interaction of β and α is eliminated with the addition of full-length UmuD, but not
UmuD'.
2.4 DISCUSSION
The disruption of replication fork progression caused by DNA damage necessitates TLS
polymerases having access to the primer terminus at the site of the damage in order to carry out
replication of damaged DNA. Two models have been proposed to explain how replicative and
TLS polymerases exchange: the “toolbelt” model and the dynamic processivity model. It has
been shown that the α subunit and pol IV can interact with the β clamp simultaneously (68),
allowing replication to alternate between the two DNA polymerases without the need for them to
dissociate from the β clamp (the “toolbelt” model). In such a model, when a DNA lesion is
encountered, α stalls and a Y family polymerase takes over replication (27). Once the DNA
lesion has been bypassed, α once again resumes replication. On the other hand, the dynamic
processivity model (69) suggests that the replicating polymerase is constantly being exchanged
with other polymerases without affecting overall processivity. DNA pol IV (DinB), a TLS
polymerase, has been shown to inhibit DNA pol III replication, which may facilitate dissociation
of the pol III α subunit and allow TLS to occur (58).
It has been shown that UmuDC inhibits DNA replication (46, 49), allowing time for accurate
DNA repair processes to occur (46). There is also evidence that the umuD gene products play a
102
direct role in regulating access to the replication fork by interacting with the α, β, and ε subunits
of DNA pol III (57). The interaction between UmuD and the β clamp (24) is believed to
contribute to a primitive DNA damage checkpoint (57). In this report, we investigate the
interaction between the α subunit and the umuD gene products, UmuD and UmuD′. Our
observations suggest that another contribution to the DNA damage checkpoint is likely to be the
disruption of the α-β complex by UmuD.
In this work, we identified two UmuD binding sites on α (Figure 2.3A): one in the N-terminal
domain (residues 1-280) and one in the C-terminal domain (residues 956-975). It has been shown
that UmuD and UmuD′ differentially bind the α and β subunits, where UmuD′ interacts more
strongly with α than with β and UmuD interacts more strongly with β than with α (57). A
comparison of the binding constant between UmuD and β (5.5 ± 0.8 µM) (24) with the binding
constant for UmuD-S60A and wild-type α (10.6 ± 2.9 µM) (Table 2.1), supports the previous
suggestion, based on affinity chromatography with cell extracts, that UmuD binds more tightly to
the β clamp. On the other hand, the interaction between α and the umuD gene products is
complicated by our observation of two UmuD binding sites on α and because the C-terminal
binding site favors binding to full-length UmuD. Notably, the Kds we determined for a variety of
UmuD and α constructs (ranging from 0.7-14 μM, Table 2.1) are similar to those obtained
between UmuD and β and between UmuD and DinB (24, 56). The cellular concentration of
UmuD and UmuD′ ranges from 0.25-1 μM depending on whether cells are SOS induced (70),
which suggests that UmuD and α can interact in vivo.
103
The C-terminal domain of α appears to be a hub for protein and DNA interactions. This apparent
regulatory domain contains the binding site for the β clamp (residues 920-924) (21), and a
binding site for τ (residues 1120-1160) (9, 19), a subunit of the clamp loader complex (Figure
1A). We observed that the C-terminal domain of α (residues 917-1160) favors binding to full-
length UmuD (Table 2.1), and that full-length UmuD is able to disrupt the binding of α to β. This
domain is also believed to contain an OB fold, which has been shown to bind preferentially to
ssDNA and acts as a sensor detecting ssDNA upstream from the primer terminus (17, 71). On the
other hand, less is known about the N-terminal domain of α. Apart from binding UmuD, as we
have demonstrated here, this domain has also been shown to bind the ε subunit (15).
Unfortunately, it is difficult to probe possible direct effects of UmuD on the α-ε interaction
because of the difficulty of acquiring purified, soluble ε.
The umuD gene products have distinct and temporally separated roles in the DNA damage
response, first by taking part in a primitive DNA damage checkpoint by inhibiting DNA
replication and then by activating TLS (46). We observed that FRET between α and β was
reduced with the addition of full-length UmuD, suggesting that the interaction between α and β is
disrupted specifically by UmuD (Figure 2.7C). FRET efficiency was not affected by the addition
of UmuD′. This selective disruption of the α-β complex by full-length UmuD together with the
preferential binding of full-length UmuD by the C-terminal domain of α suggests a specific role
for full-length UmuD in polymerase management. Because UmuD is the predominant species at
the beginning of the SOS response, the displacement of α from the β clamp most likely occurs
before the appearance of DNA pol V (UmuD′C).
104
How can α differentiate between UmuD and UmuD′ at the molecular level? The primary
difference between UmuD and UmuD′ is the absence of the N-terminal 24 amino acid residues in
UmuD′ suggesting that the N-terminal arms are involved in the interaction between α and
UmuD. As shown here, several lines of evidence, including thermal melting and cross-linking
analysis, support a model in which α binds UmuD in an “arms down” conformation. Similarly,
the interaction between UmuD and the β clamp has been shown to involve both the N-terminal
arms and the C-terminal globular domain of UmuD (50); furthermore, the β clamp can bind to a
UmuD variant that contains an artificial disulfide bridge (C24-F94C) locking the arms to the
globular domain (50). Taken together, previous and current work suggests that both α and the β
clamp bind UmuD in a similar fashion in which the N-terminal arms are bound to the C-terminal
globular domain. Collectively, our findings suggest that the already extensive role of UmuD in
responses to DNA damage includes releasing α from the β clamp.
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110
Chapter 3: Polymerase manager protein UmuD directly regulates E.
coli DNA polymerase III α binding to ssDNA
This chapter was recently published online by Oxford University Press in Nucleic Acids
Research as:
"Polymerase manager protein UmuD directly regulates E. coli DNA polymerase III α binding to
ssDNA" Kathy R. Chaurasiya, Clarissa Ruslie, Michelle C. Silva, Lukas Voortman, Philip
Nevin, Samer Lone, Penny J. Beuning, and Mark C. Williams, Nucleic Acids Research, doi:
10.1093/nar/gkt648.
It is reproduced with permission from Oxford University Press:
http://www.oxfordjournals.org/access_purchase/publication_rights.html
Rights retained by ALL Oxford Journal Authors The right, after publication by Oxford Journals, to use all or part of the Article and abstract, for their own
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The right, after publication by Oxford Journals, to use all or part of the Article and abstract, in the preparation of derivative works, extension of the article into book-length or in other works, provided that a full acknowledgement is made to the original publication in the journal;
The right to include the article in full or in part in a thesis or dissertation, provided that this not published commercially;
For the uses specified here, please note that there is no need for you to apply for written permission from Oxford University Press in advance. Please go ahead with the use ensuring that a full acknowledgment is
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111
Kathy R. Chaurasiya designed single molecule experiments, analyzed the data, and wrote the
manuscript; Clarissa Ruslie and Samer Lone performed single molecule experiments and
analyzed the data; Michelle C. Silva designed bulk biochemical experiments, purified proteins,
performed experiments, analyzed the data, and wrote the manuscript; Lukas Voortman purified
proteins and performed bulk experiments; Philip Nevin performed docking studies; Penny J.
Beuning and Mark C. Williams designed experiments and wrote the manuscript.
3.1 INTRODUCTION
DNA polymerase III (DNA pol III) holoenzyme is a ten-subunit protein complex that efficiently
and accurately replicates the entire genome of Escherichia coli (1, 2). It is composed of three
subassemblies: the polymerase core, the β processivity clamp, and the clamp loader complex.
Polymerase and proofreading activities are conducted by the core subassembly which consists of
the polymerase subunit α, the proofreading subunit ε, and the θ subunit, which has a role in
stabilizing the core (3, 4). The β processivity clamp encircles the DNA and provides a platform
for the polymerase core to bind, providing α with access to the primer-template and facilitating
processive replication. The clamp loader complex, which consists of the γ, δ, δ′, τ, χ, and ψ
subunits, loads the β clamp onto the DNA (5) with τ tethering the polymerase core to the
replisome (6), and coordinating simultaneous replication of the leading and lagging strands of the
replication fork (6, 7).
112
Although DNA pol III efficiently replicates undamaged DNA, replication is disrupted upon
encountering damaged bases (8-11). Formation of a RecA filament on accumulated ssDNA
triggers the SOS response (12), resulting in the upregulation of genes encoding numerous
proteins involved in DNA damage repair and tolerance (13). These proteins include the
potentially mutagenic Y-family polymerases DNA pol IV (DinB) and DNA pol V (UmuD′2C)
(14-16). Replication of damaged DNA can proceed once DNA pol III α is replaced with one of
these Y-family polymerases, which can replicate damaged DNA in a process known as
translesion synthesis (TLS) (17-20). DNA pol V is composed of two subunits, the cleaved form
of UmuD and the UmuC polymerase. DNA polymerase manager protein UmuD regulates the
cellular response to DNA damage in part, along with UmuC, by decreasing the rate of
replication, thereby allowing time for non-mutagenic DNA repair processes to occur (21-23).
UmuD undergoes a RecA/ssDNA-facilitated auto-cleavage of 24 amino acids of its N-terminal
“arms” to form UmuD′. UmuD forms a tight dimer (UmuD2), which is the predominant form for
the first 20–40 minutes of the SOS response after which the cleaved form UmuD′ is the
predominant species (22, 24). Although UmuD and UmuD′ are expected to be dimeric under all
the conditions studied here (25) for simplicity we will refer to these dimeric forms as UmuD
rather than UmuD2 and UmuD′ rather than UmuD′2, respectively. UmuD also regulates
mutagenesis in the cell through its interaction with the Y-family DNA polymerase DinB, by
inhibiting DinB-dependent -1 frameshift mutagenesis (14, 26, 27).
UmuD interacts with several components of DNA pol III, including the polymerase subunit α,
the β clamp, and the proofreading subunit ε (28). Recent ensemble biochemical experiments have
shown that there are two UmuD binding sites on the α subunit, one in the N-terminal domain and
113
one in the C-terminal domain (29) (Figure 3.1). The C-terminal binding site (residues 956-975),
which has higher affinity for full-length UmuD relative to the cleaved form UmuD′ (29), is
adjacent to the β clamp binding site (residues 920-924) (30), which tethers the polymerase to its
DNA template. UmuD, but not UmuD′, releases α from the β clamp, which may inhibit DNA
replication and facilitate polymerase exchange (29). The C-terminal UmuD binding site of α is
also adjacent to the OB fold (residues 975-1160), through which α binds ssDNA (31), suggesting
that UmuD may be competing with ssDNA for binding to α. We hypothesized that one way
UmuD contributes to a DNA damage checkpoint is by disrupting the interaction between α and
ssDNA, thereby inhibiting replication. To test this hypothesis, we have used single molecule
DNA stretching to quantify α binding to ssDNA in the presence of wild-type UmuD and several
UmuD variants designed from a computational docking analysis of the complex. We find that
wild-type UmuD competitively inhibits α binding to ssDNA through UmuD-α interactions, while
a single amino acid substitution, D91K, in UmuD disrupts this inhibition.
Figure 3.1 Diagram of DNA pol III α, with domain labels within the boxes and known
interaction sites above the boxes (sequence numbering shown below).
The two UmuD binding sites, one in the N-terminal domain and one in the C-terminal domain of
α, are shown in yellow (29). The CTD binding site (residues 956-975) is adjacent to the β clamp
114
binding site (residues 920-924, shown in blue) (30) and recent biochemical experiments show
that UmuD displaces α from the β clamp (29). This UmuD binding site is also adjacent to the OB
fold (red), where ssDNA binds α (31), an observation that led us to hypothesize that UmuD also
inhibits α binding to ssDNA.
3.2 MATERIALS AND METHODS
3.2.1 Proteins and plasmids
Wild-type UmuD was expressed from the pSG5 plasmid in BL21 (DE3) (Novagen) as
previously described (32, 33). UmuD D91 and G92 were changed to lysine by site-directed
mutagenesis of pSG5 using the QuikChange Kit (Agilent) and confirmed by sequencing the
resulting plasmids (Macrogen USA). Wild-type UmuD and all variants were purified as
previously described (32).
Wild-type DNA pol III α was expressed from the pET28a-α plasmid in Tuner competent cells
(Novagen) and purified using both a Nickel His-trap column (GE Healthcare) and a heparin
column (GE Healthcare) as previously described (29). Fractions collected after the heparin
column were diluted 6-fold with buffer HA (50 mM HEPES, pH 7.5; 1 M NaCl; 2 mM beta-
mercaptoethanol; 20% glycerol) and loaded onto a hydroxyapaptite column (BioRad Bioscale
Mini CHT Type 1, 5-mL, 40 μm cartridge) to concentrate the protein; protein concentrator
devices were avoided because they significantly reduced the stability and activity of DNA pol III
α. After washing with 10 column volumes (cv) of buffer HA, buffer HB (100 mM sodium
phosphate, pH 6.5; 1 M NaCl; 2 mM beta-mercaptoethanol; 20% glycerol) was used to elute the
115
protein from the column isocratically in 2 cv. Fractions containing DNA pol III α were dialyzed
against protein storage buffer (50 mM HEPES, pH 7.5; 150 mM NaCl; and 50% glycerol) and
stored at -20 °C.
3.2.2 Single molecule DNA stretching
In DNA stretching experiments with optical tweezers, a single λ DNA molecule is captured
between two polystyrene beads inside a flow cell. One bead is fixed on a micropipette tip, and
the other is held in a dual-beam optical trap (34). As the fixed bead is extended at 100 nm/s, the
tethered DNA molecule exerts a force on the trapped bead, which is measured by deflection of
the trapping laser beams. The force on the DNA molecule is measured as a function of DNA
extension. As the DNA is stretched, the dsDNA helix undergoes a force-induced melting
transition into ssDNA (Figure 3.2A). The bases anneal upon DNA release, exhibiting minimal
hysteresis, or mismatch between the extension and release curves. After the DNA is stretched
and released in buffer only (10 mM HEPES, 100 mM Na+, pH 7.5), the solution in the flow cell
is exchanged to include protein. Subsequent force-extension curves are obtained in the presence
of α, UmuD, or both proteins.
116
Figure 3.2 DNA pol III α binding to ssDNA characterized with single molecule force
measurements.
(A) Typical extension (solid black) and return (dashed black) of a single DNA molecule. The
molecule undergoes a force-induced melting transition from dsDNA (red, Equation 3.2) to
ssDNA (blue, Equation 3.3) at 62.6 ± 0.5 pN. (B) In the absence of protein (black), the DNA
molecule anneals immediately upon release, exhibiting minimal hysteresis, or mismatch between
the stretch (solid) and release (dashed) curves. The force extension curves in the presence of 500
nM UmuD are the same within uncertainty as those of DNA only (purple), which shows that
UmuD does not measurably bind DNA. In the presence of 250 nM α (green), pausing at a fixed
DNA extension (dashed arrow) after the melting transition exposes ssDNA to α for 30 minutes,
as previously described (31). Upon DNA release, α remains bound to the ssDNA (open circles),
prohibiting the two strands from annealing. The DNA molecule is therefore a combination of
dsDNA and protein-bound ssDNA, and fitting the release curve (open circles) to Equation 3.1
(solid line) yields the fraction of ssDNA bound to α (fss = 0.58 ± 0.02).
Force-extension curves in the presence of 250 nM α alone exhibit significant hysteresis, since
protein bound to the exposed ssDNA prohibits the two strands from annealing upon DNA
release. All single molecule experiments were performed by waiting at fixed extension for 30
min prior to DNA release, which has been established as a quantitative method of characterizing
α binding to ssDNA (31). The fraction of ssDNA bound fss may be described in terms of
observed length b (31):
0
10
20
30
40
50
60
70
0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60
Forc
e (
pN
)
Extension (nm/bp)
dsDNA ssDNA
250 nM α
DNA only
pause: 30 min
500 nM UmuD
0
10
20
30
40
50
60
70
0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60
Forc
e (
pN
)
Extension (nm/bp)
A B
117
ss ds ss ss ss( ) 1b F b f b f
(Equation 3.1)
where bds as a function of force F is described by the Worm-Like Chain (WLC) model:
1
2B
ds ds
ds ds
k1( ) 1
2
T Fb F B
P F S
(Equation 3.2)
with persistence length Pds, end-to-end or contour length Bds, and stretch modulus Sds. The
Freely-Jointed Chain (FJC) model describes the polymer elasticity of ssDNA:
ss Bss ss
B ss ss
2 k1( ) coth 1
k 2
P F T Fb F B
T P F S
(Equation 3.3)
The WLC and FJC polymer models shown in Figure 3.2 have typical parameter values (Bds =
0.34 nm/bp, Pds = 48 nm, and Sds = 1200 pN in Equation 3.2, Bss = 0.55 nm/bp, Pss = 0.75 nm,
and Sss = 720 pN in Equation 3.3). As previously described, the fits were confined to forces
below 40 pN (31) to eliminate effects from changes in the force-extension curve of ssDNA due
to protein binding.
118
These experiments were repeated in the presence of both α and UmuD, and the fraction of α-
bound ssDNA fss as a function of UmuD concentration cs was fit to a simple competitive DNA
binding isotherm:
sapp
dss sat
sapp
d
1
1
c
Kf f
c
K
(Equation 3.4)
where Kdapp
is the apparent equilibrium dissociation constant between UmuD and α in the
presence of ssDNA, and fsat is saturated α-ssDNA binding. A minor correction to added UmuD
concentration c accounts for UmuD bound to α in solution, so effective UmuD solution
concentration cs is (35):
s
a α1
cc
K c
(Equation 3.5)
where Ka = 9.1 x 10-5
M-1
is the equilibrium association constant between UmuD and α in bulk
solution (29) and α concentration cα is 250 nM.
3.2.3 Protein-protein docking
Protein-protein docking models were used to predict residues involved in the binding interaction
between α and UmuD. The structures used for these docking models were a homology model of
119
UmuD (33) and a homology model of full-length DNA pol III α (36). Protein complexes where
predicted by docking DNA pol III α with UmuD using ClusPro 2.0 (37-40), GRAMM-X (41),
and PatchDock (42). The top 10 results from each method were analyzed and compared. Local
docking was performed using the RosettaDock server (43).
3.2.4 Thermal stability assay
A thermal stability assay was used to determine the melting temperature of UmuD variants
relative to wild-type, as previously described (44). To determine whether mutations at positions
D91 and G92 disrupt the stability of UmuD, melting temperatures of these variants were
compared to those of wild-type. Samples containing 20 µM of each variant in 50 mM HEPES,
pH 7.5, 100 mM NaCl and 25x Sypro Orange (Invitrogen) were exposed to temperatures from 25
°C to 80 °C while monitoring the fluorescence emission intensity at 575 nm. Melting
temperatures Tm were determined by taking the first derivative of the melting curves, as
previously described (29).
3.2.5 RecA/ssDNA facilitated cleavage assay
Reactions were assembled as previously described (33) and incubated at 37 °C for 45 minutes.
After incubation, the cleaved product UmuD′ was separated from full-length UmuD using 18%
SDS-PAGE. Bands were then analyzed using the image analysis software ImageQuant TL
(Amersham Biosciences). Control reactions in the absence of RecA, ssDNA and ATPγS were
carried out to determine the amount of UmuD′ present due to spontaneous cleavage.
120
3.2.6 Tryptophan fluorescence assay
The equilibrium dissociation constant Kd between DNA pol III α and the UmuD variants were
determined with a Varian Cary Eclipse Fluorescence Spectrophotometer, as previously described
(29). DNA pol III α (5 µM in 50mM HEPES, pH 7.5 and 100 mM NaCl) was titrated with
varying volumes of 200 - 400 µM UmuD variants. Tryptophan fluorescence quenching was used
to quantitate binding constants, as previously described (29).
3.3 RESULTS
3.3.1 UmuD inhibits α binding to ssDNA
In DNA stretching experiments, a single double-stranded λ-DNA molecule was captured
between two polystyrene beads, one held in an optical trap and the other fixed on a micropipette
tip. As the distance between the beads increases, measurements of the force on the DNA
molecule yield a force-extension curve (Figure 3.2A, solid black line). At a constant force of
62.6 ± 0.5 pN, the dsDNA molecule undergoes a force-induced melting transition to ssDNA.
This overstretching transition has been established as force-induced melting in the presence of
DNA binding proteins such as α, which is the case for the experiments presented in this work
(34). When the tension on the DNA molecule is released (Figure 3.2A, dashed black line), the
ssDNA generated by force anneals immediately into dsDNA, and the curve exhibits minimal
121
hysteresis, or mismatch between DNA extension and release curves. The molecule is a well-
characterized combination of dsDNA and ssDNA along the force-induced melting transition
(45), so waiting at constant extension for a fixed time exposes ssDNA to proteins in solution.
The protein-bound ssDNA exhibits a change in observed length upon DNA release, which is a
direct measurement of protein binding to ssDNA. This single molecule technique has been
established as a quantitative method of characterizing α-ssDNA binding (31). As expected,
constant extension experiments in the presence of 250 nM α exhibit large hysteresis, indicating
significant ssDNA binding (Figure 3.2B). Fits to Equation 3.1 yield the fraction of ssDNA bound
by α, which agrees with previous results obtained by this method (31).
Introducing the DNA damage response protein UmuD disrupts the binding of α to ssDNA
(Figure 3.3A). Constant extension experiments generate ssDNA for 30 minutes in the presence
of both α and UmuD, and the fraction of ssDNA bound by α decreases with UmuD
concentration. Control experiments demonstrate that UmuD does not bind DNA, since force
extension curves in the presence of UmuD are the same within uncertainty as those of DNA only
(Figure 3.2B, purple). Therefore the interaction between UmuD and α inhibits α binding to
ssDNA. A simple DNA binding isotherm (Equation 3.4) fit to the fraction of α bound as a
function of effective UmuD concentration in solution yields the apparent equilibrium
dissociation constant Kdapp
= 340 ± 103 nM between UmuD and α in the presence of ssDNA
(Figure 3.3B).
122
Figure 3.3 The fraction of ssDNA bound by α decreases with increasing UmuD
concentration.
(A) Force extension (solid black line) and release (open circles) curves for DNA in the absence
(black) and presence of α and UmuD (open circles). Open circles are data points, and solid color
lines are fits to Equation 3.1, which yield the fraction of ssDNA bound to α at each UmuD
concentration. Fractions shown in these representative curves are 0.58 ± 0.02 in the absence of
UmuD (green), 0.44 ± 0.02 at 200 nM UmuD (blue), 0.27 ± 0.02 at 500 nM UmuD (red), 0.09 ±
0.01 at 1 μM UmuD (brown), and 0.06 ± 0.01 at 3 μM UmuD (purple). (B) Fraction of ssDNA
bound by α as a function of effective UmuD concentration in solution (Equation 3.5). Error bars
represent standard error (N ≥ 3) for all points except 2.5 μM, which represents propagated error.
The solid line is a χ2 fit to a simple DNA binding isotherm (Equation 3.4) that yields apparent
equilibrium dissociation constant Kdapp
= 340 ± 103 nM between UmuD and α in the presence of
ssDNA, and a saturated α-ssDNA binding fraction (fsat = 0.51 ± 0.05) consistent with previous
single molecule results (31).
3.3.2 Specific UmuD variants disrupt the UmuD-α interaction
To predict potential α binding sites on UmuD, we used three global protein-protein docking
methods. All three methods predicted an ensemble of complexes with UmuD binding near the N-
terminal domain and C-terminal domain of DNA pol III α, consistent with the two previously
characterized UmuD binding sites (29). At the C-terminal domain, a number of docking models
suggested the formation of a salt bridge (Figure 3.4A) between the arginine residue of DNA pol
III α at position 1068 (Figure 3.4B) and the aspartic acid residue of UmuD at position 91 (Figure
0
10
20
30
40
50
60
70
0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60
Forc
e (
pN
)
Extension (nm/bp)
0200500 3000 1000
α (nM):
DNA only
UmuD (nM):
250 250 250 250 250
0
0.1
0.2
0.3
0.4
0.5
0.6
0 500 1000 1500 2000 2500
Fra
ctio
n o
f ss
DN
A b
ou
nd
by
α
UmuD concentration in solution (nM)
A B
123
3.4C). As a result, UmuD residues D91, along with its adjacent neutral residue G92, were each
mutated to lysine in order to disrupt this salt bridge (Figure 3.4C). We did not construct
corresponding mutants in DNA pol III α at position 1068 because such a mutant would likely
disrupt DNA binding as well.
Figure 3.4 Docking model predicts residues involved in the interaction between DNA pol
III α and UmuD.
Pol III α UmuD2
D91
G92
R1068
Pol III α
R1068
fingersdomain
palm domain
thumbdomain
PHP domain
C-terminal domain
D91
G92
UmuD2
A
B
C
124
(A) The docking model predicts a salt bridge between the C-terminal domain of DNA pol III α
(pink) and UmuD (yellow) at α residue R1068 (green) and UmuD residue D91 (blue). UmuD
residue G92 (red) may also be involved in this interaction. (B) The homology model of α (36)
showing arginine residue R1068 (green). The C-terminal domain (residues 917-1160) containing
the binding sites for the β clamp, UmuD, and ssDNA is shown in pink. (C) The homology model
of full-length UmuD (33) showing the residues D91 (blue) and G92 (red) predicted to bind to
DNA pol III α by protein-protein docking studies. The arms of UmuD are shown here in a
“trans” conformation where the arm of one monomer (both arms shown in purple) is bound to
the globular domain of the other monomer (both globular domains shown in yellow). The active
site residues S60 and K97 (cyan) cleave the N-terminal arms (residues 1-24) to form UmuD′.
To verify that the mutations did not destabilize UmuD, the melting temperatures of each UmuD
variant were determined (Figure 3.5A). It should be noted that UmuD has two melting
transitions, which have been assigned to the dissociation of the arms from the globular domain
(see drawing in Figure 3.5A) and the melting of the globular domain, respectively (46). Both
melting transitions of the UmuD variants are comparable to those of wild-type UmuD, indicating
that the variants are properly folded. To test the effect of each mutation on the cleavage activity
of UmuD, a RecA/ssDNA-facilitated UmuD cleavage assay was also performed. All variants
were able to cleave efficiently to form the cleavage product UmuD′ (Figure 3.5B), so these
mutations do not alter UmuD enzymatic activity. Because UmuD can undergo spontaneous
cleavage, control reactions where RecA/ssDNA was not added to the UmuD variants were also
carried out. The amount of UmuD′ present in both the control reactions and the reactions with
RecA/ssDNA were quantified separately to distinguish the amount of UmuD′ produced only in
the RecA/ssDNA facilitated reaction (Figure 3.5B). Thus, the UmuD variants constructed show
similar stability and similar RecA-ssDNA-faciliated cleavage efficiency as wild-type UmuD.
125
Figure 3.5 UmuD variants are structurally stable and enzymatically active.
(A) Melting temperatures of all UmuD variants. The N-terminal “arms” dissociate from the
globular domain (drawing above) in the first transition (dashed), and the globular domain melts
in the second transition (solid). (B) Relative amount of UmuD′ produced by each variant in the
RecA/ssDNA-facilitated self-cleavage reaction (45 min at 37 °C, drawing above), in the absence
(dashed) and presence (solid) of RecA/ssDNA. Thermal stability and self-cleavage activity of all
variants is similar to wild-type UmuD, suggesting that the mutations had minimal impact on
protein stability and function. Error bars represent standard error (N ≥ 3).
As shown in Figure 3.6, in the absence of DNA the UmuD variant D91K disrupted the DNA pol
III α-UmuD interaction by a 15-fold increase in Kd (18 ± 1.7 µM, compared to 1.1 ± 0.6 µM for
wild-type UmuD) while no change was seen with the adjacent UmuD variant G92K (Kd = 1.3 ±
0.5 µM). When both positions were changed to lysine, D91K-G92K, a similar effect of
decreased binding to α was observed (Kd = 33 ± 9.6 µM), confirming this position as a binding
site for DNA pol III α. Although it is initially surprising that an adjacent residue mutation does
not also disrupt UmuD-α interactions, unlike D91, we predict that a side chain at 92 would be
angled away from the surface of UmuD (Figure 3.4A), so it is less likely to participate directly in
the interaction.
UmuD globular domain
UmuD “arms”
“Arms down” “Arms up”
0
10
20
30
40
50
60
Wild-type D91K G92K D91K-G92K
Tem
pe
ratu
re ( C
)
UmuD
UmuD + RecA/ssDNA
UmuD only
UmuD UmuD′
RecA/ssDNA
0
20
40
60
80
100
Wild-type D91K G92K D91K-G92K
Um
uD
′(%
)
UmuD
A B
126
Figure 3.6 Binding curves between pol III α and UmuD variants measured by tryptophan
fluorescence quenching.
(A) The equilibrium dissociation constant Kd is 1.1 ± 0.6 µM. (B) Binding to α is compromised
by the D91K (Kd = 18.1 ± 1.7 µM) mutation relative to wild-type UmuD (29). (C) The G92K
variant (Kd = 1.3 ± 0.5 µM) binds α with nearly the same affinity as wild-type UmuD. (D) In
contrast, the binding affinity of the D91K-G92K variant (Kd = 32.5 ± 9.6 µM) is significantly
weaker than that of the D91K variant. This suggests that both the D91 and G92 residues are
involved in the interaction of UmuD and α. Error bars indicate standard error (N ≥ 3).
127
3.3.3 Specific variants disrupt UmuD inhibition of α binding to ssDNA
We used single molecule DNA stretching experiments to test whether the three UmuD variants
retain the ability to disrupt binding between α and ssDNA. We previously showed using this
single molecule method that α possesses two distinct DNA binding activities: the (HhH)2 domain
binds dsDNA whereas the C-terminal domain containing the OB-fold domain binds ssDNA (31).
This method allows us to probe specifically the binding of α to ssDNA without the potential
complications of DNA secondary structure or dsDNA binding. The biochemical results show
that the D91K mutation compromises the ability of UmuD to bind α, and single molecule
experiments demonstrate that the fraction of ssDNA bound by α in the presence of 1 μM UmuD
D91K is only slightly smaller than that of α alone (Figure 3.7A). UmuD G92K has an affinity for
α similar to that of wild-type UmuD, and DNA stretching shows that this variant retains most of
its ability to inhibit α binding to ssDNA (Figure 3.7B). However, the double mutation D91K
G92K yields a UmuD variant whose binding to α is dramatically weakened, and is completely
unable to disrupt α-ssDNA binding (Figure 3.7C). UmuD residue D91 is therefore required for
UmuD to bind α and disrupt its binding to ssDNA. Residue G92 also participates in the
interactions that are responsible for UmuD inhibition of α-ssDNA binding. Collectively, these
results show that these direct interactions between UmuD and α are responsible for the ability of
UmuD to inhibit α binding to ssDNA.
128
Figure 3.7 UmuD variants have compromised ability to disrupt α binding to ssDNA.
(A-C) Force extension (solid black line) and release (open circles) curves for DNA in the
absence (black) and presence of protein (open circles). Open circles are data points, and solid
color lines are fits to Equation 3.1, which yield the fraction of ssDNA bound by α. (A) The
fraction of α-bound ssDNA in the presence of 1 μM UmuD D91K (blue, 0.54 ± 0.02) is similar
to that of α alone (green). The D91K variant lost the ability to inhibit α-ssDNA binding relative
to wild-type UmuD (brown), which indicates that the D91 residue is required for the interaction
between α and UmuD that disrupts α binding to ssDNA. (B) The G92K variant (red, 0.23 ± 0.02)
partially retains its ability to disrupt the interaction between α and ssDNA. (C) The D91K-G92K
mutation (purple) fully abolishes the ability of UmuD to disrupt α-ssDNA binding, and fraction
of ssDNA bound to α is the same without the UmuD variant (0.58 ± 0.02). (D) Fraction of
ssDNA bound to α in the presence of 1 μM UmuD variants. These single molecule results
indicate that UmuD residue D91 is essential for the interaction between UmuD and α that
disrupts α binding to ssDNA, but G92 is also involved in the interaction.
0
10
20
30
40
50
60
70
0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60
Forc
e (
pN
)
Extension (nm/bp)
DNA only
D91K1 μM UmuD:250 nM α
WT no UmuD
0
10
20
30
40
50
60
70
0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60
Forc
e (p
N)
Extension (nm/bp)
DNA only
G92K1 μM UmuD:250 nM α
no UmuDWT
0
10
20
30
40
50
60
70
0.25 0.30 0.35 0.40 0.45 0.50 0.55 0.60
Forc
e (
pN
)
Extension (nm/bp)
DNA only
no UmuDD91K-G92KWT1 μM UmuD:250 nM α
0
0.1
0.2
0.3
0.4
0.5
0.6
Wild-type D91K G92K D91K-G92K no UmuD
Fra
ctio
n o
f ss
DN
A b
ou
nd
to
α
UmuD
A B
C D
129
3.4 DISCUSSION
In this work, we used a single-molecule method to demonstrate that UmuD inhibits the binding
of α to ssDNA through its interaction with α. The apparent equilibrium dissociation constant
Kdapp
between UmuD and α is 340 ± 103 nM in the presence of ssDNA, which is similar to a
previous measurement of the equilibrium dissociation constant between α and UmuD (Kd = 1.1 ±
0.6 μM) determined by using a tryptophan fluorescence binding assay in the absence of DNA.
The reasonable agreement between apparent binding affinity and direct binding affinity implies
that direct UmuD interactions with α are responsible for ssDNA binding inhibition. In addition,
the measured apparent binding affinity between α and UmuD is within the range of cellular
UmuD concentrations, suggesting that this interaction plays an important role in regulating α
activity in vivo (47).
In addition to directly demonstrating that UmuD inhibits α binding to ssDNA, we used protein-
protein docking models to identify potential UmuD-α interaction sites. The resulting models
predicted that binding between UmuD and α involves UmuD residues D91 and G92 and α
residue R1068. Biochemical experiments confirmed that that corresponding UmuD variants
exhibit a compromised ability to bind α, despite the fact that the variants were demonstrated to
be thermally stable and active for cleavage. Single-molecule experiments showed that the D91K
variant completely fails to disrupt the α-ssDNA interaction, while the G92K variant only
partially inhibits the ability of α to bind ssDNA. These results demonstrate that a direct α-UmuD
interaction at these residues is responsible for UmuD inhibition of α-ssDNA binding. Another
UmuD variant at position 91, D91A, has been shown to disrupt the UmuD-DinB interaction (26),
130
suggesting the existence of a general binding site for polymerases on UmuD. In addition, peptide
mapping experiments show that this region of UmuD provides a binding site for the protease
ClpXP, which plays a role in modulating mutagenesis by degrading UmuD and UmuD′ as well
as DinB (48, 49). Furthermore, the UmuD G92D mutation is poorly cleavable, but when this
mutation was constructed in the context of UmuD′, cells harboring this variant were mutable to a
similar extent as cells harboring wild-type UmuD (50, 51), suggesting that the G92D mutation
does not alter the ability of UmuD′ to facilitate UmuC-dependent mutagenesis. On the other
hand, the UmuD G92C variant was as proficient for cleavage as wild-type UmuD (52).
Therefore, this region of UmuD seems to be an important site for a number of protein
interactions.
The C-terminal domain of α facilitates numerous interactions necessary for efficient replication,
as it interacts with the β clamp (30), ssDNA (31), and the τ subunit of the clamp loader (53, 54).
The interaction between α and the β clamp is essential for high processivity. However, since
replication on the lagging strand is carried out in a series of Okazaki fragments that are
synthesized in a discontinuous manner, the polymerase on the lagging strand must be recycled
for each Okazaki fragment. Although the exact mechanism of the processivity switch is
unknown, the OB fold of α has been implicated as a sensor for ssDNA such that when synthesis
of an Okazaki fragment is completed, the affinity of α for the β clamp is decreased and α is
released from the clamp and from DNA (55-57). Thus, the interaction between α and ssDNA is
proposed to act as a processivity switch. In this work, we demonstrated that SOS-induced levels
of UmuD inhibit binding of α to ssDNA. UmuD also inhibits binding of α to the β clamp, and
these two mechanisms likely work together to facilitate polymerase exchange.
131
UmuD, together with UmuC, specifically decrease the rate of DNA replication and therefore
were proposed to participate in a primitive DNA damage checkpoint (21-23). We previously
showed that UmuD, but not the cleaved form UmuD′, disrupts the binding of α to the β clamp,
which provides a possible molecular explanation for the role of UmuD in a primitive checkpoint
(29). The question then arises of the role of UmuD disruption of ssDNA binding by α. It has
been shown that DNA damage on the lagging strand does not disrupt DNA replication, whereas
DNA damage on the leading strand may severely disrupt replication or may cause leading strand
replication to occur discontinuously as the polymerase can re-initiate synthesis downstream of
the damage (8, 9, 11). Polymerases that encounter DNA damage can also become stalled in a
futile cycle of insertion and excision of nucleotides (58) as they fail to copy the damaged DNA.
Our observations suggest that UmuD would then rescue the stalled polymerase by preventing α
from binding to ssDNA or to the β clamp, thereby allowing DNA repair proteins or translesion
DNA polymerases access to the damaged DNA.
Taken together, our findings suggest that UmuD specifically disrupts the interaction between α
and ssDNA, inhibiting access to the ssDNA template by the replicative polymerase as part of the
primitive DNA damage checkpoint to allow DNA repair. UmuD may also prevent binding of α
to ssDNA to facilitate polymerase exchange, allowing an error-prone TLS polymerase to copy
damaged DNA so that DNA replication may proceed. Our results demonstrate a new mechanism
by which UmuD may regulate the cellular response to DNA damage.
132
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Chapter 4: Replication by E. coli DNA pol III α is inhibited by
direct binding to the OB domain of single stranded DNA binding
protein
Michelle C. Silva designed bulk biochemical experiments, purified proteins, performed
experiments, analyzed the data, and wrote the manuscript; Kiran Prant designed single molecule
experiments, analyzed the data, and wrote the manuscript; Jana Sefcikova designed primer-
extension assay, purified proteins, and wrote the manuscript; Susie Nimipattana performed single
molecule experiments and analyzed data; Mark C. Williams and Penny J. Beuning designed
experiments and wrote the manuscript.
4.1 INTRODUCTION
During replication, genome integrity must be efficiently maintained by DNA polymerases. In
Escherichia coli (E. coli), processive replication of undamaged DNA is accomplished by the
DNA polymerase III (DNA pol III) holoenzyme. It consists of ten subunits which are further
organized into three subassemblies: the core, the β processivity clamp, and the clamp loader
complex (1, 2). Polymerase activity is contained in the core subassembly which consists of the α
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polymerase subunit, the ε proofreading subunit, and the θ subunit. The high processivity of the
holoenzyme is conferred by the β processivity clamp which encircles DNA and tethers the core
to its DNA substrate. This clamp is loaded onto DNA with the help of a clamp loader complex
consisting of six different subunits: τ, γ, δ, δ′, χ, and ψ. The γ subunit along with δ and δ′ load the
β processivity clamp on to DNA with concomitant ATP hydrolysis (1, 2). The τ subunit
coordinates replication on both the leading and lagging strands by coupling the core to the clamp
loader complex (3) while the χ/ψ complex helps regulate replication on the lagging strand (4).
Due to the discontinuous nature of replication on the lagging strand, a significant amount of
ssDNA may be present at one time which is coated with single-stranded DNA binding protein
(SSB) preventing ssDNA from forming secondary structures and protecting the ssDNA from
chemical and nucleolyitc attack (5-7).
SSB is a homo-tetramer with a D2 axis of symmetry (8). Each 178 amino acid-long monomer
consists of an N-terminal globular domain and a disordered C-terminal tail (9). The N-terminal
globular domain of SSB possesses an oligonucleotide/oligosaccharide-binding (OB) domain that
binds ssDNA via stacking interactions with one phenylalnine (F61) and two tryptophan (W41
and W55) residues of SSB (10-15). Changing residue H55 to either tyrosine or lysine has been
shown to disrupt the tetramer interface (16). SSB binds ssDNA by wrapping the DNA around the
homo-tetramer in two binding modes: (SSB)35 and (SSB)65 where the integers 35 and 65
represent the number of nucleotides contacting the protein (17). In the (SSB)35 binding mode,
ssDNA only wraps around two monomers of the homo-tetramer, promoting high cooperativity.
On the other hand, the (SSB)65 binding mode favors limited cooperativity and ssDNA wraps
around all four monomers (17).
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In addition to binding and protecting ssDNA, SSB also serves as a hub for at least 15 proteins
involved in numerous DNA processing pathways, such as proteins involved in homologous
recombination, DNA pol IV (18) and pol V (19) in damage tolerance, and the χ subunit of DNA
pol III in replication (20). Most of these proteins have been found to bind the C-terminal tail
suggesting that SSB uses this tail to facilitate interactions with other proteins involved in DNA
maintenance (21). The temperature sensitive allele ssb-113, encoding the mutation P177S (22),
impairs DNA replication at temperatures higher than 30 °C (23, 24). The P177S mutation has
been shown to decrease the interaction with proteins that bind this C-terminal tail (20).
SSB, in addition to protecting ssDNA present at the replication fork, also has been observed to
directly impact replication activity of DNA pol III. SSB increases processivity of the polymerase
when all of the subunits of DNA pol III are present. In contrast, when only the core subunits α, ε,
θ are present, SSB inhibits replication (25). This inhibition is alleviated when the χ subunit of the
clamp loader complex is present due to a direct interaction between χ and the C-terminal tail of
SSB (4, 26-28).
Although much work has been focused on aspects that suppress inhibition of the α polymerase
subunit by SSB, factors contributing to the inhibition have not been probed in detail. In this
report, we demonstrate that the OB-fold domain of SSB directly binds to the C-terminal domain
of the α polymerase subunit of DNA pol III. We also demonstrate that the α subunit stabilizes
SSB on DNA. Taken together, our findings suggest that the inhibition of DNA pol III by SSB is
due to the direct interaction between SSB and the α subunit and indirectly due to SSB binding
ssDNA.
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4.2 MATERIALS AND METHODS
4.2.1 Proteins and Plasmids
His-tagged DNA pol III α subunit was expressed from pET28a which was provided by Dr.
Meindert Lamers and Dr. John Kuriyan (UC Berkeley) and purified using the protocol
previously described (29). The α truncations were constructed as previously described (30).The
plasmid encoding wild-type SSB, pEAW134, was provided by Dr. Mark Sutton at University at
Buffalo and expressed in BL21-DE3 competent cells (Novagen). Cells were lysed using
lysozyme (Sigma) and purified using the established protocol (31) which includes precipitation
with polymin P and ammonium sulfate, followed by an ssDNA cellulose column. Purified SSB
was stored at -20 °C in a buffer containing 50% glycerol to prevent freezing. High amounts of
glycerol can affect the outcome of various experiments, especially primer extension assays.
Consequently, before every experiment, the buffer was exchanged using Zeba spin columns
(Thermo Scientific) to a buffer appropriate for that assay. The SSB variants F61A, D91N, and
P177S (P177S the mutation encoded by the ssb-113 temperature sensitive allele; this SSB variant
is referred to as SSB-113) were created by site-directed mutagenesis using a QuikChange kit
(Agilent). The plasmid encoding the SSB-OB truncation (residues 1-113), pSSBOB, was
developed from pEAW134 by inserting stop codons in all reading frames after the 113th
residue.
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The disordered C-terminal tail of SSB contains mostly acidic residues (32). Consequently, when
this C-terminal tail is removed as with our SSB-OB truncation, the pI changes from 6 for full-
length SSB to 9 for SSB-OB (33). Therefore, we developed the following method to purify SSB-
OB. Cell pellets from 2 L of cell culture were thawed overnight at 4 °C in SA buffer (50 mM
Tris, pH 8.3; 1 mM EDTA; 20% glycerol; 0.1 M NaCl; 1 mM dithiothreitol; ¼ protease inhibitor
tablet (Roche)) and then cells were lysed using lysozyme (300 µg/mL; Sigma), DNaseI (1 µg/mL
Roche), and a freeze-thaw cycle. Lysate was separated from cell debris by centrifugation at
12,000 x g and 4 °C for 1 hr and sterile-filtered using a 0.45 µm membrane filter (GE
Healthcare).
The lysate was loaded onto a 5 mL HiTrap SP FF column (GE Healthcare) which was
equilibrated with SA buffer. The column was washed with five column volumes of SA and then
eluted with a gradient of 0-100% SB buffer (50 mM Tris, pH 8.3; 1 mM EDTA; 20% glycerol; 1
M NaCl; 1 mM dithiothreitol) over 15 column volumes. Since SSB-OB was not retained on the
column, the flow-through was collected and applied to the column a second time. This allowed
for the separation of impurities that are retained on the HiTrap SP FF column.
The column load and wash were collected and loaded onto 2 x 5 mL HiTrap DEAE FF columns
(GE Healthcare) (10 mL total column volume) equilibrated with SA buffer. The column was
washed with 10 column volumes and then eluted with a gradient of 0-100% SB buffer over 15
column volumes. This column was repeated a second time. Once again, SSB-OB was not
retained on the column and so the DEAE column load and wash were combined for further
purification.
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Ammonium sulfate was added to a final concentration of 1 M. The mixture was then loaded onto
2 x 5 mL HiTrap phenyl-sepharose FF columns (GE Healthcare) (10 mL total column volume)
pre-equilibrated with PSA buffer (50 mM Tris, pH 8.3; 1 mM EDTA; 20% glycerol; 0.1 M
NaCl; 1 mM dithiothreitol; 1 M ammonium sulfate). The phenyl-sepharose column was washed
with 10 column volumes of PSA. SSB-OB eluted approximately halfway through a gradient of
0-100% SA buffer over 15 column volumes. The fractions containing SSB-OB were pooled and
the protein was concentrated using Vivaspin 6 centrifugal concentrators (5,000 MWCO;
Vivascience) to less than 2 mL. At this point, SSB-OB was approximately 75% pure.
This partially purified, concentrated SSB was then injected onto a 300-mL Superdex 75 size
exclusion column (GE Healthcare) using SA buffer. Fractions containing SSB-OB were pooled,
concentrated using Vivaspin 6 centrifugal concentrators (5,000 MWCO) and stored in SSB
storage buffer (20 mM Tris, pH 8.3; 1 mM EDTA; 50% glycerol; 0.5 M NaCl; 1 mM
dithiothreitol) in the same manner as full-length SSB (31). We have found that SSB-OB is
insoluble in buffer that does not contain a reducing agent. After the size exclusion column, SSB-
OB was found to be approximately 90% pure, producing a total yield of 13 mg as quantified by
Bradford assay (Bio-Rad).
4.2.2 Primer-Extension Assay
The 30-nucleotide primer and 100-nucleotide template strands used for the primer-extension
assay (Eurofins MWG Operon) were purified using denaturing polyacrylamide gel
electrophoresis followed by labeling of the primer with 32
P at the 5′ end as previously described
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(34, 35). The sequences are as follows: primer, 5′
GCATATGATAGTACAGCTGCAGCCGGACGC 3′; and template, 5′
GGATAACAATTTCACACAGGAAACAGCTATGACCATGATGGTTACTCAGATCAGGC
CTGCGAAGACCTGGGCGTCCGGCTGCAGCTGTACTATCATATGC 3′. The DNA
substrate was prepared by combining equal amounts (500 nM) of each strand in annealing buffer
(20 mM HEPES, pH 7.5; 5 mM Mg(OAc)2). DNA was annealed by first denaturing at 95 °C for
2 min, incubating at 50 °C for 1 hr and then allowing to cool to 37 °C.
DNA pol III α polymerase activity on a 32
P end labeled primer-template DNA substrate was
assayed in the presence of wild-type SSB and SSB variants as previously described (34).
Reactions were assembled with 50 nM α, 0-500 nM SSB (tetramer concentration) and 100 nM
32P primer-template in primer extension buffer (30 mM HEPES, pH 7.5; 20 mM NaCl; 10 mM
MgSO4; 4% glycerol; 100 µg/mL BSA; 1 mM dithiothreitol). Replication was initiated with 100
µM dNTPs at 37 °C and then quenched with stop buffer (50 mM EDTA; 85% formamide;
0.025% bromophenol blue; 0.025% xylene cyanol) at selected time points (0, 5, 20 and 60
minutes). All reactions were analyzed on a denaturing 12% polyacrylamide gel and imaged using
a phosphor screen and a Storm 860 Phosphorimager (GE).
4.2.3 Tryptophan Fluorescence Quenching Assay
Binding constants were determined by observing the change in intrinsic fluorescence of
tryptophan residues of the α subunit in the presence of varying concentrations of SSB as
previously described (30). Varying amounts of wild-type SSB and SSB variants (100-200 µM
143
monomer stock concentrations) were added to 5 µM α subunit in W buffer (50 mM HEPES, pH
7.5 and 100 mM NaCl). Because SSB also contains tryptophan residues, fluorescence
contributed by SSB alone was subtracted. SSB/ssDNA binding was detected by titrating varying
amounts of 50 µM 24-mer ssDNA (Eurofins MWG Operon) into 5 µM SSB variants (monomer
concentration) in W buffer.
4.2.4 Single Molecule Force Spectroscopy using Optical Tweezers
To label the 3′ and 5′ ends of the same strand, lambda DNA (Roche, 48,500 bp) with two single-
stranded 12 nt 5′ overhangs was digested with ApaI (Fermentas, Fast Digest restriction enzyme)
for 15 minutes at 37 °C to create 3′ and 5′ overhangs on the same strand. The reaction was
terminated by heat inactivation for 20 minutes at 65 °C. In the next step, the nucleotides, biotin-
14-dATP, biotin-14-dCTP (Invitrogen), dTTP and dGTP (Fermentas) were incorporated into 5′-
overhang in the presence of Klenow exo- DNA polymerase for 45 minutes at 37 ˚C . The
reaction was terminated by heat inactivation for 15 minutes at 75 ˚C and was purified using
ethanol precipitation. Biotinylated oligonucleotide (cTcTcTcTctcttctctcttctcttggcc, where T
=Biotin-dT, a gift from Dr. Eriks Rozners, Binghamton University) was annealed to the 3′
overhang to the purified DNA (100:1 oligonucleotide:DNA) by heating the reaction mixture to
65 ˚C for 10 minutes, followed by slow cooling to room temperature. Then, the ligation reaction
was carried out at room temperature for 60 minutes with T4 DNA ligase (Fermentas, Fast
Ligase) and the DNA was purified using ethanol purification. The resulting DNA construct is
38,500 bp long with biotin at 3′- and 5′-ends of the same strand.
144
After successfully labeling both ends of the same stand with biotin, the DNA is captured by
attaching to two streptavidin beads (Bangs Laboratories) where one is trapped in an optical laser
trap and the other is attached to a glass micropipette by suction (Figure 4.1A and the inset of
Figure 4.1C). The force-extension measurements were carried out by measuring the force on
trapped beads as a function of distance (nm/nt) controlled by moving the pipette a known
distance away from the laser trap to stretch the DNA between the two beads (Figure 4.1B). When
DNA was stretched to ~60 pN, a constant force transition was observed in which DNA was
converted from ssDNA to dsDNA (Figure 4.1B). However, when this process was reversed, the
DNA strands anneal and double-stranded DNA is recovered. In order to obtain ssDNA between
the two beads, we used the enzymatic activity of T7 polymerase. T7 DNA polymerization and
exonucleolysis activities can be modulated by controlling the force exerted on the DNA (Figure
4.1B). Exerting a force above 40 pN increases the exonucleolysis activity of T7 DNA
polymerase which is unaffected by the presence of dNTPs (36). In our experiments, we were
able to detect T7 exonuclease activity in real time (Figure 4.1C), which converts dsDNA to
ssDNA . The newly formed ssDNA was then stretched in the presence and absence of SSB. To
investigate the binding affinity of SSB to ssDNA in the prescence of DNA pol III α, the α
subunit was added along with SSB.
145
Figure 4.1 5′ and 3′ biotin-labeled ssDNA was constructed using exonucleolysis by T7 DNA
polymerase.
(A) Schematic diagram of DNA stretching using optical tweezers. 38,500 bp long λDNA was
prepared as described and attached between two beads, one held on the tip of a glass pipette and
the other in an optical trap. (37) (B) Force-extension curves for dsDNA (black solid and dotted
lines) and ssDNA (blue solid and dotted lines). The arrows show the direction of polymerization
and exonucleolysis. (C) When dsDNA is stretched in the presence of T7 DNA polymerase and
held at ~50 pN (red solid line), dsDNA (black solid and dotted lines) is converted to ssDNA
(blue solid and dotted lines) by exonucleolysis.
146
4.3 RESULTS
4.3.1 DNA pol III α interacts with the OB fold of SSB
It has been previously determined that SSB inhibits replication by the core subunits of DNA pol
III (25). Under our conditions, in which a 5ʹ 32
P-labeled primer-template was extended by the α
subunit of DNA pol III in the presence of wild-type SSB, we observed that inhibition increases
with increasing concentration of SSB (Figure 4.2), further supporting previous work (25).
Although studies have been conducted to determine how this inhibition is overcome (4, 26-28),
the mechanism of inhibition has not been probed in detail.
Figure 4.2 SSB inhibits primer extension by DNA pol III α.
(A) A 5′ 32
P-end labeled primer is extended by DNA pol III α in the presence of wild-type SSB.
Polyacrylamide gels were used to resolve the extended primer. (B) As the SSB concentration is
increased, bands representing full or partial extension of the DNA primer become less intense
indicating inhibition of DNA replication by SSB.
147
The observations that SSB inhibits the α subunit, but stimulates the activity of other DNA
polymerases such as DNA pols II and IV (18, 38), suggest that there may be a direct interaction
between α and SSB. To test this, a tryptophan fluorescence quenching assay was used to detect
binding. When increasing amounts of wild-type SSB were added to the α subunit, a fluorescence
quenching effect was observed, confirming that wild-type SSB binds to the α subunit (Figure
4.3). The equilibrium dissociation binding constant KD was determined to be 1.8 ± 0.4 µM
(Figure 4.3), suggesting that SSB has a higher affinity for the α subunit than for the χ/ψ complex
whose binding constant KD was previously determined to be 8.9 ± 0.7 µM (28).
Figure 4.3 SSB binds DNA pol III α with a dissociation binding constant, KD, of 1.8 ± 0.4
µM.
148
To localize the α binding site on SSB, the SSB-113 variant, which contains the C-terminal tail
mutation P177S (22), was employed (Figure 4.4). This mutation, which is also associated with
the ssb-113 temperature sensitive allele (22), has been shown to disrupt the interaction between
SSB and its binding partners whose binding sites have been located, including the χ subunit of
DNA pol III (39-41). The binding constant between the α subunit and SSB-113 was determined
to be 0.18 ± 0.1 µM (Figure 4.5A), a 10-fold increase in binding affinity when compared to wild-
type SSB. This suggests that unlike other SSB binding partners, α does not bind the C-terminal
tail of SSB. To determine whether the globular OB-fold domain of SSB binds α, a truncation of
SSB that contains only the OB fold (resides 1-113; SSB-OB) was constructed (Figure 4.4). The
binding constant KD for this truncation was determined to be 2.8 ± 0.2 µM (Figure 4.5A),
suggesting that α binds this domain. To our knowledge, no other protein has yet been shown to
bind directly to this domain of SSB.
To further localize the binding site in this domain, two SSB variants were constructed; SSB
F61A and SSB D91N (Figure 4.4). It is known that the residue F61 interacts with ssDNA by
stacking with the nucleotide bases of ssDNA. Changing this position to an alanine has been
shown to disrupt this interaction (10). The binding constant between SSB F61A and the α
subunit was determined to be 2.2 ± 1.5 µM (Figure 4.5A), similar to that of wild-type SSB
suggesting that SSB binds α and ssDNA at distinct sites of SSB.
The residue D91 was selected as a possible binding site because it lies on the surface of the N-
terminal globular domain. As shown in Figure 4.4, each of the monomers are concave in shape
with the C-terminal tail extending from one end, the ssDNA binding residues located in the
149
pocket, and the D91 residue positioned at the other end. To test whether D91 is involved in
binding α, the SSB D91N variant was constructed. The D91N mutation did not affect the ability
of SSB to bind ssDNA (Figure 4.5B). The binding constant between SSB D91N and the α
subunit was determined to be 10.9 ± 2.3 µM (Figure 4.5A), an approximately five-fold decrease
in binding. We also constructed this mutation in SSB-OB. The truncated SSB with this mutation,
SSB-OB D91N, also showed a slight decrease in binding (4.0 ± 0.7 µM) when compared to
otherwise wild-type SSB-OB (Figure 4.5A). This confirms that the α subunit binds the N-
terminal globular OB fold of SSB.
Figure 4.4 The structure (9) of the SSB homo-tetramer with the SSB variants used in this
work indicated.
The phenylalanine residue at position 61 (blue) is one of the residues responsible for binding to
ssDNA. Changing the residue to an alanine resulted in decreased binding affinity between SSB
and DNA (10). P177S (orange) located in the unstructured and unresolved C-terminal peptide is
the mutation associated with the ssb-113 temperature sensitive allele (22) and has been shown to
disrupt the interaction with various proteins including the χ subunit of DNA pol III (4). D91N
(purple) disrupts binding to DNA pol III α.
150
Figure 4.5 SSB D91N disrupts the interaction with DNA pol III α but not with ssDNA.
(A) KD values were determined for the interaction between DNA pol III α and SSB variants. A 5-
fold decrease in binding was observed with the SSB D91N variant and a 10-fold increase in
binding with SSB-113, suggesting that DNA pol III α binds the OB fold of SSB and not the C-
terminal peptide. (B) The affinity of SSB D91N (♦) for ssDNA is similar to that of wild-type
SSB (▲).
4.3.2 SSB interacts with the C-terminal domain of DNA pol III α
To localize the binding site for SSB on the α polymerase subunit, we used a series of truncations
of α that contain different domains. The first truncation used contained only the N-terminal PHP
domain, residues 1-280. No binding was detected with α 1-280 (data not shown). A truncation
consisting of residues 1-835 includes the PHP domain and the polymerase domain. The binding
constant KD between this α1-835 truncation and SSB was determined to be 36 ± 18 µM (Figure
4.6), a significant 18-fold decrease in binding, suggesting that SSB only very weakly binds the
151
N-terminal 835 residues of the α polymerase subunit. In order to assure that the subunit binds
SSB protein in the active conformation, activity of both alpha subunit variants was assayed for
primer extension activity. Both -835 and -917 are active, with -917 as active as wild-type ,
but -835 significantly less active (data not shown).
The remaining α residues 835-1160 compose the C-terminal “regulatory hub” of α (30, 42-45).
To further localize the binding site in this domain, two additional truncations were used, α
residues 1-917 and α residues 917-1160. The truncation α1-917, along with the PHP and
polymerase domains, contains the two helix-hairpin-helix motifs shown to bind dsDNA (44).
The binding constant KD for α1-917 binding to SSB was 2.5 ± 1.5 µM, comparable to that of
full-length α. The C-terminal truncation, α917-1160 contains the OB-fold domain shown to bind
ssDNA (44), as well as binding sites for the β processivity clamp (42, 43), the τ subunit of the
clamp loader complex (45-48), and the SOS response protein UmuD2 (30). The equilibrium
dissociation binding constant for α917-1160 binding to SSB was 0.54 ±0.43 µM (Figure 4.6).
Taken together, these observations confirm that the C-terminal residues 835-1160 of the α
polymerase subunit of DNA pol III binds to SSB. Localizing this interaction further shows that
SSB binds at the C-terminal domain “regulatory hub” of .
152
Figure 4.6 SSB binds the C-terminal domain of DNA pol III α subunit.
The large difference between the binding constants for the truncation α1-835 and full-length α
indicate that the binding site is between residues 835-1160. The domains of α are labeled in blue
above the schematic of the protein. The previously determined binding sites for UmuD2 (30), and
the ε (49), β (42, 43), and τ (45-48) subunits of DNA pol III are labeled in black.
4.3.3 SSB inhibits DNA pol III α when bound to ssDNA
DNA pol III α activity was assayed in the presence of numerous SSB variants. As seen in Figure
4.2, approximately 150 nM wild-type SSB is needed to significantly inhibit α polymerase
activity. SSB-113 and SSB-OB, whose binding affinities to α are similar to that of wild-type
SSB, showed a similar inhibitory effect consistent with their similar binding affinities (Figures
4.7A and 4.7C). As with wild-type SSB, significant inhibition of α by SSB-113 at 150 nM is
observed and only slightly less inhibition of α by SSB-OB is observed. With the SSB variants
that disrupt binding to the α subunit, SSB D91N and the truncation SSB-OB D91N, a significant
amount of inhibition is observed only at higher SSB concentrations of 500 nM and 250 nM,
respectively, which again is consistent with their binding affinities for α (Figures 4.7B and
153
4.7D). On the other hand, the variant SSB F61A with a similar affinity for α to that of wild-type
SSB, showed less inhibition of α (Figure 4.7E). Significant inhibition was only seen with 500
nM SSB F61A. The F61A mutation has been shown to disrupt the interaction between SSB and
ssDNA but not with α (10) (Figure 4.5), suggesting that in addition to the direct SSB-α
interaction, the interaction between SSB and ssDNA also contributes to inhibition of α subunit
polymerase activity.
154
Figure 4.7 SSB F61A and SSB D91N only minimally inhibit replication by DNA pol III α.
Primer extension by DNA pol III in the presence of (A) SSB-113, (B) SSB D91N, (C) SSB-OB,
(D) SSB-OB D91N, and (E) SSB F61A. The extent of inhibition of DNA pol III α polymerase
activity by SSB-113, SSB D91N, SSB-OB, SSB-OB D91N is consistent with the observed
binding constants. On the other hand, SSB F61A is not as profficient at inhibiting DNA pol III α
as other variants even though its affinity for DNA pol III α is similar to that of WT SSB. This
suggests that both direct binding to DNA pol III α and binding to DNA contribute to the ability
of SSB to inhibit DNA polymerase activity of DNA pol III α.
155
4.3.4 DNA pol III α stabilizes SSB on ssDNA
To determine if the DNA pol III α subunit affects the interaction between SSB and ssDNA,
single-molecule force spectroscopy, a technique involving the stretching of ssDNA with optical
tweezers (37, 50-52), was used with and without SSB and the α subunit. Single-stranded DNA
was generated by using a DNA molecule with biotin on either end of the same strand and using
the exonucleolytic activity of T7 DNA polymerase to remove one strand of DNA (Figure 4.1). A
greater force is needed to stretch ssDNA in the presence of SSB than with ssDNA alone (Figure
4.8). The change in force is directly proportional to the binding affinity of SSB to ssDNA (37,
50).
The ssDNA stretching force increases by 2-3 pN in the presence of 200 nM SSB in 50 mM Na+
(Figure 4.8), indicating that SSB binds to ssDNA at low force. Once the stretching force exerted
on the ssDNA surpasses 15 pN, SSB starts to dissociate from ssDNA and is completely
dissociated around 20 pN (Figure 4.8A and 4.8B), consistent with previous observations (53). At
this point, the stretching curve of ssDNA in the presence of SSB overlaps with that of ssDNA
alone. In contrast, in the presence of 275 nM α subunit, SSB is not completely dissociated from
the ssDNA at 20 pN and more force is required to dissociate SSB (Figure 4.8C). This suggests
that the affinity of SSB for ssDNA increases in the presence of DNA pol III α as the binding of
pol III α alone to ssDNA does not result in any change in ssDNA stretching force within
experimental uncertainty (Figure 4.8B).
156
Furthermore, we have calculated the free energy (ΔG) required to disrupt the SSB-ssDNA
interaction in the absence and presence of DNA pol III α (Figure 4.8C). In the presence of
proteins that bind ssDNA, the DNA stretching force increases and therefore the free energy to
disrupt these interactions can be estimated as the area in between the red and blue solid curves
for starting points at length per nucleotide = 0.35 and ending points at force of 40 pN for both
curves (Figure 4.8C). In the presence of DNA pol III α, the free energy to disrupt the ssDNA-
SSB interaction, is 0.23 ± 0.04 kBT/nt, as compared to 0.10 ± 0.02 kBT/nt for the ssDNA-SSB
interaction only, which shows that the ssDNA-SSB complex is significantly more stable in the
presence of DNA pol III α.
Figure 4.8 DNA pol III α stabilizes the ssDNA/SSB interaction.
(A) Force-extension curves for ssDNA (blue solid line) and ssDNA/SSB complex (red solid
line). (B) Force-extension curves for ssDNA (blue solid line) and ssDNA in the presence of α
(red solid line). (C) Force-extension curves for ssDNA (blue solid line) and ssDNA/SSB
complex (red solid and dotted lines, which represent the stretch and release, respectively) in the
presence of DNA pol III α. All experiments were carried out in 10 mM Hepes, pH 7.5; 45 mM
NaCl (50 mM Na+).
157
4.4 DISCUSSION
In this work, we probed the inhibition of DNA pol III α by SSB. We determined that the N-
terminal globular domain of SSB binds the C-terminal domain of α. By doing so, the α subunit
stabilizes the interaction between SSB and ssDNA. We also observed that along with binding to
the α subunit, SSB needs to be bound to ssDNA for inhibition to occur. Taken together, these
observations provide a better understanding into how replication by DNA pol III is regulated in
E. coli.
Other E. coli polymerases have been shown to bind SSB including the Y family polymerases
DNA pol IV and DNA pol V (18, 19). In both cases, SSB increases the efficiency of these
polymerases. With DNA pol IV, in order to efficiently replicate a template strand coated with
SSB, the polymerase needs to interact with SSB (18). A similar observation, an increase in
processivity, was seen when the RB69 DNA polymerase was fused with its cognate SSB forming
a chimeric protein (54). The interaction between DNA pol V and SSB has been shown to
increase access to the primer termini (19). The major difference between these interactions and
the α-SSB interaction is that these polymerases bind the C-terminal tail (18, 19) while the α
subunit binds the N-terminal globular domain of SSB.
As demonstrated in previous works and further probed here, the effect of SSB on DNA pol III is
more complex. Even though SSB increases the processivity of the assembled holoenzyme, SSB
158
also inhibits replication when only the α polymerase subunit is present. The mutation associated
with the temperature sensitive allele ssb-113 (23), P177S (SSB-113) has been shown to disrupt
the interaction between SSB and other proteins (20). Interestingly, the opposite is seen with the α
subunit, as P177S actually increases the affinity of SSB for the α subunit. It has been previously
suggested that the C-terminal tail of SSB binds to the DNA-binding pocket in the OB fold when
no DNA is present (55). If so and if the P177S mutation also disrupts this interaction, the N-
terminal globular domain of the SSB-113 variant may be more exposed than that of wild-type
SSB. Because α binds the N-terminal globular domain, this would provide α better access to
SSB, and thus an increase in affinity is observed with the SSB-113 variant.
The χ subunit of DNA pol III has been shown to suppress the inhibition of DNA pol III caused
by SSB, by binding to the C-terminal tail of SSB (4, 26-28). As with other proteins, the SSB-113
variant disrupts the interaction with χ. When the polymerase activity of the α subunit was tested
in the presence of the SSB-113 variant, more inhibition was observed than with wild-type SSB.
This suggests that the disruption in DNA replication observed with the ssb-113 allele is a direct
result of the inability of the χ subunit to alleviate DNA pol III inhibition and the increased
affinity for the α subunit.
Taken together, our findings suggest that the inhibition of α replication by SSB is directly due to
the interaction between α and SSB and indirectly due to SSB binding to ssDNA. Moreover, the
presence of α stabilizes SSB on ssDNA, further enhancing the inhibition. Thus, DNA replication
in E coli is regulated in multiple layers by interactions between α, SSB, and ssDNA.
159
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163
Chapter 5: The E. coli single stranded DNA binding protein SSB
binds the UmuD2 polymerase manager protein
Michelle C. Silva designed bulk biochemical experiments, purified proteins, performed
experiments, analyzed the data, and wrote the manuscript; Arianna DiBenedetto, Celeste Dang,
Christine Alves, and Monyrath Chan performed experiments and analyzed the data; Penny J.
Beuning designed experiments and wrote the manuscript.
5.1 INTRODUCTION
The SOS-induced umuD gene products, full-length UmuD and the auto-cleavage product
UmuD′, contribute to the overall regulation of DNA replication in Escherichia coli. Full-length
UmuD2 is a very tight homodimer (KD <10 pM) (1) and so is expected to be in its dimer form
under all conditions reported here. For the sake of clarity, we will refer to the dimer UmuD2
simply as UmuD throughout this report. Each UmuD monomer consists of two domains; a
dynamic N-terminal arms domain containing the cleavage site (between residues C24 and G25)
and a globular C-terminal domain containing the active site (S60 and K97). Due to the dynamic
nature of the arms, they can exist in various conformations: in an “arms down” conformation
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where the cleavage site is positioned in the active site, and an “arms up” conformation where the
arms are unbound exposing the globular domain (1-4).
During the first 40 minutes after the SOS response is initiated, full-length UmuD is the
predominant form (5, 6). It is believed that during this time, UmuD participates in a primitive
DNA damage checkpoint by inhibiting DNA replication and allowing time for accurate DNA
repair processes to act (6-9). By binding to the α polymerase and β processivity clamp subunits
of DNA polymerase III (DNA pol III), UmuD inhibits interactions between α and β and α and
ssDNA, likely contributing to the primitive DNA damage checkpoint (3, 7, 10, 11).
After these initial 40 minutes, the RecA-ssDNA nucleoprotein filament-facilitated auto-cleavage
product, UmuD′ becomes the predominant form resulting in the activation of translesion
synthesis (TLS) (6, 12, 13). TLS is the potentially mutagenic process in which DNA lesions are
bypassed by a group of specialized DNA polymerases known as the Y-family polymerases (14-
21). In E. coli, two of the five DNA polymerases are part of this specialized group. They include
DNA pol IV (DinB) and DNA pol V (UmuD′2C) both of which are regulated by the umuD gene
products (22-24). It has been shown that along with RecA, UmuD prevents DNA pol IV from
creating -1 frameshift mutations (25). DNA pol V contains two subunits, UmuD′ and the UmuC
polymerase subunit (22, 23). UmuC is specifically activated for TLS in the presence of UmuD′
and RecA that has been activated by binding to ATP and ssDNA (26).
E. coli SSB has been shown to be a key player in numerous DNA processes such as DNA
replication, recombination, and repair due to its ability to bind single-stranded DNA (ssDNA)
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present at the replication fork (27-29). It is a homotetramer where each monomer contains an N-
terminal globular domain and an inherently disordered C-terminal tail (30, 31). The N-terminal
globular domain consists of an oligonucleotide binding (OB) fold containing the tryptophan and
phenylalanine residues responsible for binding ssDNA (32-36). Recently it has been shown that
this domain is also responsible for binding the α polymerase subunit of DNA pol III (Chapter 4).
The C-terminal tail has been shown to serve as a hub for numerous proteins including the Y
family polymerases DNA pol IV and DNA pol V (37-39), as the SSB-113 variant (P177S),
associated with the ssb-113 temperature sensitive allele, has been shown to disrupt the
interaction between these proteins and SSB (38-42).
It has been recently shown that SSB increases accessibility of DNA pol V to the primer-termini
through a direct interaction between the C-terminal tail of SSB and the polymerase subunit of
DNA pol V, UmuC (38). In this work, we probed the interactions between SSB and the UmuC
partner proteins full-length UmuD and the cleaved form UmuD′. We find that these interactions
contribute to the regulation of the interface between DNA replication under normal cellular
conditions and under stress.
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5.2 MATERIALS AND METHODS
5.2.1 Proteins, Strains and Plasmids
Strains and plasmids used are listed in Table 5.1. Full-length UmuD (pSG5), the cleaved product
UmuD′ (pSG4) and the variant UmuD S60A were overexpressed in BL21-DE3 (Novagen) and
purified as previously described (3, 43). The variant UmuD S60A contains a mutation in the
active site that greatly reduces cleavage efficiency (3). Dr. Mark Sutton from the University at
Buffalo provided the plasmid pEAW134 that encodes wild-type SSB. The SSB variants SSB-113
(P177S), SSB D91N, SSB-OB, and SSB-OB D91N were constructed as previously described
(Chapter 4). Wild-type SSB and full-length SSB variants were expressed in BL21-DE3
(Novagen) and purified using the established protocol (44). The SSB truncations (SSB-OB and
SSB-OB D91N) were purified as described in Chapter 4. Purified SSB protein was stored in a
buffer containing 50% Glycerol and stored at -20 °C.
The plasmids used for assays of temperature sensitivity contain the operon expressing the genes
umuDC (pGY9739), umuD′C (pGY9738), umuD (pGYDΔC), and umuD′ (pGYD′ΔC) and
encode resistance to spectinomycin (60 µg/mL). The pGYC1 plasmid, encoding only umuC, was
constructed from the pGY9739 plasmid by inserting XhoI restriction sites before and after the
umuD gene, followed by digestion with XhoI and ligation. The restriction site were inserted by
site-directed mutagenesis using a Quikchange kit (Agilent Technologies) and the primers Cxho
(CCACGTCGTTAAGCTCGAGCGCTGATGTTTGC) and DCxho
(CAGTATAACTCGAGGCAGATTATTATGTTG) (Operon). The strain PAM33 (provided by
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the Coli Genetic Stock Center at Yale University), which contains the temperature sensitive
allele ssb-113, were grown in standard Luria broth media at 30 °C unless otherwise specified.
Competent cells of this strain were prepared using the CaCl2 method (45).
Table 5.1 Strains and plasmids used.
Genotype Reference
Strain
BL21-DE3 F– ompT hsdSB(rB–, mB–) gal dcm (DE3) Lab stock
PAM33 sulA1, trpE65(Oc), [malB+]K-12(λ
S), ssb-113(ts) (40)
Plasmid
pSG5 umuD, AmpR (43)
pSG4 umuD′, AmpR (43)
pEAW134 ssb, pET21a derived, AmpR (46)
pGB2 Vector; pSC101 derived, Specr (47)
pGY9739 oC
1 umuD+; umuC
+; pSC101 derived, Spec
r (48)
pGY9738 oC
1 umuD′+; umuC
+; pSC101 derived, Spec
r (48)
pGYDΔC oC
1 umuD+; ΔumuC; pSC101 derived, Spec
r (8)
pGYD′ΔC oC
1 umuD′+; ΔumuC; pSC101 derived, Spec
r (49)
pGYC1 oC
1 ΔumuD; umuC+; pSC101 derived, Spec
r This work
5.2.2 Tryptophan Fluorescence Assay
All equilibrium dissociation constants (KD) were determined by observing the intrinsic
fluorescence of tryptophan residues as previously described (11). Before each experiment, the
SSB buffer was exchanged to Trp Assay buffer (50 mM HEPES, pH 7.5; 100 mM NaCl) using
zebaspin desalting spin columns (Thermal Scientific) to remove excess glycerol. Varying
volumes of 200-400 µM full-length UmuD or the cleaved product UmuD′ (monomer
concentration) were titrated directly into 5 µM wild-type SSB or SSB variants (monomer
concentration) in Trp Assay buffer. Samples were excited at 278 nm and the emission was
168
recorded between 300 and 400 nm. Peak maxima were used to calculate the fraction quenched
(Q), which was then was plotted versus protein concentration. All binding curves produced by
this method were fitted as previously described (11).
5.2.3 Quantitative Transformation Assay
To determine the effect of elevated levels of the umuDC gene products on the temperature
sensitivity of the ssb-113 allele phenotype, transformation assays were used. Approximately 0.2
µg of each respective plasmid was added to 40 µL of PAM33 competent cells. To induce intake,
cells underwent a 10 minute incubation on ice, a 5 minute heat shock at 37 °C, and an additional
10 minute incubation on ice, after which the cells were allowed to recover for 2 hours at 30 °C in
400 µL of Luria broth with gentle shaking. Half of each transformation sample (200 µL each)
was plated on each of two Luria broth agar plates supplemented with spectinomycin (60 µg/mL)
which were then incubated at 30 °C and 37 °C, respectively. After 48 hours, cell colonies were
counted and the ratio between the non-permissive (37 °C) versus the permissive (30 °C)
temperatures were recorded. The assay was repeated at least three times.
5.2.4 Bis-maleimidohexane (BMH) Cross-linking
Bis-maleimidohexane (BMH) is used as a cross-linking agent to analyze the dynamic N-terminal
arms region of UmuD (43). Reactions contained 10 µM UmuD S60A (monomer concentration),
40 µM SSB or SSB variant (monomer concentration) with and without 50 µM ssDNA (25-mer;
Operon) in BMH buffer (10 mM Sodium Phosphate, pH 6.8; 100 mM NaCl). To initiate cross-
169
linking, 1 mM BMH was added and reactions were incubated at room temperature for 5 minutes,
after which 50 mM DTT was added to quench the reactions. Covalently cross-linked UmuD
dimers were separated from unreacted UmuD using 18% SDS-PAGE. An immunoblot was
employed to detect UmuD-containing bands using rabbit anti-UmuD as the primary antibody as
previously described (3, 43). The variant UmuD S60A was used to eliminate the possibility of
UmuD′ contamination.
5.3 RESULTS
5.3.1 SSB binds the umuD gene products
A previous study has shown that the presence of SSB increases accessibility of the 3′ primer
terminus at the site of DNA damage for the TLS polymerase DNA pol V (38). This study also
showed by immunoprecipitation that the polymerase subunit of DNA pol V, UmuC, directly
binds to wild-type SSB (38). This study led us to hypothesize that SSB binds to the other subunit
of DNA pol V, UmuD′ and its precursor, full-length UmuD, which has been shown to regulate
replication by binding to proteins essential to DNA replication (3, 9, 11, 49, 50).
To determine if the umuD gene products bind to SSB, a tryptotphan fluorescence assay was
employed. This assay involves observing the intrinsic fluorescence of the tryptophan residues of
SSB in the presence of varying concentrations of full-length UmuD or UmuD′, which have no
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tryptophan residues. As shown in Figure 5.1, fluorescence quenching was observed with
increasing concentrations of UmuD or UmuD′. This indicates that SSB does indeed bind both
proteins. The equilibrium dissociation constants (KD) for the SSB-UmuD and SSB-UmuD′
interactions were determined to be 14 ± 1.7 µM and 6.6 ± 0.7 µM, respectively, suggesting that
UmuD′ has slightly higher affinity for SSB than full-length UmuD.
Figure 5.1 Wild-type SSB binds the umuD gene products.
The curves represent the fraction of SSB tryptophan fluorescence quenched (Q) versus the ratio
of UmuD to SSB concentration fitted as previously described (11). The UmuD′ binding curve
(red) shows a slightly greater affinity for wild type SSB then that of the binding curve for full-
length UmuD (blue) (KDUmuD
= 14 ± 1.7 µM; KDUmuD′
= 6.6 ± 0.7 µM). Binding was detected in
Trp Assay buffer. The error bars represent the standard deviation of four experiments.
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5.3.2 The variant SSB-113 disrupts the interaction with UmuD
Like other SSB binding partners, the polymerase UmuC was found to bind the disordered C-
terminal tail of SSB due to the fact that the SSB-113 variant disrupts this interaction (38). To
determine if UmuD or UmuD′ bind at the same binding site, the variant SSB-113 was utilized.
The dissociation constants for the interaction between UmuD and SSB-113 and between UmuD′
and SSB-113 were found to be 50 ± 13 µM and 34 ± 7.5 µM, respectively, a 4-5-fold decrease in
binding affinity (Figure 5.2). This suggests that, like other proteins that bind SSB, both UmuD
and UmuD′ bind the C-terminal tail of SSB.
Since mutation of the C-terminal tail of SSB was not sufficient to eliminate binding, binding was
also probed using the SSB truncation, SSB-OB. This truncation contains the first 113 residues
containing the OB fold responsible for binding ssDNA and lacks the C-terminal tail known for
binding other proteins. Surprisingly, only a 2-fold decrease in affinity was observed relative to
wild-type SSB (KDUmuD
= 27 ± 5.7 µM; KDUmuD′
= 16 ± 3.7 µM) (Figure 5.2). If the C-terminal
tail were the only binding site, a greater decrease in affinity would be expected with SSB-OB.
This suggests that both the N-terminal globular domain and the C-terminal tail of SSB are
involved in binding to full-length UmuD and UmuD′.
To date, the only other protein shown to interact with this N-terminal globular domain of SSB
has been the α polymerase subunit of DNA pol III (Chapter 4). This α-SSB interaction was
disrupted by substituting Asp91 with asparagine (SSB D91N) (Chapter 4). To test if UmuD or
UmuD′ interacted with SSB in the same manner, binding was probed with the SSB D91N
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variant. A disruption in binding was observed with UmuD′ but not with full-length UmuD
(KDUmuD
= 11 ± 1.3 µM; KDUmuD′
= 22 ± 5.6 µM) (Figure 5.2). This suggests the N-terminal
globular domain may preferentially bind the cleaved UmuD′ over full-length UmuD. The same
mutation in the context of SSB-OB eliminates discrimination between UmuD and UmuD′
(Figure 5.2).
Figure 5.2 Equilibrium dissociation constant KD between UmuD or UmuD′ and SSB
variants measured by tryptophan fluorescence quenching.
The variant SSB-113 (P177S) disrupts the binding between UmuD and UmuD′ (KDUmuD
= 50 ±
13 µM; KDUmuD′
= 34 ± 7.5 µM) suggesting that UmuD and UmuD′ bind the C-terminal tail of
SSB. The truncation of the N-terminal globular domain of SSB, SSB-OB shows only a slight
disruption in binding (KDUmuD
= 27 ± 5.7 µM; KDUmuD′
= 16 ± 3.7 µM). A mutation in this
domain SSB D91N shows a disruption in binding for only UmuD′ (KDUmuD
= 11 ± 1.3 µM;
KDUmuD′
= 22 ± 5.6 µM) when compared with wild-type SSB and UmuD′ (KDUmuD
= 14 ± 1.7
µM; KDUmuD′
= 6.6 ± 0.7 µM). The D91 mutation in the context of SSB-OB eliminates
discrimination between UmuD and UmuD′. Error bars represent dissociation constant standard
error of at least three trials.
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5.3.3 Full-length UmuD but not UmuD′ complements the phenotype of the ssb-113 allele
The ssb-113 allele is an intensely studied ssb allele that confers a temperature sensitive
phenotype for growth at 37 °C on strains that harbor it (40, 42). The PAM33 strain containing the
ssb-113 allele was transformed with plasmids expressing the umuDC genes in order to test
whether elevated levels of UmuD, UmuD′, or UmuC suppress the phenotype. As seen in Figure
5.3A and Figure 5.3B, elevated levels of UmuD and UmuC (expressed from the plasmid
pGY9739) suppress the temperature sensitive phenotype. In contrast, elevated levels of UmuD′
and UmuC (expressed from pGY9738) only partially suppress the temperature sensitivity of the
ssb-113 strain (Figure 5.3A and 5.3B). To further probe the suppression of temperature
sensitivity, PAM33 was transformed with plasmids expressing umuD, umuD′, and umuC
individually. With these plasmids, growth at 37 °C was seen with umuD and umuC but not with
umuD′ (Figure 5.3A and 5.3B). Consequently, these observations show that UmuD and UmuC
but not UmuD′ suppress the temperature sensitivity of the ssb-113 strain. This suggests that the
interactions between UmuD and SSB and between UmuD′ and SSB play different roles in vivo.
174
Figure 5.3 Overproduction of UmuD but not UmuD′ suppresses the temperature sensitive
conditional lethality phenotype of the ssb-113 allele.
pGB2 = empty vector; pGY9739 = UmuDC; pGY9738 = UmuD′C; pGYDΔC = UmuD alone;
pGYD′ΔC = UmuD′ alone; pGYC1 = UmuC alone. (A) The PAM33 strain containing the ssb-
113 allele was transformed with different plasmids. Equal amounts of each transformation
reaction were plated on two to three plates (Luria broth-agar-spec), one for each temperature
tested (30 °C, 37 °C, 42 °C). Plates were incubated at the appropriate temperature for 48 hours.
(B) The ratio of cell growth 37 °C /30 °C is shown. Error bars represent the standard deviation of
at least four independent trials.
5.3.4 SSB does not disrupt the dynamics of the N-terminal arms of UmuD
The N-terminal arms of UmuD are only weakly bound to the globular domain (4). The dynamic
nature of the UmuD arms when UmuD is bound to other proteins can be probed with the thiol-
specific, homobifunctional cross-linking agent BMH (11). Because each UmuD monomer
contains only one cysteine residue at residue position 24, two covalently linked UmuD
175
monomers result from this cross-linking reaction. In order for cross-linking to occur, the two
UmuD cysteine residues, one from each monomer, must be no more than 13 Å away from each
other, which occurs only when the N-terminal arms of UmuD are unbound from the C-terminal
globular domain. Hence this technique is useful to determine if the dynamic nature of the N-
terminal arms of UmuD is altered in the presence of SSB.
The cross-linking efficiency of the variant UmuD S60A in the presence of SSB was compared to
the cross-linking efficiency of UmuD S60A alone. As shown in Figure 5.4Ai, the presence of
wild-type SSB, as well as the variants SSB-113 and SSB-OB, did not significantly change the
cross-linking efficiency of UmuD. The addition of ssDNA had no effect as well (Figure 5.4Aii).
The reaction was repeated multiple times resulting in no statistically significant change in cross-
linking efficiency (Figure 5.4B). These results suggest that the interaction between SSB and
UmuD does not change the dynamic nature of the N-terminal arms.
176
Figure 5.4 The interaction between SSB and UmuD does not affect the dynamic nature of
the N-terminal arms of UmuD.
(A) Percent UmuD-S60A cross-linked by bis-maleimidohexane (BMH) in the presence of wild-
type SSB, SSB-113, and SSB-OB (i) without ssDNA and (ii) with SSB. (B) Bar graph showing
the average % cross-linked UmuD of three independent trials where error bars represent standard
deviation.
5.4 DISCUSSION
In this work, we show that SSB binds both full-length UmuD and the cleavage product UmuD′,
and that SSB has a slight increase in affinity for UmuD′ versus full-length UmuD. As with other
binding partners, binding was localized to the C-terminal tail due to the decrease in binding with
the SSB-113 variant. But unlike other protein binding partners, UmuD and especially UmuD′
also bind the globular N-terminal domain as binding is observed with the SSB-OB truncation. In
vivo, elevated levels of UmuD and UmuC but not UmuD′ suppress the temperature sensitive
phenotype of the ssb-113 allele, suggesting that the interactions between UmuD and SSB and
between UmuD′ and SSB have different cellular consequences.
Due to the fact that UmuD is a key player in managing proteins involved in DNA replication and
translesion synthesis, we hypothesized that UmuD would bind SSB, a protein that is also
involved these DNA processes. As previously documented, full-length UmuD participates in a
primitive DNA damage checkpoint (6, 7, 9). Furthermore, UmuD disrupts the interaction
between the α and β subunits of DNA pol III by binding the two subunits (3, 11, 21), as well as
disrupts the interaction between α and ssDNA (10). In addition, SSB has also been shown to
177
interact with various polymerases in E. coli, including the DNA pol III α subunit (Chapter 4),
DNA pol IV (39) and the UmuC subunit of DNA pol V (38).
Of the approximately 15 proteins to date that have been shown to bind SSB, most have been
shown to interact with the C-terminal tail (37). Only one protein, the DNA pol III α has been
shown to interact with the N-terminal globular domain (Chapter 4). As we have shown here, the
interactions between SSB and UmuD or UmuD′ involve both the globular domain and the C-
terminal peptide of SSB. There are two possible explanations for binding to the C-terminal tail.
Because this region binds various proteins involved with different DNA processes, it has been
suggested that this region serves as a protein hub providing access to DNA primer termini for
these proteins. This increase in accessibility was shown with the interaction between SSB and
UmuC (38) and so it seems possible that the interaction between SSB and UmuD/D′ would serve
the same purpose. The other possibility is that SSB may be required to directly interact to both
subunits of DNA pol V (UmuD′C) to efficiently replicate DNA when the template is coated with
SSB, similar to the increase in replication efficiency seen when DNA pol IV (DinB) binds SSB
(39). On the other hand, the interaction with the N-terminal globular domain of SSB suggests
that interaction of UmuD or UmuD′ with SSB may affect the interaction between DNA pol III α
and SSB.
As previously mentioned, the SSB P177S mutation associated with the ssb-113 allele has a
temperature sensitive phenotype due to the disruption of DNA replication (41). It has also been
suggested that this disruption is due to the increased affinity of SSB-113 for DNA pol III α when
compared with wild-type SSB, which in turn causes the inhibition of replication (Chapter 4). If
178
this is the case, the fact that UmuD suppresses the temperature sensitivity suggests that UmuD
may disrupt the interaction between DNA pol III α and SSB. If so, it would be interesting to see
if UmuD has an effect on the interaction of SSB with ssDNA as well. This would further solidify
the role of UmuD as a manager protein in DNA replication.
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Chapter 6: Conclusions and Future Considerations
In 1985, it was reported that elevated levels of UmuDC inhibit DNA replication (1). Thus it was
concluded that UmuD participates in a temporal primitive DNA damage checkpoint (Figure 6.1).
Since then, it was determined that UmuD and UmuD′ bind the α, β, and ε subunits of DNA pol
III suggesting a role in regulation of DNA replication (2). By far, the most extensively
investigated interaction of these has been the interaction between UmuD and the β clamp (3-6).
In contrast, the interaction with the ε subunit remains to be fully characterized. In this work, we
report that UmuD prevents the α subunit from binding the β clamp and single-stranded DNA
through the direct interaction between UmuD and the α subunit. We have also shown that the
single-stranded DNA binding protein (SSB) binds UmuD and the α subunit. Overall, these
observations suggest that UmuD participates in the primitive DNA checkpoint by removing the α
subunit from the replication fork, thus disrupting replication, and by regulating other proteins
associated with replication through protein-protein interactions.
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Figure 6.1 The umuD gene products regulate E. coli DNA replication in the presence of
DNA damage.
(A) UmuD (purple) disrupts the interactions of DNA pol III α (green) with the β processivity
clamp (blue) and ssDNA, releasing the polymerase from the primer-termini. DNA damage is
represented by a red X. (B) Full-length UmuD (purple) undergoes self-cleavage to the cleaved
form, UmuDʹ. This reaction is facilitated by the RecA/ssDNA filament (blue). (C) UmuDʹ
(purple) activates UmuC (orange) for translesion synthesis (TLS); together, they make up DNA
pol V. The interaction between UmuD and SSB also contributes to the regulation of DNA
replication (not shown).
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UmuD participates in a temporal switch by undergoing RecA/ssDNA facilitated self-cleavage to
UmuDʹ, the cleaved form (7) (Figure 6.1B). This takes approximately 20-40 minutes, after which
UmuDʹ is the predominant product of the umuD gene. Previous work has shown that UmuDʹ
activates UmuC for translesion synthesis (8, 9) (Figure 6.1C). When full-length UmuD is the
predominant product, we now know that full-length UmuD but not UmuDʹ prevents the α subunit
from binding the β clamp and ssDNA (Figure 6.1A). This suggests a timeline for UmuD
regulation of DNA replication. When expression of the umuD gene is first upregulated due to the
SOS response, the role of UmuD is to release the stalled replication fork, after which UmuDʹC or
DNA pol V bypasses the DNA damage. Thus the umuD gene products are important for DNA
regulation under cellular stress. Although much progress has been made, more questions still
remain:
6.1 Does UmuD affect the processivity of DNA pol III?
Because UmuD disrupts the interaction between the α subunit and the β clamp, which is the
major contributor to processivity of DNA pol III (10), the next step would be to ascertain if
UmuD has any effect on processivity. Processivity is referred to as the number of nucleotides
incorporated onto the growing DNA strand per binding event. The α subunit alone exerts low
processivity, approximately 1-10 nucleotides (10). When the β clamp is added, processivity
levels can increase to near the levels for the entire DNA pol III holoenzyme (>50 kb) (10-12).
Because wild-type UmuD also disrupts the α/ssDNA interaction, in order to test processivity, it
185
would be necessary to create a UmuD variant that does not affect the ability of the α subunit to
bind ssDNA.
6.2 Do these proteins bind as a complex?
Now that we have shown that UmuD binds numerous proteins associated with DNA replication,
the next question is whether these interactions occur independently or in complex with each
other. For instance, when UmuD disrupts the interaction between the α subunit and the β clamp,
is UmuD binding to only the α subunit, only the β clamp, or both? In Chapter 3, we showed that
the UmuD variants D91K and D91A disrupt the interaction between the α subunit and UmuD.
Do these variants also disrupt the interaction between the β clamp and UmuD? If these UmuD
variants still interact with the β clamp, the contributions of the UmuD-α versus the UmuD-β
interactions to prevent the binding of α to the β clamp can be tested. Methods such as gel
filtration or pull-downs can be used to determine if the α subunit, the β clamp, and UmuD bind
as a complex.
186
6.3 Does UmuD also have a role in regulating DNA repair?
As previously mentioned UmuD participates in a primitive DNA damage checkpoint by pausing
replication allowing time for other non-mutagenic DNA repair mechanisms to occur (1, 13).
Does this mean that UmuD could contribute to DNA repair mechanisms such as homologous
recombination? For instance, it may be possible that the presence of UmuD increases the
accessibility of proteins involved in repair to the replication fork, thus increasing efficiency.
UmuD is already associated with RecA which is responsible for strand exchange in homologous
recombination. The auto-cleavage of full-length UmuD to UmuD′ is facilitated by the
RecA/ssDNA filament (6, 7). RecA is also needed to activate pol V (UmuD′2C) for TLS and pol
V inhibits RecA-mediated homologous recombination (14, 15). UmuD may be the key to
understanding the transition between repair and translesion synthesis.
Even though there has not yet been a high resolution structure solved for full-length UmuD, new
findings regarding how UmuD regulates DNA replication and mutagenesis have been discovered
within the past few years, including our work presented here. Taken together, these observations
further establish the role of UmuD as a manager protein.
187
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