The intra-S phase checkpoint and its effect on replication fork dynamics in saccharomyces cerevisiae

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Content ii

THE INTRA-S PHASE CHECKPOINT AND ITS EFFECT ON REPLICATION
FORK DYNAMICS IN SACCHAROMYCES CEREVISIAE

by

Shawn Joseph Szyjka

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR BIOLOGY)

December 2008

Copyright 2008 Shawn Joseph Szyjka
ii

Dedication

I dedicate this work to my family. Their undying support, encouragement, and interest
in my interests has helped in my academic life and to shape the person that I’ve
become...

To my parents, Donald and Margaret, for teaching me the value of hard work…I’m
not sure I’ll ever be able to comprehend how they were able to juggle insane work
schedules and the needs of Courtney, Adam, Heather, and myself.

To all of those science projects in elementary school that Dad helped me with just a
“little bit”. Even though I got a little more help than I should have, I really think those
projects are what got me interested in science in the first place…

To my parents for believing that I was a genius at the age of seven when I “solved” the
rubik’s cube in less than 10 minutes…

Last (but certainly not least) I’d like to particularly dedicate this work to the most
loving grandparents I could ever ask for: Nana and Papa. They taught me to be kind,
and that no matter what, there’s always more to learn…

iii

Acknowledgements

I would like to begin by thanking Dr. Oscar Aparicio, my graduate mentor, for
playing a prominent role in my growth as a scientist. Actually, let me take that back,
because I’m not sure that any amount of “thanks” could represent how grateful I am to
Oscar for his guidance…and unparalleled patience. He is a remarkable mentor that
has an amazing knack for finding (and maintaining) the fine balance between guiding
his students and letting them grow as independent scientists. I thank Oscar for his
incredible ability to listen to his students. No matter the size of the question, time of
the day, or day of the week, Oscar always has time to answer questions in his office,
on the phone, or via email. I especially want to thank him for purchasing an espresso
machine for the lab. I’m convinced that it was specifically put in the lab to increase
productivity. (With that said, I probably owe caffeine an acknowledgement
somewhere in this section.) I’m grateful to Oscar for encouraging me to present my
data early during my graduate career at the Salk Institute DNA Replication Meeting.
It was a daunting task at the time but had I not given that talk and interacted with
several scientists at that meeting, at least two of my publications would not have been
possible. I also thank him for including me in the grant writing and paper review
processes. Its experiences like these that have enriched my experience in independent
thought and have instilled confidence in my own abilities. Finally, and most
importantly, I’d like to thank Oscar for creating a lab environment that works hard,
plays hard, and laughs even harder. I cannot imagine a better place to be trained as a
scientist...
iv

I’d like to thank past and present members of the lab; Jennifer Aparicio,
Christopher Viggiani, Yuan Zhong, Simon Knott, Dan Gibson, Fangfang Hu, and Yan
(Sarah) Gan, for creating a unique lab environment where we were able to produce
copious amounts of quality data, and have a lot of fun at the same time. There wasn’t
a day that went by where we didn’t laugh at something (or someone) in the lab. I
especially thank Jennifer Aparicio for sharing her expertise in two-dimensional gel
electrophoresis and for building our DNA microarray technology from the ground up.
Her presence in the lab has been instrumental to all of our successes and without her a
significant portion of my data simply could not exist. I also want to thank her for
numerous helpful discussions and advice on a variety of topics. I’d like to thank
Christopher Viggiani for helping me (whether he’s aware of it or not) to become a
more inquisitive scientist. His thoughtful approach and careful analysis of science has
served as a yard stick with which I constantly measure myself by. I especially want to
thank Christopher for engineering the BrdU incorporation constructs that are used
widely in our lab and in dozens of other labs. These constructs have played an integral
role in our work will continue to do so far into the future. On a less serious note, I
want to thank Christopher for telling my jokes in social settings and taking credit for
them (at least I got to find out that other people found those jokes funny). Finally, I
thank Simon Knott for his invaluable computational contribution to the Rad53 work
presented in Chapter 4. I am grateful that with the touch of a few buttons; he can
make my data look “fancy”. I also thank Simon for hours (maybe more hours than I
should mention) of ping pong games. He used to crush me, but I’m getting better.
Slowly.
v

I thank my entire thesis committee: Oscar Aparicio, Susan Forsburg, Steven
Finkel, Peter Qin and Judd Rice for helpful discussions and suggestions during my
oral exams and at my committee meeting. These meetings have been vital to my
growth as a scientist and have taught me several things: 1) to design experiments that
are informative regardless of the answer, 2) to “think outside of the box”, 3) getting
five professors into one room at the same time is a rite of passage not to be taken
lightly by any graduate student and 4) one should never, EVER say: “gee, I hadn’t
thought of that” during a screening exam.
I thank my friends outside of the lab, especially: Matt Lebo, Laura Sanders,
Daniel Ford, Tom Goldman, Justin Dalton, and Mike Hartenstine for helping to make
graduate school a fun and memorable experience. Nobody ever told me that graduate
school was going to be easy, but the ribs, big buck hunting, carpool rides, and poker
parties sure have helped. I’d also like to thank: Mark LaFratta (“Butchy”), Brian
Gaudaur (“Cheese”), Scott Kowalewski (“Otter”), Joe Kowal (“Crazyjoe”) and Steve
Kolacz who, over the years have taught me about friendship, humility, and that
distance should never define who your friends are.
I want to send a very special thanks to Mandy Benedic for her love, support,
and warm personality. She has an indescribable personality trait that makes my “rainy
days” (i.e. bad experiments) not seem so rainy and my “sunny days” (i.e. publications)
that much brighter. Her easy-going nature and smile bring out the best in me. I am
truly lucky to have her in my life and I have no idea how I got along without her…
vi

I thank the labs of Norman Arnheim, Myron Goodman, Michelle Arbeitman,
and Susan Forsburg for sharing equipment. I especially thank the Forsburg lab for
invaluable input during lab meetings and journal clubs.
Finally, I’d like to thank the Buffalo Sabres for NOT making the playoffs this
year - they must have known that I really needed to work on my thesis.
ix

Table of Contents

Dedication ii
Acknowledgements iii
List of Tables ix
List of Figures x
Abstract xiii
Chapter 1: Introduction 1
Chapter 2: Mrc1 is Required for Normal Progression 25
of Replication Forks in Saccharomyces
cerevisiae

Chapter 3: Pph3-Psy2 is a Phosphatase Complex 49
Required for Rad53 Dephosphorylation and
Replication Fork Restart during Recovery
from DNA Damage

Chapter 4: Rad53 Regulates Replication Fork Restart 73
after DNA Damage in
Saccharomyces cerevisiae

Chapter 5: Conclusions 109

Chapter 6: Materials and Methods 117

References 130

Appendices 144

Appendix A: Fluorescence Activated Cell Sorting (FACS) 144
measures DNA content:
FACS analysis measures DNA content

Appendix B: Two-dimensional gel electrophoresis analyzes 145
replication intermediates:
2D gels examine replication intermediates at
specific loci
x

Table of Contents (continued)

Appendix C: rad5-535 cells are defective for replication 146
restart after MMS exposure:
Cells containing the rad5-535 allele are
defective for replication restart after
MMS exposure

Appendix D: Deactivation of Cdc7 delays completion of 147
DNA replication:
Strains containing the temperature sensitive
cdc7-4 allele are delayed in completion of bulk
DNA synthesis in unperturbed conditions and
during recovery from MMS

Appendix E: Viability, DNA content and 2D gel analysis of 149
mrc1  rrm3
ts
cells:
mrc1 rrm3
ts
cells lose viability and display
decreased tRNA pausing at the restrictive
temperature

Appendix F: The H2A.Z histone variant affects replication 151
timing and bud emergence:
The H2A.Z histone variant affects replication
timing and bud emergence

Appendix G. 2D gel analysis of ptc  pph3 cells during 155
recovery from DNA damage:
ptc2  pph3 cells do not fire the late origin,
ARS603, during recovery from MMS-induced
DNA damage

xi

List of Tables
Table 1: Strain List for Chapter 2 123
Table 2: Strain List for Chapter 3 124
Table 3: Strain List for Chapter 4 127
Table 4: Strain List for Appendices 129

xii

List of Figures

Figure 1: The DNA Damage and Replication Stress responses in yeast 9
Figure 2: Error prone and error free repair pathways in budding yeast 24
Figure 3: Mrc1 is required for normal replication fork progression along 29
chromosome VI
Figure 4: Mrc1 is required for normal progression of DNA synthesis 32
Figure 5: Mrc1 is required for normal replication fork progression along 35
chromosome III
Figure 6: Mrc1-AQ supports normal fork progression 36
Figure 7: Slower progression of Cdc45 in mrc1 cells 39
Figure 8: Loss of Mrc1 function does not enhance fork pausing at 41
tRNA genes
Figure 9: Rrm3 is required for viability of mrc1 cells but not 44
mrc1-AQ cells
Figure 10. Psy2 and Pph3 form a complex that contributes to 52
MMS resistance

Figure 11. PSY2 and PPH3 genetically interact with cell cycle 55
checkpoint components

Figure 12. MMS sensitivities of psy2  and pph3 cells are reduced 56
in rad17  or rad24  backgrounds

Figure 13. Psy2-Pph3 regulates the phosphorylation state of Rad53 58

Figure 14. in vitro desphosphorylation of Rad53 relies on a catalytically 60
active Pph3

Figure 15. WT, pph3, and psy2 cells enter S-phase with similar 62
kinetics in the presence of MMS

Figure 16. MMS recovery kinetics of WT and ybl046w  cells are 63
indistinguishable

xiii

List of Figures (continued)

Figure 17. Psy2/Pph3 promotes efficient replication restart during 64
recovery from MMS exposure in S phase

Figure 18. Psy2-Pph3 has H2AX-independent functions 67

Figure 19: PPH3 is required for progression of replication forks 80
along Chromosome VI during constant MMS exposure
Figure 20: PPH3 is required for progression of replication forks 81
along Chromosome III during constant MMS exposure
Figure 21: PPH3 is required for replication fork restart along 83
Chromosome VI during DNA damage recovery
Figure 22: PPH3 is required for replication fork restart along 84
Chromosome III during DNA damage recovery
Figure 23: Restoration of PPH3 during DNA damage recovery facilitates 86
replication restart
Figure 24: Antagonism of Rad53 promotes fork restart along 89
Chromosome VI in pph3  cells recovering from MMS
Figure 25: Antagonism of Rad53 promotes fork restart along 90
Chromosome III in pph3  cells recovering from MMS
Figure 26: Ptc2 contributes to viability, dephosphorylation of Rad53, and 92
replication restart in pph3 cells
Figure 27: Ptc2 contributes to replication restart in pph3 cells 96
Figure 28: Antagonism of Rad53 activity restores replication and 97
viability of pph3 ptc2  cells
Figure 29: WT and rad53  cells display similar fork rates in MMS 100
Figure 30: A model of Rad53 regulation of replication fork restart 104
Figure 31: Neither Pph3 nor Ptc2 is required for viability of 108
HU-treated cells
Figure 32: A model for DNA replication in mrc1  cells 110

xiv

List of Figures (continued)

Figure 33: Pph3 and Ptc2 dephosphorylate Rad53 during recovery 114
from DNA damage
Figure 34: FACS analysis measures DNA content 144
Figure 35: Two dimensional electrophoresis examines replication 145
intermediates at specific loci
Figure 36: Cells containing the rad5-535 allele are defective for 146
replication restart after MMS exposure
Figure 37: Strains containing the temperature sensitive cdc7-4 allele 147
are delayed in completion of bulk DNA synthesis in
unperturbed conditions and during recovery from MMS
Figure 38: mrc1  rrm3
ts
cells lose viability and display decreased 149
tRNA pausing at the restrictive temperature
Figure 39: H2A.Z affects replication timing and bud emergence 153
Figure 40: ptc2  pph3 cells do not fire the late origin, ARS603, 155
during recovery from MMS-induced DNA damage

xv

Abstract

Duplication of a cell’s genetic material is of paramount importance. This task
must be completed accurately, efficiently and occur once and only once within any
given cell cycle. Origin “firing”, which occurs according to a temporal program, results
in the establishment of bidirectional replication forks. As replication forks traverse the
chromosomal landscape, their stability is threatened by endogenous “obstacles” such as
heavily transcribed tRNAs, protein-DNA complexes and “replication slow zones”. In
addition, fork stability can also be threatened by genotoxic agents. During S phase, in
response to genotoxin-induced stress, the cell activates the intra-S phase checkpoint.
The intra-S phase checkpoint is comprised of three major groups of proteins: sensors,
adaptors and effectors. Sensor proteins detect problems at the replication fork and elicit
a phosphorylation cascade that is mediated by adaptor proteins and results in the
activation of effector kinases. Together, these proteins act to inhibit late origin firing,
slow cell cycle progression, upregulate DNA repair genes, and stabilize replication
forks until the stress has been alleviated.
In response to nucleotide depletion, the adaptor protein, Mrc1 (Mediator of the
Replication Checkpoint), mediates a checkpoint signal that activates the effector kinase,
Rad53. In addition to its checkpoint role, Mrc1 also appears to play a role in
unperturbed DNA synthesis. Mrc1 travels with replication forks and mrc1 cells
display slowed bulk DNA synthesis. This phenotype could be due to decreased origin
firing, increased fork pausing at endogenous “obstacles”, or overall defective fork
progression. Using two-dimensional (2D) gel electrophoresis of replication
xvi

intermediates, we demonstrate that mrc1 cells are not defective in fork pausing at
tRNAs and that Mrc1’s replication function is required for efficient progression of
replication forks throughout the genome. We hypothesize that Mrc1 maintains
association between polymerases and helicases at the replication fork, which results in
efficient fork progression.
In response to MMS-induced DNA damage, Rad53 plays a vital role in
replication fork stabilization. It is thought that Rad53 maintains the association of
replisome components so that forks are poised for efficient restart upon checkpoint
deactivation. Despite a wealth of knowledge with respect to checkpoint activation,
relatively little is known about checkpoint deactivation and how replication forks
restart. Previous work has suggested that replication fork progression along an MMS-
damaged template is independent of the checkpoint. However, recent studies lacking
Psy2-Pph3, a Rad53 phosphatase responsible for Rad53 deactivation after DNA
damage, suggest otherwise, as the completion of S-phase after DNA damage is delayed
in the absence of Psy2-Pph3. We have examined the role of Rad53 in the control of
replication fork restart by monitoring BrdU incorporation at replication forks in the
presence of DNA damage and during recovery from damage. We find that Rad53
deactivation is a key requirement for replication fork restart as cells lacking PPH3 are
defective in fork progression in the presence of DNA damage and are delayed in
replication restart during recovery. In addition, dominant-negative inactivation of
Rad53 in pph3 cells, enables replication fork restart, arguing that deactivation of
Rad53 is sufficient for replication fork restart. Deletion of PTC2, which encodes a
xvii

second, unrelated Rad53 phosphatase, in addition to PPH3, completely eliminates
replication fork progression under DNA damaging conditions and results in lethality.
These findings suggest that replication fork stabilization and restart involve a cycle of
Rad53 activation and deactivation, and that at least two distinct phosphatases are
responsible for regulating this process.
1

CHAPTER 1

Introduction

Checkpoints safeguard genome integrity

The cell undergoes a countless number of ordered events each and every cell
cycle aimed at duplicating its genome and passing a copy onto its progeny.
Maintenance of genome integrity is one of the greatest challenges that the cell faces
each time it enters the cell cycle. Failure to faithfully duplicate a cell’s genome results
in a wide array of problems including: genomic instability, anueploidy, cancer, and
cell death. To safeguard against potential catastrophe within the cell, checkpoints
ensure that certain criteria are met before allowing the cell to proceed into the next
phase of the cell cycle.
All of the work presented in this thesis has been completed in the budding
yeast, Saccharomyces cerevisiae. Saccharomyces cerevisiae is one of the simplest
eukaryotes. It contains many genes with functional homologs in higher organisms and
in many ways provides researchers with a powerful tool to study DNA replication and
cell cycle checkpoints. Budding yeast can be easily cultured, manipulated, and have a
relatively short doubling time. Yeast are somewhat unusual because they can live in a
haploid and diploid state. While in the haploid state, researchers can easily delete
genes and study the resulting phenotypes. In addition, haploid yeast can be mated,
sporulated and dissected, making them an extremely useful organism to study genetic
interactions. Collectively, these qualities make yeast an ideal system not only because
2

they are easy to work with, but they allow researchers to make educated predictions
about proteins and pathways in higher organisms.

Checkpoints: A historical perspective
The first experiments to demonstrate that the cell cycle could be regulated
were performed by Potu Rao and Robert Johnson in 1970. Their experiments showed
that when nuclei from an S phase cell and a G
2
cell were fused, the S phase nucleus
was accelerated and mitosis was inhibited in the G
2
cell – until the two nuclei were
synchronized. Although this phenomenon was likely due to the action of CDKs, it
was the first time that anyone had demonstrated that the cell cycle could be controlled
by transacting factors (Rao & Johnson, 1970). A few years later, by showing that
DNA damaging agents could arrest the cell in G
2
, Robert Tobey postulated that a
surveillance mechanism could function as a “decision point” (Tobey, 1975). That is,
the cell could decide to repair the damaged DNA and re-enter the cell cycle, or leave
the DNA unrepaired and “convert the cell to an inviable state”. Although no data
existed to support the idea, Tobey suggested that different stoppage points within the
cell cycle could also serve as decision points. Shortly thereafter, the first checkpoint
genes in fission yeast, Schizosaccharomyces pombe, were discovered. In the study,
rad1, rad3, and rad9 (RAD = radiation sensitive) mutants failed to delay cell division
after UV-induced DNA damage, thus providing evidence that cell cycle delay is an
active process dependent on several gene products (Hannan et al, 1976). In line with
these findings, Leland Hartwell’s lab set out to further characterize a set of seven rad
mutants in budding yeast (Weinert & Hartwell, 1988). Of the seven, only rad9
3

mutants failed to arrest cell cycle progression after x-irradiation. These mutants
proceeded through the cell cycle and died in subsequent generations. Interestingly,
they found that they could increase the viability of irradiated rad9 mutants if they
were held in G
2
with a microtubule poison, thus allowing time for DNA repair.
The idea of an S phase checkpoint originated from a study in human cells
lacking ATM, an important upstream checkpoint signaling protein. These cells, when
damaged with x-irradiation, displayed increased numbers of replicons and an
increased rate of fork elongation (Painter & Young, 1980). With the cloning of the
budding yeast proteins Mec1 and Rad53, evidence began to mount demonstrating that
the S phase checkpoint was important in safeguarding genomic stability. During a
checkpoint response, Mec1 and Rad53 inhibit late origin firing, stabilize replication
forks, inhibit mitosis and upregulate DNA damage genes (reviewed in Branzei &
Foiani, 2006). Despite evidence in mammalian cells that ATM regulates fork
elongation, active fork slowing in budding yeast has been in dispute. Initially, Leland
Hartwell’s lab demonstrated that cells lacking Mec1 or Rad53 exposed to MMS
cannot restrain bulk DNA synthesis as WT cells do (Paulovich & Hartwell, 1995).
Based on their analysis, it was unclear whether this was due to increased origin firing
or increased elongation rate. In an attempt to add insight into Hartwell’s results,
Tercero and Diffley (2001) used a density hybrid shift approach to examine origin
firing and replication fork progression. They showed that late origin firing, not
elongation rate, was increased in mec1  and rad53  cells. In addition, they showed
that these cells were extremely sensitive to DNA damaging agents when they were
allowed to enter S phase. Taken together, these studies have laid the groundwork
4

describing how cells maintain viability through checkpoint activation. This thesis will
mainly focus on the intra-S phase checkpoint, its components, their role in checkpoint
activation and the proteins/factors that they affect to sustain a checkpoint response. In
recent years, it has become increasingly apparent that checkpoint deactivation is
important to cellular viability. Cells that fail to down regulate the checkpoint signal
during recovery remain arrested in the cell cycle and die. In the following pages, I
will pay close attention to how the cell protects replication forks, thus poising them for
efficient restart.

Origin firing establishes replication forks
Synthesis (S) phase, the period in the cell cycle when DNA is duplicated, is an
extremely important time during the cell cycle. The events leading up to and
including S phase follow a tightly regulated temporal program to ensure that the
genome is copied once and only once per cell cycle. During S phase, replication
forks are established at ARS (autonomously replicating sequence) elements located
throughout the genome (reviewed in Takeda & Dutta, 2005). Within each ARS
element is a specific 11 base pair sequence (ARS consensus sequence, or ACS) that
binds to a complex of six proteins, termed the origin recognition complex (ORC).
ORC remains stably bound to the ACS throughout the cell cycle. However, despite
ORC’s constant association with the ACS, the proteins that bind to ORC do so
according to a precisely controlled sequential program. Additionally, mechanisms
such as Cdc6 degradation, nuclear exclusion of Mcms and phosphorylation of ORC
5

are in place to prevent the cell from rereplicating its DNA before completion of the
cell cycle (Nguyen et al, 2001).
During G
1
, ORC recruits the proteins Cdc6 and Cdt1. Once at the origin, these
proteins are required for the loading of the heterohexameric Mcm2-7 complex, which
licenses the origin for initiation. As cells proceed from G
1
into S phase, additional
proteins such as Mcm10, Cdc45, Sld2, and the GINS complex are assembled into a
complex at the pre-RCs. Along with recruitment of these proteins, the activation of
two independent kinase complexes, CDK (cyclin dependent kinase) and DDK (Dbf4
dependent kinase) are required for origin initiation. Together, these events lead to
origin unwinding, recruitment of the replicative polymerases, RPA binding to single
stranded DNA, and establishment of bidirectional replication forks at the origin.

Fork integrity is challenged under damaging and non-damaging conditions

Endogenous threats
S phase is a vulnerable time during the cell cycle because of the constant
impediments replication forks must contend with while accurately copying the
genome. tRNAs, protein-DNA complexes, and replication slow zones (RSZs) threaten
the progression, stability, and restart of replication forks (Cha & Kleckner, 2002;
Deshpande & Newlon, 1996; Ivessa et al, 2003; Ivessa et al, 2002; Ivessa et al, 2000).
To deal with these fork impediments properly, the cell employs proteins such as the 5’
to 3’ DNA helicase, Rrm3, and the Mec1 checkpoint kinase to promote replication
through these sites. In the absence of Rrm3, cells exhibit a marked increase in
6

replication fork pausing at tRNAs and protein-DNA complexes. rrm3 cells display
elevated inter-chromosomal recombination and increased ribosomal DNA (rDNA)
circles, which arise due to breakage of arrested replication forks at the at the rDNA
locus (Ivessa et al, 2003; Torres et al, 2004). Mec1 plays an important role in
maintaining replication fork integrity at RSZs within the genome. At the restrictive
temperature, mec1 cells exhibit slowed progression through S phase, slowed
replication fork progression, and breakage at individual loci on chromosome III (Cha
& Kleckner, 2002). Similar to the genomic rearrangements found in many human
cancers, mec1  cells are also characterized by increased levels of recombination and
chromosomal translocations (Craven et al, 2002; Pennaneach & Kolodner, 2004).
In the fission yeast Schizosaccharomyces pombe replication forks pause at the
mating type locus. Pausing at the mat1 locus induces recombination, mating type
switching and requires the action of Swi1 (ScTof1) and Swi3 (ScCsm3) (Dalgaard &
Klar, 2000; Dalgaard & Klar, 2001). Swi1 and Swi3 are members of the “replication
fork protection complex”, which associates with origins during initiation, and travel
with replication forks during S phase (Noguchi et al, 2003; Noguchi et al, 2004). In
their absence, Rad22 repair foci and Holliday junctions accumulate, indicating that
these proteins play a constitutive role in maintaining fork integrity. In budding yeast,
Tof1 and Csm3 are required for pausing at the replication fork barrier within the
rDNA locus (Calzada et al, 2005; Mohanty et al, 2006). Furthermore, Tof1 is required
for fork pausing at tRNAs and centromeres (Hodgson et al, 2007). In unperturbed
conditions, Tof1 coimmunoprecipitates with Mcm6 (Nedelcheva et al, 2005).
Interestingly, during exposure to hydroxyurea (HU), tof1  cells lose association
7

between polymerases and Mcms, suggesting that Tof1 acts to slow replication forks in
response to HU (Katou et al, 2003; Tourriere et al, 2005). Taken together, these data
demonstrate that even during unperturbed DNA replication, fork integrity must be
maintained as forks encounter impediments throughout the genome. It will be
interesting to see what the exact mechanism of fork pausing is. Potentially, Mec1,
Swi1 and Swi3 act as “molecular brakes” that promote stable fork progression through
potentially dangerous loci.

Exogenous threats

In addition to endogenous threats, the cell also contends with genotoxin-
induced stresses originating in the lab that cause replication stress and DNA damage.
Hydroxyurea (HU) and methyl methanesulfonate (MMS) result in depletion of
nucleotides and methylation of purines, respectively. These agents cause replication
forks to slow or stall, and at high enough concentrations, MMS can cause breaks in the
DNA. Importantly, if the cell does not deal with these stressors in a timely and
appropriate manner, fork collapse and deleterious genomic rearrangements ultimately
result. To deal with exogenous threats to the genome, the cell employs a group of
proteins, collectively known as the intra-S phase checkpoint. In budding yeast there
are two main branches of this checkpoint. Despite responding to different stressors,
the two branches share some of the same proteins and in some cases either branch is
able to functionally substitute for one another. The first of these pathways responds to
“replication stress”, which is induced by HU. I will refer to this branch as the intra-S
phase replication stress checkpoint (Fig 1A). The second pathway is triggered in
8

response to DNA damage induced by MMS and UV light and will be referred to as the
DNA damage response (Fig 1B). Both responses are made up of three main classes of
proteins; sensors, adaptors and effectors. These proteins sense the genotoxic threat,
and transduce a phosphorylation cascade to effector proteins, which in turn, amplify
the checkpoint signal. Activating the checkpoint signal results in inhibition of cell
cycle progression, upregulation of genes responsible for DNA repair, inhibition of late
origin firing and maintenance of replication fork integrity until the replication
machinery can proceed (reviewed in Branzei & Foiani, 2005). Of these, arguably the
most important responsibility of the checkpoint is stabilization of replication forks.
Checkpoint defective cells that are damaged with HU or MMS display dramatically
reduced survival rates and accumulation of abnormal replication intermediates
indicative of collapsed replication forks (Lopes et al, 2001; Sogo et al, 2002; Tercero
& Diffley, 2001). In the following sections I will introduce the major players in the S
phase checkpoints and how they contribute to genome stability.
9

Figure 1. The DNA Damage and Replication Stress responses in yeast. Nucleotide depletion by
HU (A) and MMS-induced lesions (B) cause replisomes to stall and expose stretches of ssDNA.
“Replisome” is a simplified representation of leading and lagging strand polymerases and helicases
present at the replication fork. “P” represents phosphorylation of respective proteins. See text for
detailed description.

10

Checkpoint Signaling

Sensors trigger the intra-S phase checkpoint

Sensor proteins play perhaps the most pivotal role in the intra-S phase
checkpoint because they are among the first proteins to recognize sites of DNA
damage. In addition, their recruitment does not rely on the localization of other
proteins (Melo et al, 2001; Rouse & Jackson, 2002). In budding yeast, Mec1 and Tel1
are the major sensor proteins, and have homologs in humans known as ATR and
ATM, respectively.
Many parallels exist between Mec1/Tel1 and ATR/ATM. ATR is required for
mammalian embryonic development. Cells lacking ATR accumulate chromosomal
aberrations and die, which is reminiscent of chromosomal abnormalities present within
undamaged mec1  cells (Brown & Baltimore, 2000; Cha & Kleckner, 2002; Cortez
et al, 2001). Like Mec1/Tel1, ATR/ATM are required for checkpoint signaling.
Specifically, ATR responds to UV damage, stalled replication forks and also plays a
role in the double strand break (DSB) response, while ATM primarily responds to
DSBs (reviewed in Shiloh, 2003). Much like mec1 cells, cells lacking either ATM or
ATR are also extremely sensitive to DNA damaging agents. Mec1 and Tel1 are also
functionally divergent. Like ATR, Mec1 responds to replication stress and DNA
damage, while Tel1 responds mainly to DSBs. However, Tel1 does play a small role
in the DNA damage checkpoint (Emili, 1998). Tel1’s contribution is masked by the
prevailing activity of Mec1 and is only appreciated when analyzing the increased
sensitivity of mec1  tel1 cells to ultraviolet radiation (UV) or hydroxyurea (HU)
11

compared to mec1  cells (Morrow et al, 1995). In addition, overexpression of Tel1
suppresses mec1-1 sensitivity to UV and HU. ATR and Mec1 both require a protein
binding partner during initiation of the checkpoint response. In higher eukaryotes,
ATR is bound by ATRIP and directed to sites of damage. In yeast, Mec1 and Ddc2
colocalize to sites of DNA damage and form repair foci. This complex is essential to
initiate signaling as DNA binding by Ddc2 is severely diminished in mec1  cells.
Conversely, ddc2 cells are as sensitive to DNA damaging agents as mec1  cells
(Melo et al, 2001; Paciotti et al, 2000). Together these results demonstrate functional
similarity between Mec1/Tel1 and ATR/ATM. They also suggest that Mec1 and Tel1
act in overlapping pathways in response to DNA damage, however Mec1 plays a
larger role in guarding genomic integrity.
In addition to Mec1/Ddc2, another set of proteins is required for initiation of
the DNA damage checkpoint; the Rad17-Mec3-Ddc1 complex, which is the human
ortholog of the 9-1-1 complex (Rad9-Rad1-Hus1). This heterotrimer bears homology
to Proliferating Cell Nuclear Antigen (PCNA) and is loaded onto ssDNA coated with
RPA via the RFC-Rad24 clamp loader complex (Majka & Burgers, 2003).
Interestingly, a mutant allele of RPA, rfa1-t11, abrogates the loading of the Rad17-
Mec3-Ddc1 clamp by Rad24, resulting in lower levels of checkpoint activation and
suggests that RPA plays an important role in checkpoint signaling (Majka et al, 2006a;
Marini et al, 1997; Pellicioli et al, 2001). Once loaded via Ddc2, Mec1
phosphorylates the Ddc1 and Mec3 subunits of the clamp, the clamp loader, and the
most damage proximal protein, RPA (Kim & Brill, 2003; Majka et al, 2006b).
Phosphorylation of these proteins stimulates the Mec1 kinase leading to the activation
12

of downstream adaptor and effector proteins (Majka et al, 2006a). Together these
results demonstrate a requirement for proximity and proper loading of several players
to efficiently activate the checkpoint. Indeed, the checkpoint can be activated by
artificially colocalizing Mec1/Ddc2 and Rad17-Mec3-Ddc1 to a specific DNA locus,
which likely reflects stringency within the cell to protect against spurious checkpoint
activation (Bonilla et al, 2008).
Mec1 initiates a signaling cascade in response to MMS or HU, but what
exactly is the checkpoint responding to? Likely, when then replication fork reaches a
lesion in the DNA or is inhibited by depletion of nucleotides, fork uncoupling occurs.
That is, leading and/or lagging strand polymerases fail to remain “in step” with the
helicase. As a result, excess ssDNA is unwound. Once a certain threshold is met,
localization of checkpoint sensors occurs leading to checkpoint activation (Garvik et
al, 1995; Lee et al, 1998; Shimada et al, 2002; Tercero et al, 2003).

Adaptors transduce the checkpoint signal from Mec1 to Rad53
During S phase, activated Mec1 phosphorylates downstream components of
the checkpoint. The proteins immediately downstream of Mec1 activation make up
two pathways. In response to MMS or UV induced DNA damage, Mec1 will activate
the DNA damage response by phosphorylating Rad9 (Emili, 1998; Vialard et al,
1998). In response to replication blocks, like those induced by exposure to HU, Mec1
phosphorylates Mrc1 (Alcasabas et al, 2001; Osborn & Elledge, 2003). Ultimately,
both pathways result in the activation of the checkpoint effector kinase, Rad53
(Sanchez et al, 1996; Sun et al, 1996). Despite different cues for activation, both
13

pathways share overlapping functions. That is, mrc1 cells exposed to HU still
activate Rad53, but mrc1 rad9  cells do not (Alcasabas et al, 2001). Similarily,
rad9  cells exposed to MMS retain the ability to activate Rad53, but
mrc1rad9  cells do not. Importantly, neither single mutant is as sensitive to UV or
HU as mrc1rad9 , further supporting the hypothesis that these two proteins act in
parallel pathways, converging on Rad53 (Alcasabas et al, 2001; Osborn & Elledge,
2003).
The adaptor protein Mrc1 is required for full activation of Rad53 during the
intra-S phase replication stress response in both budding and fission yeast. mrc1
cells exposed to HU display long spindles and increased late origin firing, indicative
of a defective checkpoint response. The Mrc1 protein contains 17 S/TQ motifs, which
are essential to transduce the phosphorylation signal to Rad53. Mutation of these
motifs to AQ, results in a checkpoint deficient allele of Mrc1 known as mrc1-AQ
(Alcasabas et al, 2001; Katou et al, 2003; Osborn & Elledge, 2003; Tanaka & Russell,
2001).
In addition to its role in checkpoint activation, Mrc1 also plays an important
role at the replication fork during an unperturbed S phase. Mrc1 associates with
origins around the time of Cdc45 binding, travels with forks during S phase and
coimmunoprecipitates with Mcms (Nedelcheva et al, 2005; Osborn & Elledge, 2003).
Interestingly, mrc1  cells exposed to HU are characterized by an uncoupling of
replisome components (i.e. Cdc45, Mcms) from sites of BrdU incorporation(Katou et
al, 2003). Consequently, these forks are delayed in their restart once HU is removed,
suggesting that Mrc1 helps to maintain a stable replisome when forks slow due to HU
14

exposure (Tourriere et al, 2005). Also, during unperturbed conditions mrc1  (but not
mrc1-AQ) cells are delayed in the completion of bulk DNA synthesis. These cells also
display increased recombination, which could be a result of increased ssDNA at
replication forks due to fork uncoupling (Alcasabas et al, 2001; Osborn & Elledge,
2003; Xu et al, 2004). Delayed completion of S phase could be due to defects in
origin firing, fork elongation, or increased fork pausing at scheduled pause sites within
the genome. In Chapter 2, we test these possibilities by monitoring replication
intermediates in WT, mrc1 , and mrc1-AQ cells along extended replicons on
Chromosomes III and VI. We also investigate Mrc1’s potential role in fork pausing at
tRNAs. Finally, we address the lethality of mrc1  rrm3 cells, which was previously
attributed to a requirement for an Mrc1-dependent checkpoint response in rrm3 cells
(Torres et al, 2004).
When replication forks stall at DNA lesions created by MMS or UV, the DNA
damage checkpoint is triggered and channels a signal from Mec1 through Rad9.
Similar to Mrc1, Mec1 dependent phosphorylation of Rad9 occurs on several specific
S/TQ residues, including six located within a cluster, termed the [S/T]Q cluster
domain (SCD) (Emili, 1998; Schwartz et al, 2002; Vialard et al, 1998). Mutating a
single residue within this cluster has little effect on checkpoint function of Rad9,
which has probably evolved so that the cell could tolerate a mutation to the Rad9
protein without significantly compromising cellular viability. However, when all
seven residues are mutated to alanine, Rad9 is only partially phosphorylated by Mec1,
leading to weak binding of Rad9 to Rad53 and poor activation of Rad53. These cells
15

are almost as sensitive to MMS as rad9  cells, which underscores the importance of
Rad9 SCD phosphorylation for proper checkpoint activation (Schwartz et al, 2002).

Rad53 channels upstream checkpoint signals to protect cellular viability

The effector kinase, Rad53, is the “work-horse” of the intra-S phase
checkpoints. Rad53 is activated (hyperphosphorylated) during S phase in response to
nucleotide depletion and damaged DNA (Sun et al, 1996; Tercero et al, 2003). In its
activated form, Rad53 upregulates transcription of DNA repair genes, inhibits late
origin firing, stabilizes the replication fork, and inhibits cell cycle progression (Allen
et al, 1994; Lopes et al, 2001; Santocanale & Diffley, 1998; Shirahige et al, 1998;
Sogo et al, 2002; Tercero & Diffley, 2001; Weinert et al, 1994).
The Rad53 protein consists of three major regions: a kinase domain and two
flanking forkhead associated (FHA1 and 2) homology domains (Hofmann & Bucher,
1995) FHA domains have been evolutionarily conserved from prokaryotes to
eukaryotes and facilitate a wide variety of phospho-regulated protein-protein
interactions within the cell that are generally associated with cell cycle control and the
DNA damage response. Rad53 is unique within the Cds1/Chk2 family of checkpoint
kinases because it contains two FHA domains, not one. Based on sequence alignment,
the FHA domain present in Cds1/Chk2 family members more closely resembles
Rad53’s FHA1 domain, rather than its FHA2 domain (Hofmann & Beach, 1994).
To determine the exact role that FHA domains play in Rad53 function,
previous studies have created point mutations in the FHA domains of Rad53: rad53-
16

R70A (FHA1), rad53-R605A (FHA2) and rad53-NVS655-657AAA (FHA2) (Pike et al,
2004; Pike et al, 2003; Schwartz et al, 2003). Mutations to the FHA2 domain render
cells moderately sensitive to MMS insult; however mutations in the FHA1 domain do
not significantly affect cellular viability when exposed to HU or MMS (Pike et al,
2003; Schwartz et al, 2003). Interestingly, cells carrying FHA1 and FHA2 mutations
are more sensitive than either single mutant to HU and MMS, which suggests that the
two domains are somewhat functionally redundant (Pike et al, 2003). Also, rad53-
R70Arad9 , rad53-R70Arad17  and rad53-R70Addc1  cells are extremely sensitive
to DNA damaging agents, which further supports to the hypothesis that FHA1 can act
in the DNA damage response. The rad53-R70A allele cannot restrain late origin firing
in HU, which is the same phenotype that an mrc1  cell displays and suggests that
FHA1’s primary role is within the replication stress checkpoint; however has
overlapping function that can act in the DNA damage checkpoint (Pike et al, 2004).
Although evidence for an interaction between the FHA1 domain of Rad53 and Mrc1 is
lacking in budding yeast, reports in fission yeast demonstrate that Mrc1 binds to the
FHA domain of Cds1 in yeast two-hybrid analysis. Importantly, these studies
demonstrate that Rad3 (scMec1) activates Mrc1 on Thr645 in response to HU stress,
which is necessary for full activation of Cds1 (Tanaka & Russell, 2004; Zhao &
Russell, 2004). Experiments aimed at testing the FHA1-Mrc1 interaction in budding
yeast with wild-type Mrc1 and checkpoint defective Mrc1 (mrc1-AQ) will help to
further elucidate Mrc1’s role at the replication fork during a checkpoint response.
Located between its two FHA domains, Rad53’s robust checkpoint activity is
due its kinase domain (Pellicioli et al, 1999; Stern et al, 1991). During a DNA damage
17

response, Rad53 is phosphorylated by Mec1 and Rad9 on more than 20 residues
(Smolka et al, 2005; Sweeney et al, 2005). Despite a wealth of knowledge regarding
Rad53 interactions within the cell, an exact mechanism leading to checkpoint
activation is lacking. One hypothesis suggests that Rad9 acts as a docking site for
Rad53, thus facilitating Mec1 phosphorylation. Once the initial phosphorylation
occurs, Rad53 is released, which allows docking and activation of another Rad53
protein. Phosphorylated and released Rad53 can then autophosphorylate another
molecule of Rad53, leading to the amplification of the checkpoint signal and
maintenance of genome integrity (Sweeney et al, 2005).
Possibly the most important role of Rad53 during a checkpoint response is fork
stabilization. In a checkpoint dependent manner, Rad53 helps to maintain the
association of replisome components. The essential functions of Rad53 during a
checkpoint response are dependent on its kinase activity, which can be genetically
separated from upstream Mec1 phosphorylation using a dominant-negative kinase-
dead allele of Rad53, rad53-D339A. Interestingly, overexpression of rad53-D339A
after a normal checkpoint response expedites bulk DNA synthesis in the presence of
MMS (Paulovich & Hartwell, 1995; Pellicioli et al, 1999; Tercero & Diffley, 2001).
Pellicioli and colleagues demonstrated that Rad53 kinase activity controls
phosphorylation of the DNA pol-  primase complex, suggesting that Rad53 could
control replication fork progression/restart. Another study, which used a hypomorph
of Rad53 (rad53HA), supported these claims by demonstrating that decreased kinase
activity of Rad53 correlates with expedited bulk DNA synthesis in the presence of
MMS (Cordon-Preciado et al, 2006). The idea of checkpoint regulated fork
18

progression, which was first proposed by Painter and Young (1980), was challenged
was Tercero & Diffley (2001) when they showed that replication fork movement
along Chromosome VI was indistinguishable between WT and rad53  or mec1 cells
exposed to MMS. However, at least one important caveat exists within Tercero &
Diffley’s experiments; though their experiment was the most detailed to date, they
examined replication fork progression/restart in MMS without an active checkpoint.
In response to nucleotide depletion or DNA damage, the checkpoint is essential for
viability, presumably because of its effect on replication fork stability (Lopes et al,
2001; Sogo et al, 2002; Tercero & Diffley, 2001). rad53  cells exposed to genotoxic
agents display decreased maintenance of polymerases and Mcms at the replication
fork (Cobb et al, 2005; Lucca et al, 2004). Indeed, Cds1 (scRad53)
coimmunoprecipitates with and phosphorylates Mcm4 in response to nucleotide
depletion (Bailis et al, 2008). As a result of replisome dissociation, rad53  mutants
exhibit increased stretches of ssDNA facilitating fork reversal, recombination,
formation of pathological structures and genomic rearrangements.

The DDK-Rad53 interaction facilitates inhibits late origin firing and facilitates fork
stability

The Cdc7-Dbf4 kinase (DDK complex) plays an essential role in the initiation
of DNA replication and is normally chromatin associated, likely at origins (Weinreich
& Stillman, 1999). During an HU-induced checkpoint response, Rad53 restrains late
origin firing. Potentially this could occur through a Rad53-dependent Dbf4
19

phosphorylation event, which displaces Dbf4 from chromatin. In support of this,
overexpression of the N-terminus of Dbf4 is sufficient to derepress late origin firing in
HU. Likely, the overexpressed Dbf4 N terminus “titrates” out active Rad53, which
inhibits Rad53 from displacing chromatin bound Dbf4-Cdc7 and leads to late origin
firing (Duncker et al, 2002; Weinreich & Stillman, 1999).
Accumulating evidence suggests that DDK is also important for replication
fork stability. Mutations to Dfp1 (ScDbf4) confer sensitivity to MMS despite
relatively little effect to checkpoint activation (Fung & Brown, 2002). DDK appears
to be in a unique position to affect fork stability as some of its substrates include:
Cdc45, pol-  primase, and Mcm2 (Lei et al, 1997; Nougarede et al, 2000; Weinreich
& Stillman, 1999). In addition, Cdc7 phosphorylates Claspin (Mrc1 homolog) in
mouse embroyonic cells exposed to HU (Kim et al, 2008). Furthermore, Hsk1
associates with Swi1 (scTof1), which is part of the fork protection complex
(Matsumoto et al, 2005). At the present time it is unclear whether DDK travels with
replication forks. However, it is tempting to speculate that DDK plays a major role in
the maintenance of genome stability. Future experiments designed to detect more
substrates of DDK and its presence (or absence) at replication forks will undoubtedly
provide valuable information to researchers trying to understand how replication forks
are stabilized by the S phase checkpoints.

Checkpoint deactivation allows resumption of the cell cycle
A great deal of work has been done to elucidate the players and pathways that
activate Rad53; less clear is how Rad53 is deactivated. Cells that fail to deactivate the
20

checkpoint often remain arrested in the cell cycle and die (reviewed in Harrison &
Haber, 2006). Recent evidence has shown that in response to an irreparable DSB, the
cell does not require new protein synthesis, nor does it degrade Rad53. Instead, the
cell employs two phosphatases, Ptc2 and Ptc3, which are responsible for the
dephosphorylation of Rad53 (Leroy et al, 2003). In Chapter 3, we investigate the role
of the Psy2-Pph3 phosphatase complex and its role in checkpoint deactivation after an
MMS-induced DNA damage response.
As mentioned above, Tercero and Diffley made their conclusions on
checkpoint independent fork progression in cells that lacked an active checkpoint and
die once they enter S phase in the presence of MMS. As a result, their replication
forks were likely undergoing a number of genotoxic events, clouding accurate
analysis. In chapter 4, we examine replication fork restart kinetics under various
conditions of Rad53 activity. We also investigate an alternative pathway to deactivate
Rad53 after MMS insult, mediated by Ptc2 and the effect of its absence on fork restart
and cellular viability in a pph3 cell. Taken together, these data help us to better
understand the actions of Rad53 within the cell and provide further insight into the
intricacy of the checkpoint and its overall effect on the cell.

DNA repair during S phase occurs via error prone or error free pathways
In addition to contending with fork impediments, the cell must also maintain
fidelity of the genome. To accomplish this, the cell employs damage repair pathways.
21

In G
1
, nucleotide excision repair (NER) processes lesions present in the DNA due to
UV irradiation, whereas base excision repair (BER) processes lesions due to chemical
treatment (i.e. MMS alkylation). In both NER and BER, lesions are removed from the
DNA. Lesions that persist into or arise during S phase can block the replication fork
and are processed by post replication repair (PRR). In contrast to NER and BER, PRR
bypasses lesions instead of removing them. This is accomplished by an error free
mechanism (template switch, TS) or an error prone mechanism (translesion synthesis).
The exact mechanism leading to pathway choice has not been completely worked out;
however recent findings have elucidated how cells tolerate lesions in the DNA during
S phase.
PCNA is an essential processivity factor involved in both DNA replication and
DNA repair. PCNA can be modified posttranslationally via sumoylation and
ubiquitination (reviewed in Watts, 2006). Sumoylation occurs at K164 and K127,
which suppresses recombination at replication forks through the recruitment of Srs2,
via disruption of Rad51 filaments present on ssDNA (Hoege et al, 2002; Papouli et al,
2005). PCNA can also be ubiquitinated via the Rad6/18 complex, which results in
either error free or error prone PRR. Initially, the Rad6/18 complex is recruited to
stalled replication forks because of Rad18’s affinity to stretches of ssDNA (Fig. 2)
(Bailly et al, 1997). Monoubiquitination by Rad6/18 leads to a polymerase switch for
a less processive and less accurate polymerase. After incorporation of a few bases, a
second polymerase switch occurs, placing the normal replicative polymerase back on
the DNA, resulting in bypass of the offending lesion (i.e. error prone PRR). Further
ubiquitination by Ubc13/Mms2 and Rad5 leads to the error free pathway (Zhang &
22

Lawrence, 2005). Interestingly, Rad5 displays helicase activity and is capable of
regressing stalled replication forks, providing a damage free template (via the newly
copied sister strand) to bypass the lesion (Blastyak et al, 2007).
In addition to the two pathways described above, the DNA damage checkpoint
appears to play a role in PRR. Cells lacking Rad6/18 are extremely sensitive to DNA
damage. Cells deficient for error prone (rev1 , rev7, or rev3   and error free
(rad5 , ubc13, or mms2 ) PRR pathways are not as sensitive as rad6  or rad18 
mutants, suggesting another branch in DNA repair (Barbour et al, 2006). Indeed, when
members of the DNA damage checkpoint (rad9 , rad17 , rad24 ) are deleted in
combination with error prone and error free mutants, sensitivity to MMS approaches
levels displayed in rad6  and rad18  mutants.
A recent study has demonstrated that S. pombe Rad9 (9-1-1 complex) plays a
role in determining PRR pathway choice, which is mediated by Rad3 (scMec1)
phosphorylation (Kai et al, 2007). Briefly, when the checkpoint clamp is
phosphorylated, Rad9 (scDdc1) associates with Mms2, and promotes error free
damage repair. Interestingly, this phosphorylation event further promotes sumoylation
of PCNA, recruiting Srs2 to stalled forks, thus avoiding recombination. Despite the
implications of these results and the direct link between DNA damage signaling and
PRR, it still remains to be seen how the cell might choose error prone PRR. Due to its
hypomutagenic phenotype and synergistic MMS sensitivity with rad5  mutants, Cdc7
has been implicated in the error prone branch of PRR (Pessoa-Brandao & Sclafani,
2004). Possibly, the default pathway within the cell is error free. However, in a
situation where there is an overwhelming amount of DNA damage to the template,
23

causing both leading and lagging strands to halt (therefore providing no template for
fork regression), the cell could use error prone PRR to bypass lesions. Cdc7 appears
to be an attractive candidate as it can phosphorylate pol -primase, potentially leading
to repriming downstream of the offending lesion. Experiments aimed at dissecting
PRR pathway choice and elucidating the players and their roles in maintaining a stable
replisome will undoubtedly add to our understanding of replication fork dynamics.

24

Figure 2. Error prone and error free repair pathways in budding yeast. Upon replication fork
stalling, PCNA is monoubiquitinated signaling error prone DNA repair or polyubiquitinated to signal
Rad5-dependent fork regression and error free DNA repair. In parallel to DNA repair, PCNA is
SUMOylated, which recruits Srs2 and inhibits unscheduled recombination events.

25

CHAPTER 2

Mrc1 is Required for Normal Progression of Replication Forks Throughout
Chromatin in Saccharomyces cerevisiae

All of the contents of this chapter have been published and can be found in
Molecular Cell (2005, Vol 19, pp691-697).

Figure Contributions

I contributed the data in Figures 3,5,6,8, and 9.
Christopher J. Viggiani contributed the data in Figures 4 and 7.

Overview

The accurate duplication of chromosomal DNA presents a significant
challenge to the cell. Replication forks must contend with the presence and activities
of various chromatin structures, transcription complexes and recombination
machinery, which often cause replication forks to pause (reviewed in Rothstein et al,
2000). For example, replication forks pause at tRNA genes when the direction of
transcription is opposite that of the replication fork (Deshpande & Newlon, 1996).
Although the exact function of fork pausing at tRNA genes is not known, it may help
prevent fork dysfunction that might result from collision with the transcription
machinery, and thus, facilitate resumption of normal fork progression.
Multiple mechanisms contribute to replication fork stability and faithful
genome duplication. For example, the Rrm3 DNA helicase has been implicated as
playing an important role in the progression of replication forks through natural
impediments such as tRNA genes and chromatin complexes (Ivessa et al, 2003; Ivessa
et al, 2002; Ivessa et al, 2000). In addition, surveillance mechanisms called
26

checkpoints detect and respond to problems that arise during DNA replication. A
critically important pathway that operates during S phase is the replication stress
checkpoint (reviewed in Osborn et al, 2002). This pathway acts by maintaining
adequate deoxyribonucleotide (dNTP) levels, increasing transcription of DNA repair
genes, slowing cell cycle progression, and maintaining stable replication forks that are
poised to resume replication following perturbations.
Mediator of replication checkpoint protein 1 (Mrc1) localizes with replication
forks and acts in the replication stress response by transducing signals of stress from
the “sensor” kinase Mec1 to the “effector” kinase Rad53 (Alcasabas et al, 2001; Katou
et al, 2003; Osborn & Elledge, 2003; Tanaka & Russell, 2001). Thus, in cells
challenged with an inhibitor of DNA synthesis such as hydroxyurea (HU), which
depletes deoxyribonucleotide levels, Mrc1 is required for effective activation of Rad53
by Mec1. Remarkably, in mrc1 cells undergoing DNA replication in the presence of
HU, the progression of DNA synthesis halts, while replication fork proteins, including
Mcm proteins, Cdc45 and DNA polymerases, appear to progress farther along the
chromatin, suggesting that uncoupling of the replication apparatus from the site of
DNA synthesis has occurred (Katou et al, 2003). This has led to the conclusion that
Mrc1 functions in a pausing complex that maintains replisome integrity when
exogenous stresses are encountered.
Mrc1 also functions during normal DNA synthesis in cells that have not
received exogenous perturbation. This is evidenced by the somewhat slower rate of
bulk chromosomal DNA synthesis in mrc1 cells, which is accompanied by a Rad9-
dependent DNA damage response (Alcasabas et al, 2001). The reason for the slower
27

replication of mrc1  cells remains unclear, but does not depend on checkpoint
signaling by either Rad9 or Mrc1 (Alcasabas et al, 2001; Osborn & Elledge, 2003). It
is also unknown whether the defective replisome pausing in HU reflects loss of the
replication or checkpoint-signaling function of Mrc1. In this study we have
investigated the role of Mrc1 in DNA replication of unperturbed cells, by examining
origin initiation, fork elongation, and fork pausing in cells lacking Mrc1 function. We
also have explored the possible role of Mrc1 checkpoint signaling in response to fork
pausing by analyzing genetic interactions with Rrm3. Together, our findings
demonstrate that Mrc1 plays a central and constitutive role in the efficient function of
the replisome. Our results also suggest that Rrm3 is vital to repair DNA damage
created by defective replication forks in mrc1cells.

Results and Discussion

Mrc1 is required for a normal rate of replication fork progression
The replication defect of cells lacking Mrc1 could be due to a defect in the
initiation of replication, in elongation, or in replisome pausing. To distinguish
between these possibilities, we monitored origin initiation and the progression of
replication forks using two-dimensional gel electrophoresis (2D gel) across regions of
chromosomes III and VI. In each of these regions, one or more active replication
origins has been deleted, so that in wild-type cells the replication of these regions is
unidirectional, facilitating analysis of fork progression (Fig. 3A and Fig. 5A) (Tercero
& Diffley, 2001).
28

In wild-type cells released synchronously into S phase, initiation of ARS607
occurred at about 30 minutes. By 36 minutes, primarily larger replication structures
were present due to fork elongation (Fig. 3B). In mrc1 cells, initiation of ARS607
occurred with timing like wild-type cells, as bubbles were first observed at 30 minutes
(Fig. 3B). The shapes of the bubble arcs and their greater intensities suggested a delay
in the elongation of initiation bubbles in mrc1  cells. At 30 minutes, small bubbles
predominated (Fig. 3B), and at 36 minutes, small and large bubbles were present, as
well as large forks (Fig. 3B). These larger structures persisted near ARS607 at 48 to
60 minutes. The presence of small bubbles at 30-36 minutes together with their
absence at later times, as in wild-type cells, indicates that initiation timing of these
origins is not altered by the absence of Mrc1 (also, see below). Furthermore, the
efficiency of ARS607 is not altered by loss of Mrc1 based on the virtual absence of
smaller fork structures in these gels and on the analysis of unsynchronized cultures
(data not shown). These results suggest that replication forks emanating from
ARS607 elongated more slowly in mrc1  cells than in wild-type cells.
29

Figure 3. Mrc1 is required for normal replication fork progression along chromosome VI. (A)
Position of probes (thin bars beneath the chromosome depiction) and relevant ClaI (C) and EcoRI (E)
restriction sites used in the 2D gel analysis of modified chromosome VI are shown approximately to
scale. Thick bars indicate the positions of sequences analyzed for BrdU incorporation and Cdc45
association shown in Figures 4 and 7. The bent arrows indicate the positions and transcriptional
orientations of the tRNA genes tA(AGC)F in region A and SUP6 in region B. (B) Wild-type (SSy48)
and mrc1  (SSy47) cells were blocked in G
1
with -factor at 23°C and released synchronously into S
phase at 20°C. Cells were collected at the indicated times for 2D gel analysis by digestion with EcoRI
and ClaI. Blots were probed for ARS607, and stripped and re-probed for regions A, B, and C. Filled
and unfilled arrowheads indicate small and large bubble structures, respectively; open and double
arrowheads indicate large and small fork structures, respectively; the complete arrows indicate pause
sites. The greater signal intensity of replication structures in mrc1  cells is reproducible and appears to
reflect the longer period of time that forks are present within each analyzed fragment.
30

Next, we analyzed the progression of replication forks from ARS607 into
adjacent chromosome VI sequences, restriction fragments “A”, “B”, and “C”, whose
midpoints lie at 4, 12, and 20 kb from ARS607 (Fig. 3A). In addition, regions A and
B each contain a tRNA gene that permitted analysis of replication fork pausing
(discussed below). In wild-type cells, the peak signals for forks at A, B, and C
occurred at 36, 48, and 60 minutes, respectively (Fig. 3B). Based on the initial
appearance of replication structures at ARS607 at 30 minutes and at region C at 48
minutes, the data indicate an average fork rate of ~1 kb per minute in wild-type cells.
In the absence of Mrc1, the progression of replication forks into and throughout the
adjacent chromosomal regions was significantly delayed. The peak signals for forks at
regions A, B, and C occurred at about 48, 72, and 84 minutes, and forks were present
within each region for a longer period, consistent with their slower movement through
each region (Fig. 3B). Furthermore, the preponderance of smaller forks earlier and
larger forks later in the interval during which forks move through each region is
consistent with slow movement of replication forks throughout the region (Fig. 3B).
Based on the timing of initial appearance of replication structures at ARS607 and at
region C, we estimate that replication forks progress at about half the wild-type rate in
mrc1 cells.
Replication forks also elongated at about half the wild-type rate across a 23 kb
region of chromosome III in cells lacking Mrc1 (Fig. 5B). Together with the delayed
completion of bulk DNA replication in mrc1 cells (Fig. 4A) (Alcasabas et al, 2001),
these findings suggest a general requirement for Mrc1 in the normal progression of
replication forks throughout chromatin. Whether replication forks in mrc1 cells
31

progress at a uniformly slow rate or whether the slow progression is due to occasional
stalling at random sites (requiring restart) is not clear. However, the perdurance of
some replication forks at later times (e.g.: Fig. 3B, ARS607 at 72-96 minutes),
suggests that some forks have stalled, but remain stable.
32

Figure 4. Mrc1 is required for normal progression of DNA synthesis. Wild-type (CVy39), mrc1-
AQ (CVy40) and mrc1  (CVy29) cells, which all express GPD-TK, were blocked in G
1
with -factor at
23°C and released synchronously into S phase at 20°C. (A) DNA content analysis. (B) Cells were
collected at the indicated times for analysis of BrdU incorporation. Immunoprecipitated DNA
sequences at ARS607 and at distances of 5, 15, and 20 kb (see Fig. 3A) were detected by PCR
amplification. (C) PCR products in (B) and of input DNA (not shown) were quantified and plotted as
the ratio of immunoprecipitated DNA/input DNA (% BrdU Incorporated).
33

Figure 4.

34

As part of our analysis of chromosome III, we analyzed replication of
ARS304, an inefficient origin at a distance of 45 kb from ARS306. Unexpectedly, we
observed increased initiation of ARS304 in mrc1  cells (Fig. 5B). Because the level
of ARS304 initiation in mrc1-AQ cells was similar to wild-type (Fig. 6), we conclude
that the increased initiation frequency of ARS304 in mrc1 cells is due primarily to
the slower progression of replication forks from ARS306, which permits more time for
ARS304 initiation before it is passively replicated by a fork from ARS306. We also
have observed evidence for increased initiation of the inefficient origin near the VI-R
telomere in mrc1  cells (data not shown). The increased initiation of some origins
due to their delayed passive replication helps explain why genome duplication in
mrc1 cells does not appear to require twice as long as wild-type cells (Fig. 4A),
although our data indicate that forks progress at about half the wild-type rate.
35

Figure 5. Mrc1 is required for normal replication fork progression along chromosome III. (A)
Position of probes (thin bars beneath the chromosome depiction) and relevant ClaI (C) and EcoRI (E)
restriction sites used in the 2D gel analysis of modified chromosome III are shown approximately to
scale. (B) Blots from the experiments in Figure 3 were stripped and re-probed for ARS306, Region D,
and ARS304. See Figure 4A for DNA content analysis.

We also analyzed initiation and fork progression in a strain harboring the
mrc1-AQ allele, in which all hypothetical Mec1 phosphorylation sites have been
mutated (Osborn & Elledge, 2003). This allele is defective in checkpoint signaling to
activate Rad53, but appears to retain its function in DNA replication, based on its
36

approximately normal rate of replication as measured by DNA content analysis (Fig.
4A) (Osborn & Elledge, 2003). 2D gel analysis indicated that initiation and fork
movement in the mrc1-AQ strain was similar to wild-type cells (Fig. 6). Thus,
replication forks in mrc1-AQ cells progress with similar kinetics as in wild-type cells
and the slower replication fork movement of mrc1  cells is not due to loss of Mrc1’s
checkpoint function.

Figure 6. Mrc1-AQ supports normal fork progression. mrc1-AQ (SSy116) cells were analyzed as
in Figure 3B and Figure 5B.

To confirm our conclusion that normal fork progression depends on Mrc1, but
not its checkpoint function, we analyzed nascent DNA synthesis by measuring
incorporation of the thymidine analog bromodeoxyuridine (BrdU) along chromosome
37

VI (Fig. 4B and C). In wild-type, mrc1-AQ and mrc1 cells, BrdU incorporation
occurred with similar timing and efficiency at the origin ARS607, primarily between
24 and 36 minutes, reinforcing our conclusion that initiation is not affected by loss of
Mrc1 function. The kinetics of BrdU incorporation into DNA sequences at distances
of 5, 15, and 20 kb from ARS607 was indistinguishable between wild-type and mrc1-
AQ cells. However, BrdU incorporation into DNA sequences at distances of 5, 15,
and 20 kb from ARS607 was delayed in mrc1  cells. The degree of delay is
consistent with the results of the 2D gel analysis, with a delay of about 24 minutes at a
distance of 20kb. Thus, in the absence of Mrc1, the replication timings of DNA
sequences proximal to ARS607 were delayed in relation to their distance from the
origin, with more distal sequences experiencing a greater delay. These data are fully
consistent with the conclusion that replication forks progress more slowly in cells
lacking Mrc1 function, whereas the checkpoint signaling function of Mrc1 appears to
be dispensable for normal fork progression in unperturbed cells.

Progression of Cdc45 is slower in the absence of Mrc1
The possibility that Mrc1 plays a role in coupling replication fork components
to the site of DNA synthesis prompted us to examine whether the progression of
Cdc45 along chromatin also was slowed in the absence of Mrc1 or whether it might
move rapidly ahead of the slow-moving replication forks. To examine the movement
of Cdc45, we analyzed its association with chromosome VI by chromatin
immunoprecipitation (ChIP) (Fig. 7). This analysis was performed at 16°C to enhance
the resolution of the assay. In both wild type and mrc1 cells, Cdc45 association with
38

ARS607 peaked at 60 minutes and diminished soon after. The similar timings in wild-
type and mrc1  cells reinforce our conclusion that initiation timing of ARS607 is not
altered by loss of Mrc1 function. However, the progression of Cdc45 along chromatin
was altered in the absence of Mrc1. In wild-type cells, Cdc45 moved rapidly,
associating maximally with sequences 5, 15 and 20 kb distant from ARS607 at 72, 72,
and 84 minutes, respectively. In mrc1 cells, Cdc45 progressed more slowly,
associating maximally with the sequences 5, 15 and 20 kb from ARS607 at 72, 108
and 120 minutes, respectively. Hence, the progression of Cdc45 along chromatin
during S phase, like that of replication forks, is slowed in the absence of Mrc1.
Although the slowed progression of Cdc45 may be consistent with Cdc45 remaining
coupled with the site of DNA synthesis in unperturbed mrc1  cells, it is possible that
dissociation of Cdc45 from the site of DNA synthesis does occur, but can be resolved
by our analysis only when DNA synthesis is inhibited strongly with HU. Whether or
not a defect in coupling exists, the results make clear that Mrc1 is required for the
normal progression of Cdc45 along chromatin during DNA replication.
39

Figure 7. Slower progression of Cdc45 in mrc1  cells. Wild-type (CVy31) and mrc1  (CVy30)
cells, which both express Cdc45-HA, were blocked in G
1
with -factor at 23°C and released
synchronously into S phase at 16°C. Cells were collected at the indicated times for ChIP analysis with
anti-HA antibody. DNA sequences at ARS607 and at distances of 5, 15, and 20 kb (see Fig. 3A) were
quantified by PCR amplification. The ratio of immunoprecipitated to input DNA is plotted as “%
Cdc45 Bound”.

Resolution of naturally paused replication forks does not require Mrc1.
Next, we examined whether replication forks lacking Mrc1 were defective in
progression through endogenous pause sites, which might contribute to the slower
progression of replication forks that we have observed in mrc1  cells. We analyzed
pausing at two tRNA genes within regions A and B adjacent to ARS607 (Fig. 3A).
Fork pausing at each tRNA gene was apparent as a dark spot within each fork arc.
40

Compared to wild-type cells, the pause signals appeared reduced in mrc1  cells (Fig.
3B, arrows). However, we suspected that the decreased pause signal resulted at least
in part from the slower fork movement of mrc1  cells, which increased the length of
time spent by replication forks throughout regions A and B, and thus decrease the
relative intensity of the pause signal. Consistent with this idea, careful quantification
of the pause at the tRNA gene in region A relative to total replication structures
indicated that pausing was reduced by about half (Fig. 8). Because replication forks
progress at about half the wild-type rate, these data suggest that the duration of
pausing at this tRNA gene is not significantly altered in the absence of Mrc1. In
addition, these results strongly suggest that Mrc1 is not required for resolution of
paused forks, which would appear as an increased pause signal.
41

Figure 8. Loss of Mrc1 function does not enhance fork pausing at tRNA genes. (A) Position of
probe (thin bars beneath the chromosome depiction) and relevant SpeI (S) restriction sites used in the
2D gel analysis of tA(AGC)F are shown approximately to scale. (B) Unsynchronized wild-type
(SSy48), mrc1  (SSy47), and mrc1-AQ (SSy116) cells grown at 30°C were harvested for 2D gel
analysis with SpeI digestion. Blots were probed for tA(AGC)F. The arrows indicate the pause signal.
(C) Quantification is described in Figure 9C legend. Error bars denote standard deviation.

It was possible that paused forks in mrc1  cells were unstable, and hence,
interfered with our ability to detect the paused structure itself and/or a defect in its
resolution. To address directly the possibility that defective pausing somehow
contributes to fork dysfunction in mrc1  cells, we analyzed fork progression along the
20 kb chromosome VI region in mrc1  cells in which we deleted both tRNA genes.
42

Deletion of the tRNA genes did not significantly affect the rate of fork progression
across this region (data not shown), indicating that the slow fork movement of mrc1 
cells is not due to defective pausing or instability of forks encountering these sites. In
support of this conclusion, forks appeared to progress slowly within all the regions
that we analyzed, including the chromosome III region, which does not contain
elements known to be associated with fork pausing (Fig. 5). We also note that mrc1
cells are not highly sensitive to HU, suggesting these cells are able to restart
replication effectively (Alcasabas et al, 2001). We conclude that Mrc1 is not required
for the stability and restart of naturally stalled forks.

The replication defect of mrc1  cells creates a dependence on Rrm3 for viability
The Rrm3 DNA helicase enables the resolution of stalled replication forks. In
the absence of Rrm3, fork pausing at natural sites is prolonged and cells become
dependent on MRC1 for viability (Ivessa et al, 2003; Ivessa et al, 2002; Ooi et al,
2003; Tong et al, 2004; Torres et al, 2004). These findings have suggested that the
repair or re-start of stalled forks in rrm3 cells is dependent on the replication stress
response, which is mediated by Mrc1 (Torres et al, 2004). To determine whether the
requirement of rrm3  cells for MRC1 reflects the role of Mrc1 in checkpoint signaling
or in fork progression; we assessed the viability of rrm3 mrc1-AQ cells by dissection
of a diploid strain heterozygous for both alleles. Haploid rrm3 mrc1-AQ cells
emerged at a frequency equivalent to either single mutant strain, and grew at a similar
rate (Fig. 9A). Thus, Mrc1’s checkpoint function is dispensable in cells lacking Rrm3.
43

As expected, rrm3  mrc1  cells were inviable (Fig. 9B). These results suggest that
the lethality of rrm3  mrc1  cells reflects a dependence on Rrm3 to resolve
replication fork defects incurred by loss of Mrc1 function, rather than a vital
replication checkpoint response to prolonged fork stalling in rrm3 cells.

44

Figure 9. Rrm3 is required for viability of mrc1  cells but not mrc1-AQ cells. Diploid strains
SSy108 and SSy109, in (A) and (B), respectively, were induced to sporulate at 23°C; germination was
at 30°C. The relevant genotypes are indicated in the panels below as follows: WT= wild-type, AQ=
mrc1-AQ, m1= mrc1 , r3: rrm3 . The genotypes of inviable spores were inferred assuming 2:2
segregation of genetic loci within each tetrad. (C) Haploid strains W303-1a (WT), SSy72 (mrc1-AQ),
SSy69 (rrm3 ), and SSy108 (mrc1-AQ rrm3 ) were subjected to 2D gel analysis of tA(AGC)F by
digestion with SpeI (see Fig. 8A for position of probe and restriction sites). (D) Three sets of blots like
those in (C) were quantified by measuring the pause signal and dividing by the signal for all replication
intermediates along the fork arc (excluding the 1N spot). The data was plotted relative to wild-type,
which was arbitrarily assigned the value 1. Error bars denote standard deviation.
45

Figure 9.

46

To examine further the function of the replication checkpoint in response to
fork stalling, we analyzed fork pausing at tRNA genes in wild-type and rrm3 cells
with either an intact or defective replication checkpoint. Pausing was not altered in
mrc1-AQ cells, indicating that the replication checkpoint is not required for the
resolution of paused forks (Fig. 9C, data not shown for SUP6). As expected, rrm3
cells exhibited greatly increased stalling at the tRNA gene (Fig. 9C and D).
Furthermore, we observed a similar level of fork pausing in rrm3 mrc1-AQ cells as
in rrm3  cells (Fig. 9C and D). Thus, checkpoint signaling by Mrc1 does not appear
to participate in replication fork pausing or resolution, either in the presence or
absence of Rrm3. Because mrc1 cells are defective in fork progression throughout
chromatin and require Rrm3 for viability, we propose that Rrm3 acts throughout
chromatin in response to replication fork defects and impediments.

A central role for Mrc1 at replication forks?
Previous analysis of mrc1  cells treated with HU has led to the conclusion that
Mrc1 is required for normal fork pausing in response to replication stress by
maintaining coupling of replication proteins to the site where DNA synthesis is
inhibited (Katou et al, 2003). Our demonstration that lack of Mrc1’s replication
function, but not its checkpoint function, causes defective fork progression throughout
chromatin in untreated cells is consistent with a coupling role for Mrc1 in the
replisome, and further suggests this role is constitutive, rather than checkpoint
induced. Mrc1 co-immunoprecipitates with Cdc45, Mcm2, and Mcm3 (Katou et al,
2003; Nedelcheva et al, 2005). Furthermore, recent biochemical analyses of S. pombe
47

Mrc1 and its apparent human homolog, Claspin, indicate that Mrc1/Claspin can bind
branched DNA structures in vitro (Sar et al, 2004; Zhao & Russell, 2004). Together,
these data suggest that Mrc1 links Cdc45 and/or Mcm proteins to replication forks in
vivo. Depletion of Claspin from Xenopus replication extracts also modestly reduced
the rate of DNA synthesis suggesting that Claspin may play a similar role at
replication forks in vertebrates (Lee et al, 2003).
The slower rate of fork progression in cells lacking Mrc1 may reflect
uncoupling of replication factors. Studies of bacterial and viral eukaryotic DNA
replication have demonstrated synergistic stimulation of helicase and polymerase
activities that depend upon their physical association (Dong et al, 1996; Kim et al,
1996; Yuzhakov et al, 1996). If loss of Mrc1 function disrupts similar interactions in
the eukaryotic replisome, slower fork progression might result. In addition to
compromising the efficacy of DNA synthesis at the replication fork, loss of Mrc1 from
the site of DNA synthesis may expose normal replication structures to DNA damage
sensors, DNA repair proteins, and recombination factors. This could explain some of
the defects associated with loss of Mrc1, such as activation of the DNA damage
response and increased levels of homologous recombination (Alcasabas et al, 2001;
Xu et al, 2004). These phenotypes may reflect the presence of excess unwound DNA,
or DNA structures that may serve as effective recombination substrates such as
double-strand breaks, or 3’ ends. The presence of such structures in mrc1  cells is
suggested by their vital dependence on the Srs2 and Rrm3 DNA helicases (Ooi et al,
2003; Tong et al, 2004; Torres et al, 2004; Xu et al, 2004), which appear to function in
the repair of stalled or damaged replication intermediates, perhaps by preventing their
48

subversion into recombination pathways. The elucidation of a general role of Mrc1 in
the function of replication forks provides insight into the mechanisms that promote
genome stability during DNA replication.

49

CHAPTER 3

Psy2 and Pph3 Form a Phosphatase Complex Required for Rad53
Dephosphorylation and Replication Fork Restart During Recovery from DNA
Damage

All of the contents of this chapter have been published and can be found in the
Proceedings of the National Academy of Sciences (2007, Vol 104, pp 9290-9295).

Figure Contributions

Bryan M. O’Neill contributed the work found in Figures 10-14 and 18.
I contributed the work found in Figures 15-17.

Overview

Faithful maintenance of genome integrity is essential for cellular viability.
Failure to efficiently recognize and repair DNA damage can lead to genomic
instability and, in higher eukaryotes, cancer (Kolodner et al, 2002). To coordinate the
response to genotoxic stress, eukaryotic cells use intricate signaling networks, called
checkpoints, that control cell cycle progression, transcription of DNA damage
response genes, activation of DNA repair pathways, and recruitment of proteins to
sites of damage (Nyberg et al, 2002). An efficient checkpoint response is especially
important during S phase, as replication forks are particularly vulnerable to DNA
damage (Branzei & Foiani, 2005; Lambert & Carr, 2005).
The response to DNA damage in S phase is orchestrated by the intra-S cell
cycle checkpoint response, and has been particularly well characterized in
Saccharomyces cerevisiae where it is mediated by the overlapping replication
checkpoint and DNA damage checkpoint pathways (Kolodner et al, 2002). A central
50

component of both intra-S pathways is the protein kinase Rad53. Activation of Rad53
depends on the ATM and ATR-like kinases, Mec1 and Tel1, as well as a host of
adapters and damage sensors, such as the replication stress-specific proteins, Mrc1 and
Tof1; and the DNA damage-specific proteins Rad9, Mec3/Ddc1/Rad17 (PCNA-like
complex), and Rad24/Rfc2-5 (RFC-like complex) (Nyberg et al, 2002). After
activation by Mec1 and/or Tel1, Rad53 amplifies its own activity by
autophosphorylation in trans (Ma et al, 2006).
Once activated by phosphorylation in S-phase, Rad53 protects damage-stalled
replication forks from collapse and inhibits activation of late-firing origins of
replication (Branzei & Foiani, 2005; Lambert & Carr, 2005). Fork stabilization is
thought to occur by reinforcing the association of the replisome components with the
fork and inhibiting the activity of recombination enzymes at these sites. These events
are likely regulated through Rad53-mediated phosphorylation of different targets such
as RPA, Mrc1, Srs2, Mus81, and the DNA Polymerase -Primase complex (Ma et al,
2006). The mechanism of origin inhibition may include Rad53-dependent
phosphorylation of Dbf4, which inhibits the kinase activity as well as chromatin
association of the Dbf4-dependent kinase, Dbf4-Cdc7, which is required for
replication initiation (Duncker & Brown, 2003).
In contrast to its activation and its roles in the checkpoint response, relatively
little is known about deactivation of Rad53. New protein synthesis is not required for
the appearance of unphosphorylated Rad53 during recovery from checkpoint arrest
(Tercero et al, 2003), indicating that the activated kinase is dephosphorylated, not
degraded. This model is supported by the observation that Ptc2 and Ptc3, two PP2C-
51

like phosphatases, are required for Rad53 dephosphorylation following prolonged
exposure to a persistent double-strand break (Leroy et al, 2003). Recently, it was
suggested that Rad53 deactivation during prolonged recovery from a double-strand
break is dependent on H2AX (H2AX phosphorylated at Ser129) dephosphorylation
by the type 2A-like phosphatase, Pph3, in conjunction with both Psy2 and Ybl046w
(Keogh et al, 2006).
We now report that Pph3 and Psy2 form a phosphatase complex (Psy2-Pph3)
that negatively regulates Rad53 activity independent of Ybl046w. We present in vitro
and in vivo evidence suggesting that Psy2-Pph3 performs this function by directly
binding and dephosphorylating Rad53. We also find that psy2  and pph3 cells do not
efficiently resume DNA synthesis during recovery from DNA damage and attribute
this to the defective restart of stalled forks. However, DNA replication in psy2  and
pph3  cells is completed by the initiation of late-firing replication origins during
recovery from DNA damage despite the presence of hyper-phosphorylated Rad53.

Results and Discussion
Identification of the Psy2-Pph3 phosphatase complex
We originally identified PSY2 as a gene involved in tolerating and/or
regulating the cellular response to DNA damage (O'Neill et al, 2004). To better
understand the function of Psy2, we used multidimensional protein identification
technology (MudPIT) (Washburn et al, 2001) to identify proteins that associate with a
Psy2-TAP fusion protein expressed at endogenous levels from the PSY2 promoter.
Consistent with previous reports, three potential binding partners of Psy2 were
52

identified: Pph3, Ybl046w, and Spt5 (Gavin et al, 2002; Gingras et al, 2005; Ho et al,
2002). Coimmunoprecipitation experiments confirmed that Psy2 interacts with Pph3
and Ybl046w (Figure 10A). Spt5 did not immunoprecipitate with Psy2 and was not
further characterized.

Figure 10. Psy2 and Pph3 form a complex that contributes to MMS resistance. (A) Whole-cell
extracts were prepared from yeast strains expressing Psy2-TAP, Pph3-13Myc, and/or Ybl046w-13Myc
and incubated with IgG-sepharose. Whole-cell extracts (W, 10 μg of protein) and precipitated proteins
(P, 5% of precipitated fraction) were analyzed by Western blotting with the indicated antibodies. (B-D)
Survival assays of yeast containing wild-type or null alleles of PSY2, PPH3, and/or YBL046W. (B and
D) Colony forming units (cfu) were counted 3 days after deposition onto YPD-agar media containing
the indicated concentration of MMS. Strains are as listed in Table 2 and are isogenic with BY4741. (C)
Fivefold serial dilutions of wild-type (BY4741) or pph3  (FR1046) cells harboring YEplac195 or
YEplac195 carrying a wild-type or mutant allele of PPH3 were deposited on SC-URA plates containing
MMS.

We disrupted PPH3 and YBL046W to determine if these genes contribute to
MMS resistance, as observed for PSY2(O'Neill et al, 2004), and found that pph3
cells are hypersensitive to MMS (Figure 10B), but ybl046w  cells are not. The lack of
sensitivity of ybl046  cells argues that YBL046W does not function with PSY2 or
53

PPH3 in response to MMS damage (Figure 10B). The MMS sensitivity of pph3 is
complemented by wild-type PPH3 but not by pph3-Y121N, which encodes a
catalytically inactive mutant of Pph3 (Figure 10C), demonstrating that the phosphatase
activity of Pph3 is required for tolerance of MMS. We also examined the genetic
interactions between psy2 , pph3, and ybl046w  to determine whether these genes
function in the same pathways. psy2  pph cells are no more MMS sensitive than
pph3  cells (Figure 10D), indicating that PSY2 and PPH3 function in the same
pathway(s). psy2  ybl046w  cells are marginally more sensitive to MMS than psy2 
cells, consistent with the above conclusion that YBL046W does not function with PSY2
and PPH3 in the response to MMS (Figure 10D). Together, these data suggest that the
Psy2-Pph3 phosphatase complex promotes viability following MMS-induced damage,
and that this function is independent of Ybl046w.

Psy2-Pph3 antagonizes the DNA damage checkpoint response in a Rad53-
dependent manner
To study the contribution of Psy2-Pph3 to replication and/or cell cycle control,
we examined the genetic interactions of PSY2 and PPH3 with genes involved in the
DNA damage and replication checkpoints. Loss of PSY2 or PPH3 results in a
dramatic decrease in the MMS sensitivity of strains deficient for genes that function
upstream of Rad53 activation (RAD9, RAD17, RAD24, or MEC1), a synergistic
increase in the sensitivity of strains lacking genes that function downstream of Rad53
(DUN1 or PTC2), and no effect in cells lacking RAD53 itself (Figure 11 and 12).
However, it should be noted that Dun1 has Rad53-independent functions (Bashkirov
54

et al, 2003). The epistatic relationship between psy2  or pph3 and rad53  is
particularly interesting as it suggests that Psy2-Pph3 does not function outside of the
Rad53-mediated checkpoint pathway to promote viability in response to MMS
damage. In addition, Psy2-Pph3 activity appears to be specific for the DNA damage
checkpoint pathway, as cells deficient for PSY2 or PPH3 are not hypersensitive to
treatment with hydroxyurea (Figure 12C), which induces the replication checkpoint.
These data indicate that Psy2-Pph3 acts downstream of Rad53 to antagonize the
activation of the DNA damage checkpoint. Interestingly, the synergistic relationship
of psy2  and pph3  with ptc2 , suggests that Psy2-Pph3 and Ptc2 have overlapping
roles in Rad53 dephosphorylation.

55

Figure 11. PSY2 and PPH3 genetically interact with cell cycle checkpoint components. Survival
assays of yeast containing wild-type or null alleles of PSY2, PPH3, and/or YBL046W in combination
with wild-type or null alleles of cell cycle checkpoint components. Colony forming units (cfu) were
counted three days after deposition onto YPD-agar media containing the indicated concentration of
MMS. Strains are as listed in Table 2 and are isogenic with BY4741 (A and D) or RDKY3615 (B, C,
and E).

56

Figure. 12. MMS sensitivities of psy2  and pph3  cells are reduced in rad17  or rad24 
backgrounds. Colony forming units (cfu) were counted three days after deposition onto YPD-agar
media containing the indicated concentration of MMS. psy2  and pph3  cells are not hypersensitive to
HU. (C) Fivefold serial dilutions of wild-type (BY4741), psy2  (FR288), or pph3  (FR1046) cells
were deposited on YPD agar with or without 200 mM HU and grown at 30°C for 48 hours. Strains are
as listed in Table 2 and are isogenic with BY4741.

Psy2-Pph3 interacts with and directly dephosphorylates Rad53
Although not detected by MudPIT, previously published yeast two-hybrid
studies indicate that Psy2 interacts with Rad53 (Gingras et al, 2005; Uetz et al, 2000).
To confirm this interaction and determine whether it is domain-specific, we cloned
PSY2 into the bait vector pGBT9, and expressed Rad53 from the prey vector pACTIIst
as full-length protein, kinase domain, or individual Forkhead-Associated (FHA)
phosphothreonine-binding domains, FHA1 and FHA2 (Leroy et al, 2003). An
interaction was detected between Psy2 and the Rad53 kinase domain, as well as
between Psy2 and the full-length protein, but not between Psy2 and either FHA
domain (Figure 13A). The interaction could not be detected by co-
immunoprecipitation experiments from undamaged or MMS-treated cells (data not
shown). The ability to detect a Pph3-Psy2-Rad53 complex only under the high protein
C
57

concentrations of the yeast two-hybrid system is consistent with an enzyme-substrate
interaction, for which only modest affinities are expected.

58

Figure 13. Psy2-Pph3 regulates the phosphorylation state of Rad53. (A) Yeast two-hybrid analysis
of Psy2 interaction with Rad53. pGBT9 expressing the Gal4BD-Psy2 fusion protein was introduced
into the Y190 tester strain along with pACTIIst expressing the Gal4AD fused to full-length Rad53 or
fragments encompassing the Rad53 FHA1, kinase (KD) or FHA2 domains. (B-F) Western blot analysis
of Rad53 phosphorylation state. (B and C) in vitro assay of Rad53 dephosphorylation by Psy2-Pph3.
Incubation of Rad53 isolated from E. coli (B) or MMS-treat yeast (C) cells with immunoprecipitates
from whole-cell lysates of the indicated yeast strains (isogenic with W1588-4C). (D) Effect of psy2 
and pph3  on Rad53 activation in response to MMS treatment. Log phase cultures of the indicated
strains (isogenic with BY4741 or RDKY3615) were treated with 0.1% MMS for 0, 1, and 2 hours.
Western blot analysis of phosphoglycerate kinase (PGK) was used for loading control. (E) Effect of
psy2  and pph3  on spontaneous Rad53 phosphorylation in tof1  and mrc1  cells. Undamaged log
phase cultures of the indicated strains (isogenic with BY4741) were analyzed. (F) Effect of psy2  and
pph3  on dephosphorylation of Rad53 during recovery from MMS exposure in S phase. The indicated
strains (isogenic with BY4741) were synchronized in G1 at 30°C, released into 0.033% MMS for 1
hour and then shifted to YPD. Rad53* denotes phosphorylated Rad53.
59

Figure 13.

We next determined whether the Psy2-Pph3 complex can directly dephosphorylate
Rad53 in vitro. Using recombinant expression in E. coli, we obtained Rad53 that was
autophosphorylated on many of the sites induced by MMS in vivo (Gilbert et al, 2001;
Smolka et al, 2005), and incubated it with immunoprecipitates of whole-cell extracts
from yeast containing untagged or C-terminally TAP-tagged Psy2 or Pph3.
Immunoprecipitate from untagged cells results in no observable Rad53
dephosphorylation, however, immunoprecipitate from Pph3-TAP cells results in the
60

efficient dephosphorylation of Rad53 (Figure 13B). Immunoprecipitate from Psy2-
TAP cells also dephosphorylates Rad53, but not when purified from pph3  cells.
Finally, immunoprecipitate from Psy2-TAP pph3 cells harboring a plasmid
expressing wild-type Pph3, but not the catalytically inactive Y121N mutant,
dephosphorylates Rad53 (Figure 14). Immunoprecipitates from Pph3-TAP and Psy2-
TAP cells also dephosphorylate Rad53 isolated from MMS-treated yeast cells (Figure
13C). Taken together, these results suggest that Psy2-Pph3 is a Rad53 phosphatase.

Figure 14. in vitro desphosphorylation of Rad53 relies on a catalytically active Pph3. Incubation of
Rad53 isolated from E. coli with immunoprecipitates from whole-cell lysates of PSY2-TAP pph3  cells
(FR1044) harboring YEplac195 or YEplac195 carrying a wild-type or mutant allele of PPH3. Rad53*
denotes phosphorylated Rad53.

Psy2-Pph3 promotes dephosphorylation of activated Rad53 during recovery from
MMS treatment

We next examined the role of Psy2-Pph3 in regulating Rad53 phosphorylation
in vivo. Asynchronous wild-type cells accumulate partially phosphorylated Rad53
after one hour of treatment with MMS and fully phosphorylated Rad53 after two hours
(Figure 13D). Loss of PSY2 or PPH3 results in the more rapid accumulation of fully
phosphorylated Rad53, relative to wild-type cells. Consistent with the observed
suppression of MMS sensitivity, loss of PSY2 or PPH3 partially restores the MMS-
induced phosphorylation of Rad53 in rad9  and mec1  mutants (Figure 13D).
Additionally, disruption of PSY2 or PPH3 increases the amount of phosphorylated
Rad53 in undamaged tof1 and mrc1 cells (Figure 13E), which are known to have
61

elevated levels of spontaneous DNA damage due to defective replication forks (Foss,
2001; Katou et al, 2003; Osborn & Elledge, 2003; Szyjka et al, 2005; Tourriere et al,
2005). These effects are consistent with previously observed genetic relationships
between tof1  or mrc1  and psy2  or pph3 mutants (Keogh et al, 2006; O'Neill et
al, 2004). To determine whether Psy2-Pph3 regulates Rad53 activation specifically in
the intra-S phase checkpoint, we synchronized cells in G1 and released them into S
phase in the presence of MMS. pph3 and psy2  mutants accumulate more hyper-
phosphorylated Rad53 than wild-type cells (Figure 13F), consistent with a hyperactive
intra-S checkpoint response.
We next examined the phosphorylation state of Rad53 in cells recovering from
MMS treatment in S phase. Following removal of MMS from the media, virtually full
dephosphorylation of Rad53 was observed in wild-type cells by 60 minutes (Figure
3F). In contrast, Rad53 remained hyper-phosphorylated in pph3 and psy2 mutants
throughout the entirety of the two-hour experiment. These data demonstrate that the
Psy2-Pph3 phosphatase is required for the dephosphorylation of Rad53 during
recovery from the intra-S DNA damage checkpoint.

62

Figure 15. WT, pph3 , and psy2  cells enter S-phase with similar kinetics in the presence of
MMS. Budding index analysis of wild-type (SSy187), pph3  (SSy188), and psy2  (SSy189) cells that
were blocked in G1 with -factor, released into S phase in the presence of 0.033% MMS for 1 hour,
and then shifted to YPD. All steps were conducted at 30°C.

Psy2-Pph3 is required for efficient DNA synthesis during recovery from MMS
treatment
To determine the impact of misregulated Rad53 on the kinetics of DNA
replication, we monitored DNA content by flow cytometry during and after exposure
to MMS. Figure 17A shows that wild-type, psy2  and pph3 cells synchronized in G1
and released into YPD in the absence of MMS progress similarly through S-phase.
Cells were also synchronized in G1, released into YPD containing 0.033% MMS for 1
hour, and then transferred to YPD. All three strains enter S-phase in the presence of
MMS with similar kinetics based on budding morphology (Figure 15). As expected,
DNA synthesis is inhibited soon after entry into S phase in wild-type, psy2 , and
pph3  cells. However, psy2 and pph3 cells synthesize significantly less DNA than
wild-type cells during incubation in MMS, consistent with hyperactivation of the
checkpoint (Figure 17B, time=60/0). In addition, psy2  and pph3 cells require
63

significantly more time than wild-type cells to complete replication following removal
of MMS, with wild-type cells largely completing replication by 60 minutes and cells
lacking Psy2-Pph3 requiring at least 180 minutes (Figure 17B and data not shown).
psy2  and pph3 cells also require significantly more time than wild-type cells to re-
enter the cell cycle (Figure 16). These data demonstrate that cells deficient for Psy2-
Pph3 elicit a hyperactive intra-S DNA damage response and fail to resume DNA
synthesis and re-enter the cell cycle normally following the removal of MMS.

Figure 16. MMS recovery kinetics of WT and ybl046w  cells are indistinguishable. DNA content
analysis (A) and budding index (B and C) of wild-type (SSy187), pph3  (SSy188), psy2  (SSy189),
and ybl046w  (SSy235) cells that were blocked in G1 with -factor, released into S phase in the
presence of 0.033% MMS for 1 hour, and then shifted to YPD in the presence of -factor. All steps
were conducted at 23°C.

64

Figure 17. Psy2/Pph3 promotes efficient replication restart during recovery from MMS exposure
in S phase. (A) DNA content analysis by flow cytometry of wild-type (SSy187), pph3  (SSy188), and
psy2  (SSy189) cells that were blocked in G1 with -factor and released into S phase in YPD. All steps
were conducted at 30°C. (B) DNA content analysis by flow cytometry of wild-type (SSy187), pph3 
(SSy188), and psy2  (SSy189) cells that were blocked in G1 with -factor, released into S phase in the
presence of 0.033% MMS for 1 hour, and then shifted to YPD. All steps were conducted at 30°C. (C)
BrdU incorporation analysis of wild-type (SSy205) and pph3  (SSy210) cells treated as in panel B but
grown at 23°C. Immunoprecipitated DNA sequences at ARS607 and at distances of 15, 33, 53, and 73
kb were detected by PCR amplification. (D and E) 2D gel analysis of samples collected in panel B that
were digested with BamHI and NcoI. Blots were probed for ARS1 (D) or ARS603 (E). Solid
arrowheads indicate large replication bubbles, open arrowheads indicate large replication forks, and
caret indicates small and medium replication forks.

65

Psy2-Pph3 promotes restart of stalled replication forks
Rad53 activation during S phase inhibits the initiation of unfired replication
origins and prevents the collapse of replication forks encountering a damaged
template, presumably by maintaining the association of replisome components with
the fork structure (Cobb et al, 2003; Lopes et al, 2001; Santocanale & Diffley, 1998;
Shirahige et al, 1998; Tercero & Diffley, 2001). These functions of Rad53 enable the
rapid resumption of DNA synthesis following removal of DNA damage, which is
normally correlated with dephosphorylation of Rad53 (Tercero et al, 2003). Thus, we
hypothesized that dephosphorylation of Rad53 by Psy2-Pph3 facilitates the
resumption of normal DNA synthesis following removal of MMS by regulating
replication fork restart and/or the firing of replication origins. To directly examine
replication fork progression in the absence of Psy2-Pph3 function, we used BrdU
incorporation to monitor a replication fork that initiates at ARS607 and progresses
across a 70 kb, origin free region of chromosome VI. Upon entry into S-phase, BrdU
is efficiently incorporated into ARS607 in wild-type and pph3 cells, reflecting
normal entry into S-phase and initiation at this early origin (Figure 17C). During
incubation with MMS, both wild-type and pph3  cells incorporate BrdU at a DNA
sequence 15 kb from ARS607, but do not incorporate BrdU at more distal sites,
consistent with a reduced rate of elongation in the presence of DNA damage. Upon
release from MMS, BrdU incorporation in wild-type cells at DNA sequences 33, 53,
and 73 kb from the origin is detected at approximately 15, 45, and 60 minutes,
respectively. In pph3  cells, BrdU incorporation at these sequences is delayed at least
30 minutes, with replication at the most distal sequences beginning only after 90
66

minutes. These results demonstrate that Psy2-Pph3 promotes the resumption of DNA
synthesis after MMS-induced fork arrest and suggest that this function is mediated by
dephosphorylation of Rad53.
We next analyzed initiation of both early and late origins during and after
exposure to MMS. As above, cells were released synchronously into S phase in the
presence of 0.033% MMS. Initiation of the early origin ARS1 occurs with similar
timing and efficiency in wild-type, psy2 , and pph3  cells (Figure 17D). In addition,
initiation of the late origin ARS603 is blocked in wild-type, psy2 , and pph3 cells
(Figure 17E), demonstrating that Psy2-Pph3 is not required to inhibit late origin firing.
Upon release of wild-type cells from MMS, ARS603 replication occurs “passively” by
replication fork restart (Figure 17E). In contrast, upon release of psy2  and pph3
cells from MMS, ARS603 replication is delayed. Replication in these mutants occurs
not by restart of stalled replication forks, but rather by initiation of ARS603
establishing new forks (Figure 17E). Some replication forks persist at ARS1 in psy2 
and pph3 cells (Figure 17D), consistent with delayed replication restart of forks that
initially stall near ARS1 during MMS treatment. Nevertheless, these forks appear
stable, as we observed no evidence of fork collapse, such as broken bubbles or forks,
which would appear as aberrantly migrating forms on the 2D gels. Together, these
data strongly suggest that replication forks established at early origins in psy2 and
pph3  cells are stable, but unable to efficiently resume DNA synthesis following
removal of MMS, and instead, new forks are initiated from late origins to complete
genome replication, despite the persistence of hyper-phosphorylated Rad53.

67

H2AX-independent regulation of Rad53 by Psy2-Pph3
During the course of this study it was reported that Psy2 forms a complex with
Pph3 and Ybl046w that dephosphorylates H2AX, and that this function is required to
inactivate Rad53 during prolonged recovery from a double-strand break (Keogh et al,
2006). To further examine the suggested connection between H2AX and Rad53
dephosphorylation, we examined the MMS sensitivity of an H2AX mutant that cannot
be converted to H2AX by phosphorylation (hta1-S129A). An additive increase in
MMS sensitivity is observed when PPH3 is disrupted in the hta1-S129A mutant,
demonstrating that Pph3 has H2AX-independent functions (Figure 18A).

Figure 18. Psy2-Pph3 has H2AX-independent functions. (A) Colony survival assays of
yeast containing wild-type (YMV2) or mutant alleles of PPH3 and/or HTA1 (H2AX). Colony forming
units (cfu) were counted three days after deposition onto YPD-agar media containing the indicated
concentration of MMS. (B) Western blot analysis of H2AX or phosphoglycerate kinase (PGK, loading
control) from log phase cultures of the indicated strains (BY4741 background) that were treated with
0.1% MMS for 0 or 1 hour. Strains are as listed in Table 2.

68

In addition, deletion of YBL046W, which is required for H2AX dephosphorylation,
does not impart the phenotypes observed with deletion of PPH3 or PSY2, including:
sensitivity to MMS (Figure 10B), genetic interactions with cell cycle checkpoint
mutants (rad9 , mec1 , dun1, or ptc2 ; Figure 11A-D), and delayed reentry into the
cell cycle during recovery from MMS treatment (Figure 16). Most importantly,
ybl046w  does not show a Rad53 dephosphorylation defect (Figure 13F), despite its
H2AX dephosphorylation defect (Keogh et al, 2006 and Figure 18B). Thus, while
Psy2-Pph3 requires Ybl046w to dephosphorylate H2AX, it does not require Ybl046w
to dephosphorylate Rad53, mechanistically separating Rad53 dephosphorylation from
that of H2AX.
Activation of Rad53 by phosphorylation and the initiation of cell cycle
checkpoints is essential for the maintenance of genome integrity. While the
phosphorylation of Rad53 has been intensively studied, its deactivation, which is
thought to be necessary for cells to re-enter the cell cycle, has received little attention
and is much less understood. To date, the only phosphatases implicated in Rad53
deactivation are Ptc2 and Ptc3 (Leroy et al, 2003). In their absence, cells suffering a
persistent double-strand break are unable to adapt to Rad53-mediated G2/M arrest. In
addition, Rad53 dephosphorylation after induction of a persistent double-strand break
has been suggested to depend on dephosphorylation of H2AX, which is mediated by
a complex of Pph3, Psy2, and Ybl046w.
We demonstrate that Psy2 and Pph3 form a complex that regulates Rad53
activity in MMS-treated cells, in a manner independent of its role with Ybl046w in
69

H2AX dephosphorylation. Our results suggest that Ybl046w is a specificity factor for
the Psy2-Pph3 phosphatase complex. That Rad53 dephosphorylation following repair
of a double strand break via single-stand annealing is independent of Pph3 in h2a-
129A cells (Keogh et al, 2006) may reflect switching to an alternative, Pph3-Psy2-
independent repair pathway (Downs et al, 2000; Redon et al, 2003), perhaps an
adaptation pathway involving Ptc2 and/or Ptc3 (Leroy et al, 2003). This is consistent
with the synergistic increase in MMS sensitivity of ptc2  cells caused by disruption of
PSY2 or PPH3. That Psy2-Pph3 regulates Rad53 is supported by a body of evidence
that includes: genetic analyses placing Psy2-Pph3 downstream of Rad53 in a role that
antagonizes the DNA damage checkpoint response; yeast two-hybrid data indicating
that Psy2 interacts with Rad53; and biochemical results demonstrating that Psy2-Pph3
can directly dephosphorylate Rad53 in vitro and that Psy2-Pph3 is required for
dephosphorylation in vivo of activated Rad53 during recovery from MMS treatment in
S phase.
DNA damage checkpoint-mediated activation of Rad53 regulates DNA
synthesis by stabilizing replication forks that are already initiated and by delaying the
initiation of late-firing origins. Both functions appear normal in cells bereft of Psy2-
Pph3 (Fig 17D and E). However, replication forks fail to restart normally in psy2  or
pph3  cells after removal of the genotoxic stress. The severe delay in replication fork
restart is likely the result of Rad53 remaining in a hyper-phosphorylated state;
however a previous study indicated that fork progression rates are unaffected by the
presence or absence of Rad53 (Tercero & Diffley, 2001), suggesting that
dephosphorylation of other factors may be involved. DNA replication eventually
70

resumes through firing of late origins. This late origin firing is consistent with, and
likely enabled by, the delayed restart of early-established forks that would otherwise
passively replicate the late origin, as is observed with wild-type cells, and is
particularly remarkable given the persistence of activated Rad53. These results suggest
a functional uncoupling of the Rad53-dependent processes of fork stabilization and
late-origin control. Precedence for such an uncoupling is provided by studies of the
mec1-100 hypomorphic allele, which is proficient in stabilizing replication forks, but
defective in preventing late origin firing (Tercero et al, 2003).
How might activated Rad53 independently regulate replication fork restart and
late origin firing? One possibility is that fork stabilization and the inhibition of late
origin firing are mediated by distinct Rad53 phosphorylation patterns, and that these
different phosphorylation patterns are recognized and regulated by different
phosphatases. According to this model, Psy2-Pph3 recognizes a subset of Rad53
phosphorylation sites that are required to stabilize stalled replication forks, and thus, it
is required to restart forks. However, the efficient activation of late origins during
recovery from DNA damage appears to result from regulation of different Rad53
phosphorylation sites by another phosphatase(s). This model is supported by previous
work that has shown that different facets of the checkpoint response controlled by
Rad53 are mediated by different domains of the protein (Pike et al, 2004; Pike et al,
2003; Schwartz et al, 2003). Further support for this model is provided by the
observation that different Rad53 residues are phosphorylated in response to different
types of DNA damage (Smolka et al, 2005; Sweeney et al, 2005). Thus, specific
Rad53 phosphorylation sites may be induced by different types of damage, be
71

regulated by different mechanisms, and mediate different aspects of the DNA damage
response.
Another plausible mechanism separating Rad53-dependent fork stabilization
from origin control is that the hyper-phosphorylated Rad53 apparent in the absence of
Pph3-Psy2 is sequestered at stalled forks, and thus unavailable to act elsewhere.
Therefore, newly synthesized Dbf4 (and perhaps other factors) might escape inhibition
by activated Rad53, and thus promote late origin firing. This model is analogous to
that 16 proposed for hypomorphic Mec1-100 which is proficient for the stabilization
of stalled forks, but defective in late origin control (Tercero et al, 2003).
Checkpoint pathways are well conserved among eukaryotes. Rad53 is the S.
cerevisiae homolog of the mammalian tumor suppressor Chk2 (reviewed in Bartek et
al, 2001). In the presence of DNA damage, Chk2 is phosphorylated and activated by
ATM (Bartek et al, 2001), and then phosphorylates downstream effectors, including
the tumor suppressors p53, BRCA1, and Cdc25. Homologs of Pph3 and Psy2 have
also been identified in higher eukaryotes; in humans the homologs are encoded by PP4
and PP4R3, respectively (Cohen et al, 2005; Gingras et al, 2005). Interestingly, PP4R3
complements the cisplatin hypersensitivity of yeast lacking PSY2, indicating
functional conservation through evolution (Gingras et al, 2005). Furthermore, a
conserved role of PP4 and PP4R3 as a Chk2 phosphatase complex is suggested by a
yeast two-hybrid interaction between PP4R3 and Rad53 (Gingras et al, 2005).
Further understanding the regulation of Rad53 in S. cerevisiae will provide
additional insights into the checkpoint response. In particular, how different
phosphorylation patterns of Rad53 are induced by different forms of genotoxic stress,
72

how these sites are regulated (i.e. sequence specifically or by localization to specific
replication structures), and their functional significance is of great interest. Further
characterizing the role of Psy2-Pph3 in these processes, and determining whether
Psy2-Pph3 dephosphorylates additional replication factors is also crucial to gaining a
clearer understanding of the mechanisms of fork stabilization and origin control.

73

CHAPTER 4

Rad53 Regulates Replication Fork Restart after DNA Damage in Saccharomyces
cerevisiae

All of the contents of this chapter have been published and can be found in
Genes and Development (2008, Vol 22, pp 1906-1920).

Figure Contributions

I contributed the work found in Figures 19-29, and 31.
Oscar M. Aparicio contributed the model found in Figure 30.
Simon R.V. Knott contributed to the computational analysis and plotting of the data in
Figures 19-22, 24-27, and 29.

Overview
S-phase is a particularly vulnerable period for the genome as environmental agents
that damage DNA and intrinsic replication defects can interfere with the replication
process, potentially giving rise to mutations and genomic instabilities. The presence
of DNA damage during chromosomal DNA replication results in a significantly
reduced rate of DNA synthesis in organisms ranging from bacteria to humans. In S.
cerevisiae, the slowed S-phase resulting from DNA alkylation by MMS depends on
checkpoint signaling by Mec1 and Rad53, indicating that slowing of DNA synthesis is
a regulated process (Paulovich & Hartwell, 1995; Paulovich et al, 1997). Similar
radioresistant DNA synthesis occurs in mammalian cells lacking the Mec1-related
protein ATM (Painter & Young, 1980). The exquisite DNA damage sensitivity of
checkpoint defective cells, and the increased cancer susceptibility of organisms with
checkpoint defects, emphasizes the importance of these surveillance mechanisms
(reviewed in Kolodner et al, 2002).
74

Generation of a DNA damage signal during S-phase requires activation of
replication forks (Shimada et al, 2002; Tercero et al, 2003), suggesting that the
encounter of replication forks with damaged sites creates aberrant DNA structure(s),
such as excess ssDNA, which is recognized by checkpoint sensors (reviewed in
Paulsen & Cimprich, 2007). In S. cerevisiae, MMS-induced DNA damage leads to
recruitment of Mec1 kinase and the PCNA-like Rad17-Ddc1-Mec3 complex to sites of
damage, whereupon these proteins further engage Rad9, which mediates Mec1-
dependent phosphorylation of the effector kinases Chk1 and Rad53 (reviewed in Melo
& Toczyski, 2002). Chk1 and Rad53 target downstream factors to delay mitotic
progression, and Rad53 additionally targets factors to induce expression of DNA
metabolism genes, stimulate dNTP production, suppress the initiation of additional
(late-firing) replication origins, and stabilize the function of replication forks.
Activation of Rad53 is critical for survival of yeast cells subjected to DNA
damage or replication stress (reviewed in Branzei & Foiani, 2006). In the absence of
Rad53 (or Mec1), MMS treatment does not slow S-phase and DNA content
approximately doubles; however, these cells are inviable (Paulovich & Hartwell,
1995). Analysis of an individual replicon reveals a significant degree of incomplete
replication, suggesting that replication forks irreversibly collapse in MMS-treated
rad53  and mec1  cells (Tercero & Diffley, 2001). Recovery of cells from inhibition
of DNA synthesis with hydroxyurea (HU) also requires Rad53, as HU-stalled
replication forks degenerate in rad53  cells and these cells are unable to resume DNA
synthesis after removal of the drug (Desany et al, 1998; Lopes et al, 2001; Sogo et al,
2002). These studies have led to the paradigm that a critical function of Rad53 in the
75

S-phase checkpoint pathways is the stabilization of stalled or stressed replication forks
(Branzei & Foiani, 2006).
The role of Rad53 in stabilization of stressed forks and the Rad53-dependence
of S-phase slowing in response to DNA damage implies that fork stabilization by
Rad53 involves direct inhibition of the replication fork. However, careful analysis of
DNA synthesis across a well-characterized chromosome VI replicon indicates that
whereas replication forks progress slowly due to the presence of MMS, they
nevertheless progress with similar slow kinetics in wild-type, mec1, and rad53  cells
(Tercero & Diffley, 2001). These findings have suggested that Mec1 and Rad53 do
not regulate fork progression as a consequence of replication fork stabilization, and
further that replication initiation of normally dormant and late-firing origins must
account for the accelerated S-phase of rad53  and mec1 cells. Indeed, analysis of
cells carrying the hypomorphic mec1-100 allele supports this idea (Paciotti et al, 2001;
Tercero et al, 2003). These cells fail to restrain late origin firing and do not slow
replication in MMS; however, mec1-100 cells remain viable and show minimal
evidence of fork dysfunction in MMS, suggesting that fork stabilization operates
normally. Thus, defective fork stabilization correlates with drug sensitivity, whereas
deregulation of origin firing correlates with the failure to slow S-phase.
The conclusion that Rad53 does not directly modulate the rate of fork
progression is challenged by the recent characterization of cells lacking the Psy2-Pph3
phosphatase, which acts to dephosphorylate, and hence deactivate, Rad53 during
recovery from MMS exposure (O'Neill et al, 2007). After transient exposure to MMS
during early S-phase, ps y 2∆ and pph3∆ cells are delayed in completing bulk DNA
76

replication, and analysis of BrdU-incorporation along the aforementioned
chromosome VI replicon indicates that replication fork progression is delayed in the
absence of Psy2-Pph3. The correlation between the delayed replication restart and
delayed dephosphorylation of Rad53 suggests that deactivation of Rad53 is required
for replication restart following DNA damage. The failure to dephophosphorylate
H2a after DNA damage does not account for the replication restart defect of psy2∆ or
pph3∆ cells (Keogh et al, 2006; O'Neill et al, 2007). Nevertheless, the possibility
remains that the role of Psy2-Pph3 in replication fork restart reflects
dephosphorylation of a different, still unrecognized, Psy2-Pph3 substrate.
In this study, we have tested the hypothesis that Rad53 controls replication
fork restart by monitoring the progression of replication forks in MMS-damaged cells,
under different conditions of Rad53 activity. We show that replication forks progress
more slowly in pph3 ∆ cells in the presence of MMS, and in cells recovering from
MMS damage. In contrast, antagonism of Rad53 activity in these pph3∆ cells restores
rapid DNA synthesis at forks during recovery, indicating that deactivation of Rad53 is
sufficient to allow fork restart. We also reveal the involvement of Ptc2 in
dephosphorylation of Rad53 in MMS-treated pph 3∆ cells, explaining how these cells
eventually complete DNA synthesis and resume growth, and further supporting the
connection between Rad53 deactivation and replication restart. These results provide
important new insights into the mechanism of replication fork stabilization and restart,
as well as coordination with DNA repair.

77

Results and Discussion

Pph3 is required for replication fork progression through damaged DNA
To examine the role of Rad53 in regulation of replication fork dynamics on a
damaged DNA template, we have analyzed replication fork activity in pph3∆ cells,
which lack the Rad53 phosphatase, Psy2-Pph3. Previous work showed that cells
lacking PPH3 or PSY2 replicate normally in the absence of DNA damage, but are
delayed in completing replication during recovery from MMS-induced DNA damage,
suggesting a defect in replication fork restart (O'Neill et al, 2007). To obtain a
comprehensive view of replication fork dynamics, we have monitored replication of
two well-characterized expanded replicons on chromosomes III and VI (Labib et al,
2000; Szyjka et al, 2005; Tercero & Diffley, 2001), using improved methods for BrdU
incorporation into S. cerevisiae (Viggiani & Aparicio, 2006), immunoprecipitation of
BrdU-labeled DNA (Katou et al, 2003; Szyjka et al, 2005), and analysis with
oligonucleotide-tiling-microarrays, which together we refer to as BrdU-IP-chip.
We began by comparing replication of wild-type and pph3∆ cells during
constant exposure to MMS. G
1
-synchronized cells were released from -factor arrest
into rich medium containing 0.033% MMS; aliquots of this culture were exposed to
BrdU for 15 minute pulses and harvested for DNA isolation. BrdU incorporation
occurs at the early origins ARS606 and ARS607 and at surrounding sequences during
the 30-45 minute pulse period, in both wild-type and pph3∆ cells (Fig. 19A). Little if
any BrdU incorporation is detected at these origins subsequently. These results
indicate that both strains initiate chromosomal DNA replication with similar
78

dynamics, as demonstrated previously by analysis of replication initiation structures
with 2D-gels (O'Neill et al, 2007). Analysis of budding kinetics also supports the
conclusion that pph3∆ cells progress through the G1-S transition like wild-type cells
(Fig. 19B). Thus, replication initiation at early origins appears normal in the absence
of PPH3.
The progression of replication forks away from ARS607 is impaired in the
absence of PPH3. During the 45-60 minute period, little BrdU incorporation is
observed at ARS607. However, two flanking peaks of BrdU incorporation are
observed, reflecting DNA synthesis at each replication fork emanating from ARS607
(Fig. 19A, the leftward-moving fork partially converges with the rightward-moving
fork from ARS606). These patterns appear similar in wild-type and pph3∆ cells;
however, the rightward-moving ARS607 replication fork appears to progress ~10kb
further from the origin in wild-type cells. The more distant progression of this
replication fork in wild-type cells becomes more pronounced over the time-course. By
the 105-120 minute interval, BrdU incorporation occurs over a large region of
chromosome VI extending ~65kb from ARS607 toward the telomere in wild-type
cells, whereas BrdU incorporation in pph3∆ cells extends only ~35kb from ARS607.
Wild-type cells also complete replication of the region between ARS606 and ARS607
by ~90 minutes, while BrdU incorporation in this region continues through 120
minutes in the pph3∆ cells.
The chromosome VI results are supported by analysis of the expanded
chromosome III replicon, demonstrating limited progression of the leftward-moving
replication fork from ARS306 in pph3∆ cells compared with wild-type cells (Fig. 20).
79

We used a data normalization method that allows us to make semi-quantitative
comparisons between hybridizations, and we note that pph3∆ cells reproducibly
incorporate less BrdU than wild-type cells during each pulse (especially later times)
based on the total area of BrdU enrichment (Fig. 19A and 20 and see below). The
chromosome III and VI data are also consistent with the kinetics of bulk DNA
synthesis as determined by FACScan analysis, with a higher rate of bulk DNA
synthesis occurring in wild-type versus pph3∆ cells (Fig. 19C). Together, these data
clearly demonstrate limited progression of replication forks in MMS-treated pph3∆
cells, confirming a role for Pph3 in the function of replication forks encountering
damaged DNA (O'Neill et al, 2007).

80

Figure 19. PPH3 is required for progression of replication forks along Chromosome VI during
constant MMS exposure. Wild-type (WT) (SSy419) and pph3  (SSy420) cells were blocked in G
1
-
phase with -factor and released synchronously into YEPD containing 0.033% MMS at 23
o
C (Time =
0). (A) Aliquots of each culture were incubated with BrdU for the indicated time intervals and
harvested for BrdU-IP-chip analysis using an oligonucleotide-tiling array covering the indicated portion
of chromosome VI. Probes representing statistically significant regions of BrdU incorporation were
determined by a two-state Hidden Markov Model (Xu et al, 2006) and are highlighted in blue. Aliquots
of each culture were harvested at the indicated times for determination of budding index (B) and DNA
content (C).

81

Figure 20. PPH3 is required for progression of replication forks along Chromosome III during
constant MMS exposure. BrdU-IP-chip analysis of the indicated chromosome III region in the
experiment described in Figure 19 is shown. The cluster of inefficient origins associated with the HML
locus is indicated.

Pph3 is required for restart of stalled replication forks during DNA damage
recovery
Cells lacking PPH3 also are delayed in resumption of DNA synthesis
following removal of MMS, suggesting that continued presence of DNA damage is
not the cause of delayed replication restart. To examine replication fork restart
kinetics during DNA damage recovery, we released G
1
-synchronized cells into rich
medium containing MMS for 60 minutes to allow early origin initiation and fork
stalling, after which the MMS was quenched and washed-out. BrdU-IP-chip analysis
shows replication forks flanking ARS607 during the 45-60 minute BrdU pulse in
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MMS in both wild-type and pph3∆ cells (Fig. 21A). Upon removal of MMS (Time =
0), replication forks progress rapidly in wild-type cells with the leading edge of BrdU
incorporation progressing ~40kb in 45 minutes (Fig. 21A), which is significantly more
rapid than its replication during constant MMS exposure (comparing total time after
G
1
release) (Fig. 19A). In contrast, the leading edge of BrdU incorporation in pph3∆
cells progresses ~20kb in 60 minutes (Fig. 21A).
Analysis of the expanded chromosome III replicon buttresses the chromosome
VI results. The ARS306 replication fork exhibits limited progression during DNA
damage recovery in pph3∆ cells compared with wild-type (Fig. 22). Furthermore,
bulk DNA synthesis during MMS recovery also is reduced significantly in pph3∆
cells, consistent with the reduced progression and levels of BrdU incorporation in
these cells (Fig. 21B) and (O'Neill et al, 2007). Together, these results strengthen and
extend our previous conclusion that replication fork restart during DNA damage
recovery requires PPH3 (O'Neill et al, 2007).
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Figure 21. PPH3 is required for replication fork restart along Chromosome VI during DNA
damage recovery. WT (SSy419) and pph3  (SSy420) cells were blocked in G
1
-phase with -factor
and released synchronously into YEPD containing 0.033% MMS for one hour at 23
o
C; MMS was
washed out (Time = 0) and cells were suspended in YEPD. (A) Aliquots of each culture were incubated
with BrdU for the indicated periods and analyzed as described in Fig. 19. (B) Aliquots of each culture
were harvested at the indicated times for DNA content analysis. “MMS” samples are after one hour of
incubation with MMS.

84

Figure 22. PPH3 is required for replication fork restart along Chromosome III during DNA
damage recovery. BrdU-IP-chip analysis of the indicated chromosome III region in the experiment
described in Figure 20 is shown.

Restoration of Pph3 function during DNA damage recovery enables rapid
replication restart
Deregulation of Rad53 activity in pph3∆ cells by elimination of Pph3 function
might disrupt the normal stabilization of forks, resulting in the observed defect in fork
progression (Fig. 19A). However, a number of findings argue that replication forks
are stable in pph3∆ cells, including relatively modest lethality in the presence of MMS
and no evidence of the replication fork collapse observed in rad53∆ cells by 2D-gel
analysis (O'Neill et al, 2007). If forks indeed remain stable in pph3∆ cells, we
reasoned that restoration of PPH3 function during recovery should enable rapid
resumption of DNA synthesis.
85

To test the idea that stalled replication forks in pph3∆ cells remain stable and
capable of rapid restart, we placed the endogenous PPH3 gene under control of the
GAL promoter to enable its induction subsequent to replication fork stalling by MMS.
G
1
-synchronized wild-type, pph3∆, and GAL-PPH3 cells grown in raffinose (to
repress GAL-PPH3) were released from arrest into raffinose medium containing
MMS. After 90 minutes to allow initiation of early origins and stalling of replication
forks, the cultures were split into fresh media lacking MMS and containing raffinose
or galactose (to induce GAL-PPH3) (Fig. 23A). In raffinose or galactose, wild-type
cells resume rapid DNA replication and complete bulk DNA synthesis between 120
and 180 minutes after removal of MMS, whereas pph3∆ cells do not complete DNA
synthesis by 240 minutes (Fig. 23B and C). In raffinose, GAL-PPH3 cells complete
DNA synthesis with kinetics similar to pph3∆ cells, demonstrating effective repression
of PPH3 activity (Fig. 23B). However, in galactose, GAL-PPH3 cells complete DNA
synthesis by 180 minutes, only slightly behind wild-type cells, and significantly
sooner than pph3∆ cells (Fig. 23C). The ~30 minute delay in replication restart of
GAL-PPH3 cells compared with wild-type (Fig. 23C, compare WT 90’ with GAL-
PPH3 120’) correlates with the time required to induce expression of PPH3 from the
GAL promoter (data not shown). The rapid restart of replication upon restoration of
PPH3 function is consistent with the conclusion that stalled replication forks in MMS-
treated pph3∆ cells are stable and poised for restart.
86

Figure 23. Restoration of PPH3 during DNA damage recovery facilitates replication restart. (A)
WT (SSy187), GAL-PPH3 (SSy250), and pph3  (SSy188) cells were blocked in G
1
-phase with -factor
in YEP-Raffinose and released into YEP-Raffinose containing 0.033% MMS for 90 minutes at 23
o
C.
MMS was washed out (Time = 0) and cells were split into YEP-Raffinose (RAF) (B) and YEP-
Raffinose + 0.5% Galactose (GAL) (C). (B, C) Cells were harvested at the indicated time points for
DNA content analysis.

Deactivation of Rad53 is sufficient to restart stalled replication forks in pp h 3∆
cells
The role of Pph3 in Rad53 deactivation by dephosphorylation implies that
replication fork restart requires deactivation of Rad53. However, the role of Pph3 in
replication restart may depend on dephosphorylation of other Pph3 substrates, either
instead of, or in addition to Rad53. If Rad53 deactivation is the sole requirement for
87

replication restart in pph3∆ cells, then elimination of its activity during the DNA
damage recovery period should enable fork restart. However, it is necessary to
preserve Rad53 activity to stabilize forks encountering DNA damage during the initial
period of MMS exposure. To exert this control over Rad53 activity, we have used a
dominant-negative, kinase-dead allele of RAD53 (rad53-K221A,D339A, herein
referred to as rad53-KD), under control of the GAL promoter. Galactose-induced
overexpression of Rad53-KD suppresses the kinase activity of previously DNA-
damage-activated, endogenous Rad53 (Pellicioli et al, 1999). Our approach was to
expose pph3∆ cells to MMS to stall replication forks in the presence of wild-type,
endogenous Rad53, and following removal of MMS, to induce Rad53-KD to
antagonize the activated, wild-type Rad53.
Antagonism of activated Rad53 by expression of Rad53-KD restores rapid
replication kinetics during DNA damage recovery. Analysis of bulk DNA synthesis
shows delayed completion of DNA replication after MMS damage in pph3∆ cells
harboring empty vector (Fig. 24A). However, pph3∆ cells harboring pGAL-rad53-KD
completed DNA replication between 120-150 minutes after the removal of MMS and
addition of galactose. These replication kinetics were similar to those of wild-type
cells recovering from MMS-treatment (Fig. 24A, compare with 23B). We confirmed
that endogenous Rad53 kinase activity was suppressed by overexpression of rad53-
KD by in situ Rad53 kinase assay (Fig. 24B). Thus, artificial deactivation of Rad53
can replace Pph3 function in replication restart during DNA damage recovery,
indicating that deactivation of Rad53 is the critical function of Pph3 in replication
88

restart during DNA damage recovery. These results strongly suggest that Rad53
deactivation regulates replication restart.
To examine the effect of Rad53-KD expression specifically on replication fork
restart, we used BrdU-IP-chip. In pph3∆ cells harboring empty vector, very limited
replication fork progression occurs during two hours of recovery from MMS-treatment
(Fig. 24C). In contrast, pph3∆ cells expressing Rad53-KD exhibit accelerated
replication fork progression. Rightward-moving replication forks reach the telomere
during the 60-90 minute pulse covering ~30kb during the 60-90 minute pulse.
Termination of the converging ARS606 and ARS607 replication forks occurs before
the 60-90 minute interval, as minimal BrdU incorporation occurs in this region in cells
expressing Rad53-KD. Analysis of chromosome III-L replication also shows
accelerated replication fork restart resulting from Rad53-KD expression during DNA
damage recovery in pph3∆ cells (Fig. 25). These strains contain ARS305, and
therefore the rightward ARS305 fork converges with the leftward ARS306 fork and
replication termination occurs in the middle of this region between ARS305 and
ARS306. While BrdU incorporation in the termination region continues through the
90-120 minute period in pph3∆ cells harboring only vector, BrdU incorporation in this
region ceases before the 60-90 minute interval in pph3∆ cells expressing Rad53-KD.
Taken together, the replication fork kinetics of chromosome III and VI replicons, as
well as the kinetics of bulk DNA replication, support the conclusion that deactivation
of Rad53 regulates replication fork restart.

89

Figure 24. Antagonism of Rad53 promotes fork restart along Chromosome VI in pph3  cells
recovering from MMS. pph3  cells transformed with pGal (SSy395) or pGal-rad53-KD (SSy396)
were blocked in G
1
-phase with -factor in YEP-Raffinose and released into YEP-Raffinose containing
0.033% MMS for 90 minutes at 23
o
C (MMS). MMS was washed out (Time = 0) and cells were
suspended in YEP-Raffinose + 0.5% Galactose. Aliquots of each culture were harvested at the
indicated times for DNA content analysis (A), in situ Rad53 kinase assay (B), or incubated with BrdU
for the indicated periods and analyzed as described in Fig. 19 (C).

90

Figure 25. Antagonism of Rad53 promotes fork restart along Chromosome III in pph3  cells
recovering from MMS. BrdU-IP-chip analysis of the indicated chromosome III region in the
experiment described in Figure 24 is shown.

Overlapping functions of Pph3 and Ptc2 in Rad53 deactivation and replication
restart
Our results strongly suggest that restart of DNA damage-stalled replication
forks requires Rad53 deactivation by Pph3. Nevertheless, pph3∆ cells eventually
appear to complete chromosome replication during recovery from MMS based on bulk
DNA content analysis (O'Neill et al, 2007 and see below). We determined the
viability of pph3∆ cells recovering from transient exposure to MMS, analogous to the
earlier MMS recovery experiments. G
1
-synchronized cells were released into S-phase
in the presence of MMS, and plated on rich medium lacking MMS to determine
viability (Fig. 26). Even after three hours of MMS exposure, pph3∆ cells show
91

minimal loss of viability when permitted to recover, although the colonies appear
somewhat smaller after longer exposure to MMS. These data indicate that pph3∆ cells
eventually complete chromosomal replication and resume proliferation, and suggest
that an alternative means exists for deactivation of Rad53.

92

Figure 26. Ptc2 contributes to viability, dephosphorylation of Rad53, and replication restart in
pp h3 cells. (A) WT (SSy187), pph3  (SSy188), ptc2  (SSy248), and ptc2  pph3  (SSy249) cells
were arrested in G1-phase with -factor at 30
o
C and released into YEPD containing 0.033% MMS at
30
o
C (Time = 0). At the indicated times, cells were ten-fold serially diluted and plated onto YEPD.
Plates were incubated two days at 30
o
C and imaged. (B, C) Strains in (A) were arrested in G1-phase
with -factor at 30
o
C and released into YEPD containing 0.033% MMS for 45 minutes (MMS). MMS
was washed out, and cells were suspended in YEPD at 30
o
C (Time = 0). Cells were harvested at the
indicated time points for immunoblot analysis with anti-Rad53 antibody (B) and DNA content analysis
(C). (D, E) ptc2  pph3  cells harboring plasmid pGal (SSy526) or pGal-rad53-KD (SSy527) were
arrested in G
1
-phase with -factor at 25
o
C in YEP-Raffinose and released into YEP-Raffinose
containing 0.033%MMS and BrdU for 90min at 23
o
C (MMS). MMS was quenched and washed out
and cells were suspended in YEP-Raffinose + 0.5% Galactose containing BrdU (Time = 0). Cells were
harvested at the indicated times and analyzed for BrdU incorporation (D) and DNA content analysis
(E).
93

The Ptc2 and Ptc3 phosphatases have been implicated in the dephosphorylation
of Rad53 during prolonged recovery from a repairable dsDNA break or adaptation to
an irreparable dsDNA break (Leroy et al, 2003). Hence, we tested the effect of
deleting PTC2 (the major activity in dsDNA break recovery) on the viability of wild-
type and pph3∆ cells subjected to transient MMS exposure (Fig. 26A). The ptc2∆
cells show no sensitivity to transient MMS exposure; however, ptc2∆ pph3∆ cells
show sensitivity to acute MMS exposure, with over ten-fold loss of viability per hour
of MMS exposure. Given the previously characterized role of Ptc2 in Rad53
dephosphorylation after dsDNA breaks and the present data, we conclude that Pph3
and Ptc2 play overlapping roles in maintaining viability during recovery from MMS-
induced DNA damage through deactivation of Rad53.
To confirm that the lethality caused by PTC2 deletion reflects its function as a
Rad53 phosphatase, we analyzed Rad53 phosphorylation by immunoblotting. Wild-
type, pph3∆, ptc2∆, and ptc2∆ pph3∆ cells were synchronized in G
1
-phase with
-factor, released into S-phase in the presence of MMS for one hour to allow
replication initiation and Rad53 activation, and then allowed to recover from MMS.
All strains show hyperphosphorylated Rad53 after MMS treatment (Fig. 26B). Upon
removal of MMS, wild-type cells show a decreasing proportion of
hyperphosphorylated form(s) of Rad53 with the band corresponding to
unphosphorylated Rad53 predominating by 2 hours. A similar kinetic of Rad53
dephosphorylation occurs in ptc2∆ cells, though there appears to be a slight delay (~30
minutes) based on analysis of multiple experiments (Fig. 26B and data not shown).
This result suggests that Ptc2 plays a minor role in Rad53 dephosphorylation during
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MMS recovery. As shown previously, pph3∆ cells are delayed in Rad53
dephosphorylation, with the hyperphosphorylated form(s) predominating for at least 2
hours and remaining abundant for at least 4 hours. Consistent with Pph3 and Ptc2
playing overlapping roles, Rad53 dephosphorylation is further delayed in cells lacking
both PPH3 and PTC2, with the hyperphosphorylated forms predominating after 6
hours of recovery (Fig. 26B). Notably, reduction in the proportion of the slowest
migrating forms of Rad53 (as opposed to complete dephosphorylation) appears to
correlate with replication restart in each of these strains (see below). These results
show that a severe defect in Rad53 dephosphorylation accompanies the lethality of
ptc2∆ pph3∆ cells after MMS treatment.
The severe defect in Rad53 dephosphorylation and lethality of DNA-damaged
cells deficient in both Pph3 and Ptc2 further supports the conclusion that Rad53
deactivation is required for DNA damage recovery, in particular, the restart of stalled
replication forks. This notion predicts that the replication fork restart defect should be
exacerbated in ptc2∆ pph3∆ cells. We examined total DNA content in wild-type,
pph3∆, ptc2∆, and ptc2∆ pph3∆ cells during recovery from MMS (Fig. 26C). Wild-
type, ptc2∆ and pph3∆ cells complete replication after about 1.5, 2, and 3 hours of
recovery, respectively. Strikingly, ptc2∆ pph3∆ cells do not appear to complete
replication in this time course, showing limited increase of DNA content during 6
hours of recovery (Fig. 26C). Consistent with this, fork progression completely stalls
in ptc2∆ pph3∆ cells based on BrdU-IP-chip analysis (Fig. 26D). In fact, we were
unable to measure BrdU incorporation in ptc2∆ pph3∆ cells during MMS recovery
using the pulse-labeling approach, likely because these cells synthesize very little
95

DNA during recovery (Fig. 26C). Thus, we used a cumulative labeling approach with
BrdU present throughout the time-course, including S-phase entry in the presence of
MMS, and during recovery from the MMS (Fig. 26D). In ptc2∆ pph3∆ cells exposed
to MMS, BrdU incorporation ceases ~15-20kb from ARS607, and very little further
incorporation is observed during 2 hours of recovery. Restoration of Pph3 activity in
these cells by induction of GAL-PPH3 during recovery permits resumption of DNA
synthesis from the stalled forks (Fig 26D) and completion of bulk DNA synthesis (Fig.
26E). Analysis of the chromosome III replicon shows similar results (Fig. 27). The
failure of ptc2∆ pph3∆ cells to restart replication provides a compelling cause for the
lethality of these cells after exposure to MMS. Furthermore, the synergistic effect of
loss of two different Rad53 phosphatases on replication fork restart directly supports
the notion that Rad53 deactivation is a key step in this process.

96

Figure 27. Ptc2 contributes to replication restart in pph3  cells. BrdU-IP-chip analysis of the
indicated chromosome III region in the experiment described in Figure 26D is shown.

To verify that failure of Rad53 deactivation causes the failed replication restart
and lethality of ptc2∆ pph3∆ cells, we induced expression of Rad53-KD in wild-type,
pph3∆, ptc2∆, and ptc2∆ pph3∆ cells beginning recovery from MMS-induced DNA
damage (Fig. 28A, “+rad53-KD”). Expression of Rad53-KD has little effect on
kinetics of DNA replication during recovery of wild-type and ptc2∆ cells. As shown
above (Fig. 24), expression of Rad53-KD largely eliminates the delay in replication
restart of pph3∆ cells (Fig. 28A). Consistent with a direct link between defective
Rad53 dephosphorylation and replication restart, expression of Rad53-KD enables
replication restart in ptc2∆ pph3∆ cells, with these cells largely completing replication
by 4 hours. The longer time (compared with pph3∆ cells) required for replication
restart in ptc2∆ pph3∆ cells probably reflects the lack of Ptc2 activity contributing to
97

Rad53 deactivation by Rad53-KD. Indeed, we confirmed that overexpression of
Rad53-KD suppressed the endogenous Rad53-associated kinase activity in ptc2∆
pph3∆ cells (Fig. 28B), however, the rate of Rad53 deactivation was lower than in
pph3∆ cells (Fig. 24B), consistent with Ptc2 contributing to Rad53 deactivation by
Rad53-KD. Finally, expression of Rad53-KD partially rescues the viability of MMS-
treated ptc2∆ pph3∆ cells (Fig. 28C), which is fully consistent with the conclusion that
Rad53 deactivation is necessary and sufficient to restart replication forks stalled by
DNA damage, and thereby avert lethality.

Figure 28. Antagonism of Rad53 activity restores replication and viability of ptc2  pph3  cells.
WT +pGal (SSy385), +rad53-KD (SSy386); pph3  +pGal (SSy387), +rad53-KD (SSy388); ptc2 
+pGal (SSy389), +rad53-KD (SSy390); and ptc2 pph3  +pGal (SSy391); +rad53-KD (SSy392) cells
were blocked in G
1
-phase with -factor in YEP-Raffinose and released into YEP-Raffinose containing
0.033% MMS for 90 minutes at 23
o
C (MMS). MMS was washed out and cells were suspended in
YEP-Raffinose + 0.5% Galactose (Time = 0). (A) DNA content was analyzed at the indicated times.
(B) In situ Rad53 kinase assay of SSy391 and SSy392. (C) % Viability of SSy391 and SSy392 was
determined relative to the -factor time point; standard error for three independent experiments is
shown.

98

Activated Rad53 regulates replication fork restart
Rad53 plays a crucial role in the stabilization of replication forks encountering
DNA damage. The mechanism of stabilization remains vague, but is thought to
involve Rad53-dependent phosphorylation of replication proteins to maintain their
association with the fork and thereby prevent fork collapse and formation of dsDNA
breaks, which occurs in the absence of Rad53 (Branzei & Foiani, 2006). We exploited
the defect of pph3∆ cells in deactivating Rad53 to examine replication fork activity
under constant Rad53 control. The slower replication fork progression in MMS-
damaged pph3∆ cells strongly suggests that activated Rad53 directly impedes fork
progression or restart by phosphorylating replication factors. We further showed that
deactivation of Rad53 is sufficient to allow replication fork restart in cells recovering
from DNA damage. This finding links replication restart with dephosphorylation of
Rad53, which normally results from diminution of the checkpoint signal as DNA
damage is repaired. By inhibiting replication fork restart as well as additional origin
firing, Rad53 provides a better opportunity for repair of damaged DNA prior to arrival
of a replication fork.
Our conclusion that Rad53 directly regulates fork activity contrasts with the
conclusion of Tercero and Diffley who observed similarly slow replication fork
kinetics across the chromosome VI replicon in wild-type and rad53  cells in the
presence of MMS, leading them to conclude that while DNA damage slows replication
fork progression, Rad53 does not regulate the rate of fork progression in response to
DNA damage (Tercero & Diffley, 2001). Using BrdU-IP-chip, we also find that fork
rates are similar in wild-type and rad53∆ cells (Figure 29). Based on our conclusion
99

that deactivation of Rad53 is required for fork restart, a seemingly straightforward
prediction is that rad53∆ cells would exhibit more rapid fork progression in the
presence of MMS than wild-type cells, which is not observed. However, elimination
of Rad53 complicates the situation by the resulting failure to stabilize stalled forks.

100

Figure 29. WT and rad53  cells display similar fork rates in MMS. WT (SSy529) and rad53 
(SSy530) cells were arrested in G
1
-phase with factor at 23
o
C and released into YEPD containing
0.033% MMS at 23
o
C. Cells were collected at the indicated time points for DNA content analysis (A)
or incubated with BrdU for the indicated intervals and analyzed as described in Fig. 19 for the indicated
portions of chromosome VI (B) or chromosome III (C). ARS608 and ARS609 are not deleted in these
strains and some BrdU incorporation indicative of initiation of these origins is observed in (B), and the
normally inefficient origin ARS304 displays a significant level of BrdU incorporation in the rad53 
cells (C).
101

Figure 29.

102

Thus, we suggest (in agreement with Tercero and Diffley) that DNA damage
intrinsically slows or stalls fork progression independently of Rad53, perhaps by
provoking uncoupling of the fork from the damaged site and generating a DNA
damage signal (see below). Subsequent fork stabilization by Rad53 ensures that
stalled forks are channeled into an efficient, direct restart pathway, but only upon
deactivation of Rad53. In the case of rad53  cells, the initial failure to stabilize the
correct fork structure may preclude rapid and direct restart by the preferred
pathway(s), and rely instead on alternative restart mechanisms. Indeed, fork collapse
stimulates repair by homologous recombination, which is normally suppressed by
Rad53 (reviewed in Lambert et al, 2007). Ultimately, although fork progression
through the chromosome VI region occurs with approximately wild-type kinetics in
MMS-treated rad53  cells, unreplicated DNA remains, and these cells are inviable
(Tercero & Diffley, 2001). In contrast, pph3∆ cells maintain stable replication forks
in the presence of DNA damage and remain viable (Fig. 26A) (O'Neill et al, 2007).

A model of Rad53 regulation of replication fork restart
Current models for replication of a damaged template invoke uncoupling of
leading and lagging strand synthesis when leading strand synthesis is blocked by a
lesion, while a block of the lagging strand is automatically uncoupled as an Okazaki
fragment and does not block fork progression (reviewed in Heller & Marians, 2006).
Uncoupling allows continued template unwinding and lagging strand DNA synthesis
that enables replication restart beyond the blocking lesion through a re-priming event
on the leading strand template. As a result of uncoupling from the blocking lesions
103

and downstream re-priming of DNA synthesis, unreplicated gaps are left behind the
fork, presumably generating a DNA damage signal. These gaps can be repaired by
one of several post-replication repair (PRR) mechanisms, including recruitment of
translesion polymerase(s), template switching, or homologous recombination(Ulrich,
2005). However, some restart mechanisms such as fork regression by Rad5 and
translesion synthesis (TLS) may act directly in fork restart without the re-priming
mechanism, which remains speculative in eukaryotes. How the repair pathway is
chosen remains unclear, but likely depends on the nature of the lesion and whether
Rad53 stabilizes the uncoupled fork.
Rad53 may limit progression of the uncoupled fork by inhibiting helicase
activity or lagging strand synthesis to prevent the formation of long unreplicated gaps
(Figure 30). By limiting the extent of lagging strand synthesis, Rad53 may facilitate
Rad5-dependent fork regression as a restart mechanism, which may not act over very
long distances in vivo; although Rad5 can regress fork-like structures ~1 kb in vitro
(Blastyak et al, 2007), its activity may be constrained to lesions near the fork in vivo.
The Rad5 mechanism is error-free and bypasses the blocking lesion without producing
gaps in the DNA. Hence, it is interesting that MMS does not appear to provoke the
same level of gap formation, suggestive of fork uncoupling, as UV treatment (Sogo et
al, 2002), raising the possibility that Rad53 inhibits fork uncoupling in MMS, thereby
facilitating direct restart of stalled forks by Rad5. Different mechanisms may act
depending on the level of Rad53 activity in the cell when the fork encounters the
lesion. For example, early in S-phase before Rad53 has been activated, extensive fork
progression after uncoupling may occur, requiring re-priming and gap-repair, whereas
104

later in S-phase after Rad53 has become fully active, fork progression may be rapidly
halted after uncoupling, or uncoupling may be prevented to enable Rad5-dependent
restart.

Figure 30. A model of Rad53 regulation of replication fork restart. Arrows on the DNA structures
represent 3’ ends, the asterisks represent a fork-blocking lesion, and dashed lines indicate new DNA
synthesis. Thin curved lines represent protein function, ending with arrows for activation or
perpendicular lines for inhibition. See text for additional details of regulation.

Fork uncoupling may intrinsically regulate the rate of fork progression by
modulating helicase activity. Helicase rate and processivity are stimulated by coupled
polymerase activity (Dong et al, 1996; Hamdan et al, 2007; Kim et al, 1996; Stano et
al, 2005; Yuzhakov et al, 1996). Thus, uncoupling of a blocked leading strand
polymerase may enable a limited amount of continued unwinding and lagging strand
synthesis, which facilitates restart, while preventing excessive formation of
unreplicated gaps in the DNA. It is unclear whether Rad53 regulates fork coupling,
however, Mrc1, a Rad53-activating protein that functions at replication forks appears
105

to regulate this process, as delocalization of the replication apparatus from the site of
stalled DNA synthesis occurs in HU-treated mrc1 cells (Katou et al, 2003).
Interestingly, cells lacking Mrc1 exhibit significantly slower fork progression in the
absence of DNA damage consistent with the notion that fork uncoupling slows fork
progression (Hodgson et al, 2007; Szyjka et al, 2005; Tourriere et al, 2005). Mrc1
appears to be phosphorylated by activated Rad53 (Osborn & Elledge, 2003), which
may prevent uncoupling to limit ongoing replication of a heavily damaged template
while providing an opportunity for repair.
The regulation of Cdc7-Dbf4 may be a key feature of Rad53 regulation of
replication restart (Figure 30). Rad53-dependent inhibition of late origin firing is
thought to occur through phosphorylation of Dbf4, resulting in reduced Cdc7 kinase
activity and dissociation from chromatin (Duncker et al, 2002; Weinreich & Stillman,
1999). However, recent studies in human cancer cells suggest Cdc7 kinase is active
under replication stress conditions (Tenca et al, 2007). A possible explanation for this
apparent discrepancy is that Rad53 may inhibit Cdc7-Dbf4 activity in normal
replication initiation at origins while stimulating Cdc7-Dbf4 activity in replication
fork restart. Consistent with this idea, Cdc7-Dbf4 has been implicated in error-prone
TLS. Cdc7 is required for induced mutagenesis by UV and MMS, and combined
mutations of CDC7 and RAD5 cause synergistic increases in UV and MMS
sensitivities (Njagi & Kilbey, 1982; Pessoa-Brandao & Sclafani, 2004). This indicates
that Rad5 and Cdc7-Dbf4-dependent TLS function in parallel, and suggests that
Rad53 may regulate this choice (Figure 30). In S. pombe, mutations in hsk1 or dfp1
(CDC7 and DBF4 homologs, respectively) cause MMS sensitivity and chromosomal
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instability, consistent with a role in replication fork stabilization or restart, and Cds1
(RAD53 homolog) appears to directly target Hsk1-Dfp1 for regulation (Fung et al,
2002; Matsumoto et al, 2005; Snaith et al, 2000; Sommariva et al, 2005; Takeda et al,
1999; Takeda et al, 2001). Consistent with the idea that Cds1 controls the choice of
replication restart pathways is the finding that cds1 deletion exacerbates induced
mutagenesis resulting from a defective allele of DNA Polymerase , which likely
causes frequent fork stalling (Kai & Wang, 2003). Together, these findings suggest
that Cdc7-Dbf4 is required for replication restart through the TLS pathway, and that
Rad53 modulates this activity.

Distinct and overlapping roles of Rad53 phosphatases in checkpoint regulation
In constant MMS, replication forks progress more slowly in pph3∆ cells than
in wild-type cells (Fig. 19), suggesting that Pph3 dephosphorylation of Rad53
constitutively opposes Rad53 activation. Thus, the slow but continuous fork
progression in wild-type cells (in MMS) likely reflects a cycle of fork stabilization by
Rad53 and fork restart through Rad53 dephosphorylation. Complete Rad53
dephosphorylation may not be required if specific hyperphosphorylated form(s) inhibit
restart, or if a graded response to the level of Rad53 phosphorylation occurs. The data
support this idea, as replication restart appears to correlate with reduction in the level
of the slowest migrating forms of Rad53 (Fig. 26B, C). An attractive possibility for
regulation of Pph3 activity may be through its targeting to replication forks (by Psy2)
to dephosphorylate Rad53 only when these forks are prepared for restart, for example,
after lesion bypass by Rad5.
107

The progression of replication forks in MMS-treated pph3∆ cells slows but
does not arrest completely (Fig. 19A), indicating either that activated Rad53 does not
completely block fork progression or restart, or that an alternative means of Rad53
deactivation occurs in pph3∆ cells. Consistent with the latter hypothesis, reduced
levels of phosphorylated Rad53 eventually are observed in pph3∆ cells (Fig. 26B).
We tested the involvement of PTC2, which encodes a Rad53 phosphatase previously
characterized as having a role in recovery from cell cycle arrest caused by a long-
lived, dsDNA break (Leroy et al, 2003). We found that combined disruption of Pph3
and Ptc2 activity virtually eliminates replication fork restart and causes lethality after
MMS-induced damage, reinforcing the conclusion that Rad53 deactivation is the key
function of Pph3 in replication restart. Furthermore, we showed that direct
antagonism of activated Rad53 by overexpression of Rad53-KD enables replication
restart and rescues the lethality of pph3∆ ptc2∆ cells, which are otherwise unable to
deactivate Rad53. Based on these results, we conclude that Rad53 deactivation is
necessary for replication fork restart after damage-induced stalling and Rad53-
dependent fork stabilization.
Pph3 and Ptc2 appear to play differential roles in Rad53 deactivation
depending on the source of checkpoint activation. We find only a minor requirement
for Ptc2 in dephosphorylation of Rad53 and replication restart during recovery from
MMS, except in cells lacking Pph3 (Figs. 26 and 28). Ptc2 and Pph3 interact with
different domains of Rad53, and therefore, may preferentially dephosphorylate distinct
phospho-sites (Leroy et al, 2003; O'Neill et al, 2007). Specific phospho-sites may
regulate distinct aspects of Rad53 function, so the ability to deactivate specific sites
108

with a unique phosphatase potentially allows for modulation of the checkpoint
response for the specific circumstances (reviewed in Heideker et al, 2007). Along
these lines, Pph3 may prefer Rad53 phospho-sites that regulate replication fork restart
after MMS damage, while Ptc2 may favor sites that regulate G
2
-arrest and adaptation
after dsDNA break (Fig. 26B and Leroy et al, 2003; O'Neill et al, 2007). Furthermore,
neither Pph3, nor Ptc2, nor Ptc3, alone or in combination, is required for viability after
HU treatment suggesting that Rad53 phospho-sites may be differentially targeted in
response to different types of replication stress (Fig. 31). Similar findings were
reported while this paper was in revision (Travesa et al, 2008). Obviously,
determining the Rad53 phospho-site specificities of Pph3 and Ptc2 will be highly
illuminating. Little is known about the regulation of these phosphatases in DNA
damage responses, knowledge of which will also be crucial to understanding fully
checkpoint recovery.

Figure 31. Neither Pph3 nor Ptc2 is required for viability of HU-treated cells. WT (SSy187),
pph3  (SSy188), ptc2  (SSy248), ptc2  pph3  (SSy249) ptc3  ptc2  pph3  (SSy405), and ptc3 
ptc2  (SSy411) cells were arrested in G
1
-phase with -factor at 30
o
C and released into YEPD
containing 0.2 M HU at 30
o
C. At the indicated times cells were ten-fold serially diluted and plated onto
YEPD. Plates were incubated two days at 30
o
C and imaged.

109

CHAPTER 5

Conclusions

Mrc1 is required for normal progression of replication forks in Saccharomyces
cerevisiae

In chapter 2, we demonstrated that in the absence of Mrc1, replication forks
proceed slowly throughout chromatin. This was not a result of increased fork pausing
at tRNAs or decreased origin firing. In fact, we showed that early origin firing is not
altered in mrc1  cells and that ARS304, a normally late and inefficient origin, fires in
mrc1 cells. This is likely a result of slow fork progression in mrc1  cells, which
allows time for the firing of normally dormant origins. Also, contrary to a previous
report, we demonstrated that rrm3 cells do not rely on Mrc1’s checkpoint function
for survival.
Mrc1 localizes to origins around the time of initiation, coimmunoprecipitates
with Mcms and travels with replication forks. During an unperturbed S phase, mrc1 
cells display increased recombination and an active DNA damage checkpoint. When
mrc1 cells are exposed to HU, BrdU incorporation halts while Cdc45 proceeds ~2.5-
3kb further along the chromosome. This data suggests a role for Mrc1 in replisome
coupling during response to HU treatment. Based on our data and the data
mentioned above, we propose that Mrc1 also plays a role in fork coupling during an
unperturbed S phase. Our model suggests that in mrc1 cells the helicase unwinds
ahead of leading and lagging strand polymerases. Unwinding that occurs ahead of the
polymerases exposes ssDNA, which, at a certain threshold, triggers the DNA damage
response. Consequently, exposure of ssDNA renders the fork susceptible to strand
110

breaks and recombination. Indeed, these phenotypes are observed in mrc1  cells.
Furthermore, we suggest that uncoupled unwinding and synthesis is the reason for
reduced fork rate in mrc1  cells, which is in line with previous studies in bacteria that
have demonstrated coupling between the helicase and polymerases facilitates efficient
replication (see Figure 32 for details). At this point, it is unclear whether the
polymerases and helicases move along the chromosome at a constant (“slow”)
distance from each other or if the polymerase and helicase play “molecular bumper
cars” along the chromosome where they associate for a short time and then dissociate
due to the lack of Mrc1-dependent coupling.

Figure 32. A model for DNA replication in mrc1  cells. In wild-type cells (A), the helicase unwinds
double stranded DNA which is then copied by leading and lagging polymerases. Efficient unwinding
and copying is ensured through Mrc1-dependent coupling. In mrc1  cells (B), association between the
helicase and polymerase is lost resulting in decreased fork progression. This also results in extended
tracts of RPA-bound ssDNA, which triggers the DNA damage response. See text above for details.
111

To more specifically elucidate Mrc1’s role at the replication fork, I propose the
following experiments:

I. Examine RPA binding in WT, mrc1-AQ and mrc1  cells
Based on our prediction, if Mrc1 couples polymerases and helicases at the
replication fork, mrc1  cells should have increased amounts of ssDNA. Since ssDNA
at replication forks is bound by RPA, mrc1  cells should have increased amount of
RPA at replication forks. I propose an experiment that utilizes RPA-ChIP-chip, BrdU-
IP-chip and NimbleGen tiling arrays. Exposing cells to short pulses of BrdU, and
analyzing RPA binding by ChIP-chip we can obtain a snapshot of active replication
forks and compare it to regions of RPA binding. In theory, this experiment could be
carried out using DNA combing and antibodies for RPA and BrdU to obtain lengths of
RPA tracts in our three strains.

II. Ascertain the domain(s) of Mrc1 responsible for Mcm and polymerase association.
I propose truncation experiments that examine the regions of Mrc1 and regions
of Mcms (and perhaps pol    in a yeast-2-hybrid study aimed at elucidating the
specific region(s) of Mrc1 responsible for interactions with replication fork proteins to
maintain coupling. Following identification of these regions, I think it would be
informative to introduce an allele lacking one or all of the interacting domains into an
mrc1 cell and analyze: association with the replication fork, bulk DNA synthesis,
Rad53 activation during HU exposure, and fork restart after HU exposure.
112

Additionally, these alleles could be tested to see if they rescue the lethality of an
mrc1 rrm3 cell.

III. Examine replication intermediates in mrc1 rrm3
ts
strains.
The reason for synthetic lethality of mrc1  rrm3  cells is still in question. In
Chapter 2, we proposed that mrc1 cells rely on the Rrm3’s helicase activity to
resolve problems at the replication fork. Indeed, a recent study has shown that
replication forks in mrc1  cells pause due to hairpin intermediates that accumulate at
inverted repeats (Voineagu et al, 2008). Likely, hairpins are a result of extended
ssDNA regions that anneal back on themselves in mrc1 cells. Situations like these
may require the activity of the Rrm3 helicase and allow for progression of replication
forks. Potentially, in the absence of both Mrc1 and Rrm3, these replication forks may
irreversibly stall. Additionally, mrc1 rrm3  cells may experience a lethal amount of
recombination, fork break down and fork stalling at endogenous pause sites. To get a
better picture of what happens at the replication fork in these cells, I have constructed
and begun to characterize a set of temperature sensitive Rrm3 alleles in an mrc1  cell.
Using these strains, we can examine replication intermediates at endogenous pause
sites, inverted repeats, and fork progression, which will give us a clearer picture of the
catastrophic events that occur at the replication fork in mrc1 rrm3 cells.

113

Rad53 regulates replication fork restart after DNA damage in Saccharomyces
cerevisiae

In Chapters 3 and 4, we demonstrated through genetic and biochemical
experiments that the Psy2-Pph3 phosphatase complex is responsible for
dephosphorylating Rad53 after an MMS-induced DNA damage response. In the
absence of Psy2 or Pph3, hyperphosphorylated Rad53 persists during recovery and
replication forks, though stable, are delayed in their restart. Using BrdU-incorporation
and oligonucleotide microarrays (BrdU-IP-chip), we demonstrated that antagonism of
Rad53 activity, through the use of the dominant-negative rad53-KD allele, is sufficient
to promote replication fork restart. Interestingly, in the absence of Pph3, Rad53 is
eventually dephosphorylated. This led us to examine another Rad53 phosphatase,
Ptc2. In ptc2  pph3  cells recovering from a DNA damage response, Rad53 is not
dephosphorylated, replication forks arrest and cells die. Both fork restart and viability
can be restored in ptc2  pph3 cells when Rad53 is antagonized, which demonstrates
that decreased viability is not due to mass amounts of fork collapse, but instead is a
result of a Rad53-dependent cell cycle block. These experiments have begun to shed
light on the topic of checkpoint-dependent fork slowing and have led to the following
model:

114

Figure 33. Pph3 and Ptc2 dephosphorylate Rad53 during recovery from DNA damage. In wild
type cells recovering from a DNA damage response (A), the Pph3 phosphatase dephosphorylates
Rad53, which promotes replication restart and cell cycle recovery. In pph3  cells recovering from a
DNA damage response (B), hyperphosphorylated Rad53 persists, which is eventually
dephosphorylated, perhaps by Ptc2. These cells are characterized by slowed replication fork restart and
delayed completion of S phase. Interestingly, pph3  cells recovering from DNA damage also fire the
late origin, ARS603, despite the presence of hyperphosphorylated Rad53. Perhaps Ptc2
dephosphorylates residues of Rad53 responsible for restraining late origin firing. In ptc2  pph3  cells
recovering from a DNA damage response (C), Rad53 remains hyperphosphorylated, resulting in
replication fork arrest, cycle arrest and cell death. Curved arrows represent Rad53 dephosphorylation
by the respective phosphatase. Horizontal arrows between forms of Rad53 represent relative length of
time.

In addition to the proposed model, I propose the following to obtain help further
elucidate Rad53’s at the replication fork during a checkpoint response:

115

I. Determine the specific phosphorylation sites (if any) that Pph3 or Ptc2
dephosphorylate during recovery from a DNA damage response. Determine if the
different sites control different aspects of replication (i.e. origin firing and fork
slowing).
In pph3 cells recovering from a DNA damage response, the late origin,
ARS603 fires despite the presence of hyperphosphorylated Rad53. In ptc2  cells
recovering from a DNA damage response, replication forks restart with wild type
kinetics. However, in a ptc2  pph3 cell recovering from DNA damage, late origin
firing, replication fork restart and cell cycle progression are inhibited resulting in cell
death. Therefore, I hypothesize that in a pph3 cell, Ptc2 may partially
dephosphorylate Rad53, leading to late origin firing. Similarly, in a ptc2  cell, Pph3
may act to dephosphorylate Rad53 on specific residues resulting in efficient fork
restart (see Fig 33). To address this hypothesis, I propose experiments using inducible
PTC2 or PPH3 genes in a ptc2  pph3 background. By inducing Ptc2 or Pph3 during
the recovery period, and analyzing fork restart and late origin firing, we can obtain a
clearer picture as to specific roles for each phosphatase during DNA damage recovery.
In addition, using mass spectrophotometry we can analyze specific Rad53 residues
that are dephosphorylated in ptc2  or pph3  during MMS recovery and
mechanistically link specific residues to their function during a checkpoint response.

116

II. Determine the phosphatases responsible for Rad53 deactivation in HU.
The phenotypes observed for pph3, ptc2 and ptc2  pph3  cells are
specific to MMS-induced DNA damage. When these cells recover from HU exposure,
Rad53 is dephosphorylated, replication forks restart with similar kinetics to WT cells,
and cellular viability is relatively uncompromised. Perhaps there are other
phosphatases that are responsible for Rad53 dephosphorylation during an HU
response. As mentioned in Keogh et al 2006, there are nine nuclear phosphatases.
Therefore, I propose a screen of these phosphatases for HU sensitivity and Rad53
dephosphorylation kinetics. This screen could also be carried out for UV, NQO and
other DNA damaging agents. Taken together, experiments such as these can elucidate
the how the cell may differentiate amongst different forms of damage and trigger the
appropriate pathway for recovery and repair.

117

CHAPTER 6
Materials and Methods

Plasmid and strain constructions
All strains are derived from W303, except where noted, and are listed in Tables
1-3. Gene knockouts were constructed by PCR-based methods (Guldener et al, 1996;
Longtine et al, 1998). pRS306-Mrc1-AQ (Chapter 2) contains the NotI-KpnI mrc1-
AQ fragment from pRS405-Mrc1-AQ (Gibson et al, 2004) inserted into pRS306.
mrc1-AQ was introduced at its native locus by a pop-in pop-out strategy, or by a
simple pop-in (Chapter 2). To construct pGal-rad53-KD (Chapter 4), the 1.6kb
BstEII-SphI fragment of RAD53 (SPK1) containing the kinase-inactivating mutations
K227A and D339A was isolated from pRS316-SPK1-(K227A,D339A) (Fay et al,
1997), and subcloned into BstEII-SphI digested pJA98 (Allen et al, 1994). To
construct pGal-PPH3 (Chapter 4), genomic DNA from SSy250 (P
GAL
-HA::PPH3) was
amplified with a primer (containing an SphI site) upstream of the GAL promoter and a
primer just downstream of an endogenous MscI site within the PPH3 gene. The PCR
product was digested with SphI and MscI and ligated into pFR071 (pRS306-PPH3;
O'Neill et al 2007). To construct p415-rrm3
ts
, pAI20 (Ivessa et al, 2003) was digested
with PstI and HindIII to liberate a 3.2kb fragment containing the entire RRM3 gene.
This fragment was cloned into pRS415, and subsequently digested with NsiI resulting
in a gapped plasmid that was gel purified. To introduce random mutations into the
RRM3 gene, RRM3 was amplified under the following conditions: 5U Taq
Polymerase, dNTPs (dCTP, dGTP, dGTP: 1mM, dATP: 0.2mM), 4.2mM MgSO
4
,
118

0.5mM MnCl
2
, Vent Buffer (New England Biolabs), and 100ng RRM3 template DNA
(pIA20). 25 cycles of: 94
o
C (1.5min), 45
o
C (1min) and 72
o
C (2.5min) yielded
RRM3
ts
DNA. Gel-purified gapped plasmid plus column purified RRM3
ts
DNA was
combined in a 6:1 (vector:insert) ratio and transformed into SSy203. Transformants
were grown on selective media at 23
o
C, patched to FOA at 23
o
C (to select for loss of
pRS316-MRC1), and replica plated to two YEPD plates; 1 placed at 23
o
C, 1 placed at
37
o
C. Patches that did not grow 37
o
C were selected and analyzed further.

Yeast Methods
YEPD medium was used for all experiments, except where noted. Raffinose
was present at 2% in YEP-Raffinose. Cell culturing, synchronization, DNA content
analysis (FACScan), spore analysis, and Rad53 analysis have been described
(Aparicio et al, 2004; Gibson et al, 2004), except we used anti-Rad53 antibody at
1:1000 (Santa Cruz Biotechnology, SC6749). 22 cycles of PCR were performed for
analysis of the ChIP experiment (Chapter 2). The in situ Rad53 kinase assay has been
described previously (Pellicioli et al, 1999). For DNA damage recovery experiments,
MMS (Sigma) was quenched by addition of sodium thiosulfate to 0.5%, cells were
immediately harvested by centrifugation and resuspended as indicated in the figure
legends. Budding index was determined by counting 200 cells per time point. Colony
survival assays (Chapter 3) were performed by counting colony forming units (cfu)
three days after deposition onto YPD-agar media containing the indicated
concentration of MMS. Each point represents the mean of at least three independent
experiments. Error bars represent standard deviations of the independent experiments.
119

The Matchmaker yeast two-hybrid system (Clontech) was employed to analyze Psy2-
Rad53 binding interactions (Chapter 3).

Analysis of replication structures (Chapters 2 and 3).
2D gel analysis was performed as described previously, except that 30 g of
DNA was used for each sample (Gibson et al, 2004). Quantitative comparisons were
performed on samples run in the same gels and blotted and hybridized together to
minimize any errors that might result from differential transfer or hybridization
efficiencies.

Analysis of BrdU incorporation (Chapter 2).
Strains expressing seven copies of HSV-TK were grown in the presence of 400
g/mL BrdU as described (Lengronne et al, 2001). Immunoprecipitation of BrdU-
substituted DNA was carried out as described in (Katou et al, 2003) and references
therein, with the following modifications: DNA was extracted by glass bead beating,
sheared by sonication to an average size of ~500 bp, and isolated by
phenol/chloroform extraction and ethanol precipitation. DNA was treated with
RNAseA and Proteinase K and further purified on Qiaquick PCR purification spin-
columns (Qiagen). 500 ng of DNA was combined with 20 g of sheared salmon
sperm DNA, denatured, and incubated with anti-BrdU antibody (Oxford
Biotechnology) followed by G-Sepharose beads (Amersham). 22 cycles of PCR were
performed on the immunoprecipitated material and on a 1:10 dilution of the input
material.
120

Viability analysis (Chapter 4).
Approximately 7.5  10
6
cells were removed from culture, sonicated, ten-fold
serially diluted, plated onto YEPD and incubated at 30
o
C for 2 days. Plates were
imaged using a ChemiDoc XRS 170-8070 (Bio-Rad) and Quantity One Analysis
software (Bio-Rad). Quantification of viability was performed by duplicate platings
onto YEPD of appropriate culture dilutions that result in 100-200 colonies per plate.

BrdU-IP-chip (Chapter 4).
Strains containing the BrdU-Inc construct (Viggiani & Aparicio, 2006) were
incubated with 800 g/ml BrdU (Sigma) and harvested with the addition of ice-cold
TBS and 0.1% NaN
3
. DNA isolation and BrdU immunoprecipitation were performed
as previously described (Szyjka et al, 2005), except that anti-BrdU (GE Healthcare)
was used at 1:400 and incubated overnight at 4
o
C. To obtain a reference “total DNA”
sample, DNA was isolated from a G
1
-arrested culture. Amplification of
immunoprecipitated and total DNA was performed as described (O'Geen et al, 2006),
except that amplified samples were subjected to Klenow extension for 4 hours at 37
o
C
in the presence of amino-allyl-dUTP (48 M, Ambion) and dNTPs (dATP, dGTP,
dCTP: 120 M, dTTP: 72µM). Also spiked into each amplification reaction was 0.25
µL of ten Drosophila cDNA clones (corresponding to Drosophila oligonucleotides on
the mircoarrays, see below) that span a 100-fold concentration range (1-100 pg/µL),
which produce a range of signal intensities for data normalization.
Immunoprecipitated and total DNA samples were coupled with Cy5 or Cy3 dyes (GE
Healthcare) at room temperature for 1 hour, respectively. After purifying DNA
121

samples using Qiaquick spin columns (Qiagen), 1 g total DNA and 1 g of each
immunoprecipitated sample were combined, dried in a speedvac (Thermo),
resuspended in 10mM EDTA, and denatured for 2 minutes. Pre-warmed (50
o
C)
hybridization buffer (30% formamide, 5x SSC, 0.1% SDS, 100 g/ml salmon sperm
DNA) was added to denatured samples and hybridized to pre-warmed microarrays for
18 hours at 50
o
C; one slide was used for each experimental time point. Slides were
washed with gentle shaking in 1x SSC, 0.1% SDS, 1mM DTT (pre-warmed to 50
o
C)
for 5 minutes, submerged several times in 0.2x SSC, 1mM DTT (23
o
C), and washed
two times for 3 minutes in 0.1x SSC, 1mM DTT (23
o
C). Slides were dried by
centrifugation for 45 seconds in a microfuge and scanned in an Axon scanner using
Genepix 5.0 to capture and save the image.

Microarray design and production (Chapter 4).
Oligonucleotide probes (60 bp, Tm range = 80-90°C) were designed to analyze
chromosome VI (coordinates: 143,000-270,000; one probe per 270 bp) and part of the
left arm of chromosome III (coordinates: 11,300-77,700; one probe per 100 bp) using
OligoArray 2.1 (Rouillard et al, 2003). 50 oligonucleotides (five unique
oligonucleotides for each of the ten Drosophila cDNA clones described above) were
similarly designed to detect the spiked-in Drosophila cDNAs for data normalization.
Oligonucleotides were printed in quadruplicate on poly-lysine slides (Erie Scientific)
using either a MicroGrid I BioRobotics or MGuide printer
(http://cmgm.stanford.edu/pbrown/mguide/).
122

Post-processing utilized succinic anhydride as blocking reagent as described in
(http://cmgm.stanford.edu/pbrown/protocols/3_post_process.html).

Microarray analysis (Chapter 4)
ImaGene image analysis software (http://www.biodiscovery.com/index/imagene)
was used to quantify raw spot intensities, and low-level analysis of the arrays was
performed using the limma software (Smyth, 2004), available from Bioconductor and
R (R Development Core Team, 2007). Local background correction was applied, and
normalization was performed using the Drosophila spike-in probes on the arrays.

123

Table 1
Strains used in Chapter 2
Strain Genotype Source
Strains are identical to W303-1a, except as indicated
W303-1a MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 R. Rothstein
can1-100
YJT80 ars608 ::HIS3 ars609 ::TRP1 ars305 ::kanMX (Tercero & Diffley, 2001)
ADE2 rad5-535
E1000 GPD-TK(7x)::URA3 (Lengronne et al, 2001)
SSy45 YTJ80 ars305 ::loxP (Szyjka et al, 2005)
SSy46 SSy45 mrc1 ::kanMX “
SSy47 SSy46 bar1 ::LEU2 “
SSy48 SSy45 bar1 ::LEU2 “
SSy56 SSy46 bar1 ::URA3 “
SSy62 SSy56 mrc1 ::kanMX::mrc1-AQ (LEU2) “
SSy69 MAT  rrm3 ::TRP1 RAD5 “
SSy72 SSy185 mrc1 ::HIS5::mrc1-AQ (LEU2) RAD5 “
SSy81 mrc1 ::HIS5::mrc1-AQ (LEU2) rrm3 ::TRP1 (SSy108 segregant) “
SSy108 SSy69 X SSy72 diploid “
SSy109 SSy69 X SSy185 diploid “
SSy116 SSy47 mrc1-AQ “
SSy185 mrc1 ::HIS5 RAD5 “
CVy29 E1000 mrc1 ::kanMX ars608 ::HIS3 bar1 ::LEU2 “
CVy30 CVy31 mrc1 ::kanMX “
CVy31 E1000 bar1 ::LEU2 cdc45 ::CDC45-HA3 (TRP1) “
rad5-535
CVy39 E1000 ars608 ::HIS3 bar1 ::TRP1 “
CVy40 CVy39 mrc1 ::HIS5::mrc1-AQ (LEU2) “

124

Table 2

Strains used in Chapter 3

Strain Genotype Source
BY4741 MATa his3∆1 leu2∆0 met15∆0 ura3∆0 ATCC
orf  BY4741 orf ::kanMX4 Open Biosystems
FR288 BY4741 psy2 ::LEU2 (Kolodner et al, 2002)
FR319 BY4741 tof1::LEU2 “
FR818 BY4741 rad9∆::LEU2 (O'Neill et al, 2007)
FR334 BY4741 psy2∆::kanMX4 tof1∆::LEU2 (Kolodner et al, 2002)
FR293 BY4741 rad9∆::kanMX4 psy2∆::LEU2 (O'Neill et al, 2007)
FR583 BY4741 rad24∆::kanMX4 psy2∆::LEU2 “
FR932 BY4741 rad17∆::kanMX4 psy2∆::LEU2 “
FR494 BY4741 mrc1∆::kanMX4 psy2∆::LEU2 “
FR1098 BY4741 ptc2∆::kanMX4 psy2∆::LEU2 “
FR1046 BY4741 pph3∆::his5
+
“
FR962 BY4741 pph3∆::kanMX4 psy2∆::LEU2 “
FR984 BY4741 pph3∆::kanMX4 tof1∆::LEU2 “
FR986 BY4741 pph3∆::kanMX4 rad9∆::LEU2 “
FR1135 BY4741 rad24∆::kanMX4 pph3∆::his5
+
“
FR1133 BY4741 rad17∆::kanMX4 pph3∆::his5
+
“
FR1048 BY4741 mrc1∆::kanMX4 pph3∆:: his5
+
“
FR1100 BY4741 ptc2∆::kanMX4 pph3∆:: his5
+
“
FR1195 BY4741 ybl046w∆::his5
+
“
FR964 BY4741 ybl046w∆::kanMX4 psy2∆::LEU2 “
FR988 BY4741 ybl046w∆::kanMX4 tof1∆::LEU2 “
FR990 BY4741 ybl046w∆::kanMX4 rad9∆::LEU2 “
FR1235 BY4741 ptc2∆::kanMX4 ybl046w∆::his5
+
“
RDKY3615 MATa ura3-52 leu2∆1 trp1∆63 his3∆200 lys2∆Bgl (Nyberg et al, 2002)
hom3-10 ade2∆1 ade8 hxt13::URA3
RDKY3733 RDKY3615 sml1∆::G418 “
RDKY3735 RDKY3615 sml1∆::G418 mec1∆::HIS3 “
RDKY3739 RDKY3615 dun1∆::HIS3 “
125

Table 2, continued

Strain Genotype Source
RDKY3749 RDKY3615 sml1∆::G418 rad53∆::HIS3 (Nyberg et al, 2002)
FR623 RDKY3615 psy2∆::LEU2 (O'Neill et al, 2007)
FR1057 RDKY3615 sml1∆::G418 psy2∆::TRP1 “
FR625 RDKY3615 sml1∆::G418 mec1∆::HIS3 psy2∆::LEU2 “
FR980 RDKY3615 dun1∆::HIS3 psy2∆::LEU2 “
FR982 RDKY3615 sml1∆::G418 rad53∆::HIS3 psy2∆::LEU2 “
FR1028 RDKY3615 pph3∆::TRP1 “
FR1059 RDKY3615 sml1∆::G418 pph3∆::TRP1 “
FR1030 RDKY3615 sml1∆::G418 mec1∆::HIS3 pph3∆::TRP1 “
FR1032 RDKY3615 dun1∆::HIS3 pph3∆::TRP1 “
FR1034 RDKY3615 sml1∆::G418 rad53∆::HIS3 pph3∆::TRP1 “
FR1205 RDKY3615 ybl046w∆::TRP1 “
FR1207 RDKY3615 sml1∆::G418 ybl046w∆::TRP1 “
FR1209 RDKY3615 sml1∆::G418 mec1∆::HIS3 ybl046w∆::TRP1 “
FR1211 RDKY3615 dun1∆::HIS3 ybl046w∆::TRP1 “
FR1213 RDKY3615 sml1∆::G418 rad53∆::HIS3 ybl046w∆::TRP1 “
W1588-4C MATa ade2-1 can1-100 his3-11,15 leu2-3,112 (Lambert & Carr, 2005)
trp1-1 ura3-1
FR796 W1588-4C PSY2-TAP::kan
r
(O'Neill et al, 2007)
FR1038 W1588-4C PPH3-TAP::kan
r
“
FR946 W1588-4C PPH3-13MYC::TRP1 “
FR950 W1588-4C YBL046W-13MYC::TRP1 “
FR948 W1588-4C PSY2-TAP::kan
r
PPH3-13MYC::TRP1 “
FR952 W1588-4C PSY2-TAP::kan
r
YBL046W-13MYC::TRP1 “
FR1044 W1588-4C PSY2-TAP::kan
r
pph3∆::his5
+
“
FR1180 W1588-4C Rad53-TAP::kan
r
psy2∆::LEU2 pph3∆::his5
+
“
SSy187 MATa ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 “
ura3-1 RAD5 bar1::hisG ars603.5::URA3
SSy188 SSy187 pph3∆::his5
+
“
SSy189 SSy187 psy2∆::his5
+
“
SSy235 SSy187 ybl046w∆::his5
+
“
126

Table 2, continued

Strain Genotype Source
SSy210 SSy205 pph3∆::his5
+
(O'Neill et al, 2007)
SSy205 MATa ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 “
ura3-1 RAD5 bar1::hisG ars603.5∆::URA3
ars608∆::HIS3 ars609∆::TRP1 LEU2::BrdU-Inc.
DD355 MATa∆::hisG ho∆ hml∆::ADE1 hmr∆::ADE1 (Keogh et al, 2006)
(YMV2) his4::URA3-leu2(XhoI-to-Asp718)-pBR322-his4
ade1 lys5 ura3-52 trp1::hisG leu2::HOcs ade3::GAL::HO
DD813 DD355 pph3∆::KanMX (Ma et al, 2006)
DD983 DD355 hta2∆::KanMX hta1-S129A “
DD1039 DD355 hta2∆::KanMX hta1-S129A pph3∆::NatMX “

127

Table 3

Strains used in Chapter 4

Strain Genotype Source
All strains are bar1 ::hisG RAD5 derivatives of W303-1a unless noted

W303-1a MATa ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 R. Rothstein
can1-100
232178 sml1 ::HIS3 BAR1 “
SSy187 ars603.5 ::URA3 (O'Neill et al, 2007)
SSy188 ars603.5 ::URA3 pph3 ::HIS5 “
SSy248 ars603.5 ::URA3 ptc2 ::KanMX (Szyjka et al, 2008)
SSy249 ars603.5 ::URA3 pph3 ::HIS5 ptc2 ::KanMX “
SSy250 ars603.5 ::URA3 KanMX-P
GAL1
-HA::PPH3 “
SSy385 ars608 ::HIS3 ars609 ::TRP1 leu2::BrdU –Inc (LEU2) “
pGAL (URA3)
SSy386 ars608 ::HIS3 ars609 ::TRP1 leu2::BrdU –Inc (LEU2) “
pGAL-rad53-KD (URA3)
SSy387 ars609 ::TRP1 pph3 ::HIS5 leu2::BrdU –Inc (LEU2) “
pGAL (URA3)
SSy388 ars609 ::TRP1 pph3 ::HIS5 leu2::BrdU –Inc (LEU2) “
pGAL-rad53-KD (URA3)
SSy389 ptc2 ::KanMX pGAL (URA3) “
SSy390 ptc2 ::KanMX pGAL-rad53-KD (URA3) “
SSy391 pph3::HIS5 ptc2 ::KanMX pGAL (URA3) “
SSy392 pph3::HIS5 ptc2 ::KanMX pGAL-rad53-KD (URA3) “
SSy395 ars608 ::HIS3 ars609 ::TRP1 leu2::BrdU –Inc (LEU2) “
pph3::lox-KanMX-lox pGAL (URA3)
SSy396 ars608 ::HIS3 ars609 ::TRP1 leu2::BrdU –Inc (LEU2) “
pph3::lox-KanMX-lox pGAL-rad53-KD (URA3)
SSy405 pph3::HIS5 ptc3 ::HIS5 ptc2 ::KanMX “
SSy411 pph3::HIS5 ptc3 ::HIS5 “
128

Table 3, continued

Strain Genotype Source
SSy419 ars608 ::HIS3 ars609 ::TRP1 ars305 ::loxP (Szyjka et al, 2008)
leu2::BrdU –Inc (LEU2)
SSy420 ars608 ::HIS3 ars609 ::TRP1 ars305 ::loxP “
leu2::BrdU –Inc (LEU2) pph3 ::loxP
SSy526 ars608 ::HIS3 ars609 ::TRP1 ars305 ::loxP “
leu2::BrdU –Inc (LEU2) pph3 ::loxP ptc2 ::loxP pGAL (URA3)
SSy527 ars608 ::HIS3 ars609 ::TRP1 ars305 ::loxP “
leu2::BrdU –Inc (LEU2) pph3 ::loxP ptc2 ::loxP
pGAL-PPH3 (URA3)
SSy529 sml1 ::HIS3 bar1 ::LEU2 ars305 ::BrdU-Inc (URA3) “
SSy530 sml1 ::HIS3 bar1 ::LEU2 ars305 ::BrdU-Inc (URA3) “
rad53 ::KanMX

129

Table 4

Strains used in Appendices

Strain Genotype Source
All strains are derivatives of W303-1a unless noted

SSy187 bar1 ::hisG ars603.5::URA3 RAD5 (O’Neill et al 2007)
SSy188 bar1 ::hisG ars603.5::URA3 pph3::HIS5 RAD5 “
SSy203 mrc1 ::HIS5 rrm3 ::TRP1 pRS316-MRC1 RAD5 This study
SSy224 bar1 ::URA3 cdc7-4 RAD5 “
SSy248 ars603.5 ::URA3 ptc2 ::KanMX (Szyjka et al, 2008)
SSy249 ars603.5 ::URA3 pph3 ::HIS5 ptc2 ::KanMX “
SSy325 mrc1 ::HIS5 rrm3 ::TRP1 RAD5 pRS415-rrm3
ts
(1) “
SSy326 mrc1 ::HIS5 rrm3 ::TRP1 RAD5 pRS415-rrm3
ts
(2) “
SSy327 mrc1 ::HIS5 rrm3 ::TRP1 RAD5 pRS415-rrm3
ts
(3) “
SSy328 mrc1 ::HIS5 rrm3 ::TRP1 RAD5 pRS415-rrm3
ts
(4) “
DGy368 bar1 ::hisG ars603.5::TRP1 rad5-535 D. Gibson
FR1162 bar1 ::hisG ars603.5::TRP1 pph3D::KanMX rad5-535 F. Romesberg
ROy1945 MATa lys2 ::HisG bar1 ::LEU2 RAD5? (Dhillon et al, 2006)
BUy828 MATa lys2 ::HisG bar1 ::LEU2 RAD5? htz1 ::KanMX “

130

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144

Appendix A: Fluorescence Activated Cell Sorting (FACS) measures DNA
content

Figure 34. FACS analysis measures DNA content. (A) Cells were collected from an asynchronous
population, fixed with ethanol, treated with RNAse A, Proteinase-K, and stained with Sytox Green, a
fluorescent DNA-binding dye. Two main peaks depict cells in G
1
(1C) or G
2
/M (2C) phases of the cell
cycle. Cells that contain greater than 1C and less than 2C are in S phase, which is denoted
(approximately) by the blue bracket. (B) A population of cells was synchronized in late-G
1
phase
(large peak at 1C), time = 0. Cells were then released into S phase and proceed until they reach 2C
DNA content. Based on this type of analysis, one can determine approximately how long a given strain
takes to progress through S phase.

145

Appendix B: Two-dimensional gel electrophoresis analyzes replication
intermediates

Figure 35. 2D gels examine replication intermediates at specific loci. (A) Cartoon representation
of a hypothetical chromosome. Replication forks emanate from an “origin” and progress
bidirectionally. Replication intermediates at the origin and the “+20kb” locus can be analyzed by
digesting chromosomal DNA with EcoRI. Brackets and arrows below EcoRI restriction sites depict
expected replication intermediates at respective loci. Digested DNA is run in first dimension, which
separates based on size. Lanes are cut from original gel, repositioned 90
o
and run in second dimension,
which separates based on shape. Gels are blotted and probed (location of probes are denoted by thick
black bars within restriction fragments). (B) Loci that are “actively” replicated (i.e. origin firing within
locus) contain small, medium, and large bubbles which create a pattern termed “bubble arc”. Loci that
are “passively” replicated contain small, medium and large forks (pictured on right). Arrow heads
denote fastest migrating replication intermediates within respective dimensions.

146

Appendix C: rad5-535 cells are defective for replication fork recovery

Figure 36. Cells containing the rad5-535 allele are defective for replication restart after MMS
exposure. WT (SSy187), pph3  (SSy188), rad5-535 (DGy368) and rad5-535 pph3  (FR1162) cells
were arrested in late G
1
with -factor for 2 hours at 30
o
C and released into 30
o
C YEPD containing
0.033% MMS for 60min. After which, MMS was quenched, cells were pelleted and resuspended in
30
o
C YEPD. Samples for DNA content analysis were collected at the indicated time points. Consistent
with Figure 10, bulk synthesis in pph3  cells lags behind WT cells (~45min). Interestingly, this
difference is masked when the rad5-535 allele is introduced. In the absence of an error free pathway,
error prone repair is required, which is probably less optimal (from a mutagenic standpoint), and
possibly slower than error free repair. Together this data suggests that error free repair plays an
important role in replication and that defects in DNA repair delay completion of DNA synthesis.

147

Appendix D: Deactivation of Cdc7 delays completion of DNA replication

Figure 37. Strains containing the temperature sensitive cdc7-4 allele are delayed in completion of
bulk DNA synthesis in unperturbed conditions and during recovery from MMS. (A) WT
(SSy187) and cdc7-4 (SSy224) cells were arrested with -factor and released into YEPD at 23
o
C. After
allowing time for early origin firing, half of each culture was shifted to 34
o
C (restrictive temperature,
red time points). Samples were collected at the indicated times for DNA content analysis. (B) Strains
from (A) were arrested in late G
1
with -factor and released into YEPD containing 0.033% MMS at
23
o
C for 15min. Cultures were shifted to 34
o
C for 45min while still in MMS (total time in MMS =
60min). After which, MMS was quenched, cells were pelleted and resuspended in 23
o
C YEPD, which
was then allowed to “warm up” to 34
o
C. Samples were collected at the indicated times for DNA
content analysis. Cdc7 is a key component of DDK and is required for origin initiation. The replication
defect of cdc7-4 cells after temperature shift (A) likely reflects faulty origin initiation. Completion of
replication is delayed >2.5hrs during MMS recovery compared to WT cells (B). Potentially, Cdc7
could be required for fork stabilization or fork restart during recovery from MMS insult. However, I
should note that Cdc7 was inactivated 15 minutes after -factor release. Possibly, early inactivation of
Cdc7 could have allowed only the earliest of early origins to fire. Therefore, the delay in cdc7-4 could
be due to only a subset of early origins firing or a defect in fork stabilization/restart. BrdU
incorporation and 2D gel experiments analyzing initiation and fork progression in these mutants should
be performed to discern between these possibilities.
148

Figure 37.

149

Appendix E: Viability, DNA content and 2D gel analysis of mrc1 rrm3
ts
cells

Figure 38. mrc1 rrm3
ts
cells lose viability and display decreased tRNA pausing at the restrictive
temperature. (A) WT (SSy211) and four strains containing isolates of pRS415-rrm3
ts
(SSy325-328)
were grown at the permissive temperature (23
o
C) and shifted to the restrictive temperature (37
o
C) for
the indicated amount of time. Cells were then ten-fold serially diluted and plated onto YEPD. Plates
were photographed after three days at 23
o
C. (B) WT (SSy211) and mrc1 rrm3
ts
(SSy325) strains were
grown asynchronously at the permissive temperature (25
o
C). After overnight growth, half of the culture
was shifted to the restrictive temperature (37
o
C) for five hours. Samples were collected for DNA
content analysis (B) and 2D gel analysis (C). DNA isolated from strains in (B) was digested with SpeI
and probed for the specified tRNA.
150

Figure 38.

151

Appendix F: The H2A.Z histone variant affects replication timing and bud
emergence

The contents of this appendix have been published and can be found in
Molecular and Cellular Biology (2006, Vol 26, pp 489-501).

Overview

In all eukaryotes, DNA is packaged into chromatin which consists of
histones and DNA. Histones can be modified in a number of different ways. These
modifications, which include: acetylation, methylation, phosphorylation, and
ubiquitination affect compaction of the chromatin. In turn, chromatin compaction
affects processes within the cell such as: DNA repair, cell cycle progression, gene
expression and DNA replication. In addition to histone modifications, histone variants
also exist which play roles in genomic integrity, transcription and chromosome
segregation. Histone H2A variants are highly conserved proteins found ubiquitously
in nature and are thought to perform specialized functions in the cell. Studies in yeast
on the histone H2A variant H2A.Z have shown a role for this protein in transcription
as well as chromosome segregation. The publication that resulted from a collaboration
with Rohinton Kamakaka’s lab, Dhillon et al, focuses on understanding the role of
H2A.Z during cell cycle progression. We found that htz1  cells were delayed in DNA
replication and progression through the cell cycle. Furthermore, cells lacking H2A.Z
required the S-phase checkpoint pathway for survival. We also found that H2A.Z
localized to the promoters of cyclin genes, and cells lacking H2A.Z were delayed in
the induction of these cyclin genes.

152

Contribution to the Study
Our initial goal in this collaboration was to examine replication
intermediates in htz1  cells. Given that we knew htz1  cells were delayed in
completion of S phase, we reasoned that this delay was due to decreased origin firing
or increased pausing at heavily transcribed tRNA genes. To this end, we analyzed
replication intermediates in WT and htz1  cells at ARS305 and SUP6, which is a
tRNA near ARS607. The experiment was carried out at 15
o
C to slow replication in
these cells and the results presented in Appendix F demonstrate that replication
intermediates persist at ARS305 and SUP6, which indicates that replication proceeds
asynchronously in htz1  cells. This is consistent with delayed entry into S phase (see
Fig 39B) and delayed in bud emergence (see Fig 39C). Because this experiment was
carried out at a low temperature, it allowed for careful analysis of cell cycle
progression, which could have been masked by faster kinetics at a higher temperature.
Based primarily on the results from this experiment, our collaborators examined
Htz1’s contribution to cell cycle progression. Interestingly, they found that htz1  cells
are delayed in Sic1 degradation and induction of Cln2 and Clb5 transcription. Taken
together, these results demonstrate that Htz1 plays an important role at the G1-S
transition. Potentially, studies like these performed in yeast could lead to new cancer
drug targets that inhibit the transcription of genes necessary for cell cycle transition in
human tumors.

153

Figure 39. The H2A.Z histone variant affects replication timing and bud emergence. Wild-type
(ROY1945) and htz1  (BUy828) cells were arrested in late G1 with -factor at 25
o
C and released into
pre-chilled 15
o
C YEPD. Samples were collected at the indicated times for 2D gel analysis (A) and
probed for the early firing origin, ARS305. Blots were then stripped and reprobed for the tRNA, SUP6,
which is located at 212kb on Chromosome VI. Samples were also collected for DNA content analysis
(B) and budding analysis (C). See text above for details.
154

Figure 39.

155

Appendix G. 2D gel analysis of ptc2  pph3  cells during reocovery from DNA
damage

Figure 40. ptc2  pph3  cells do not fire the late origin, ARS603, during recovery from MMS-
induced DNA damage. WT (SSy187), pph3  (SSy188), ptc2  (SSy248) and ptc2  pph3  (SSy249)
cells were arrested in -factor at 30
o
C for 2 hours and released into pre-warmed 30
o
C YEPD media
containing 0.033% MMS for 60 min. After MMS exposure, cells were pelleted and resuspended in pre-
warmed 30
o
C YEPD media lacking MMS. At the indicated times samples were collected for 2D gel
analysis (A) and DNA content analysis (B). (A) Samples collected for 2D gel analysis were probed for
replication intermediates at the early firing origin, ARS1 and the late firing origin, ARS603.
Replication bubbles were detected at similar times at ARS1 in all four strains demonstrating that
initiation was not affected by the phosphatase mutants. In WT and ptc2  cells, ARS603 is passively
replicated by replication forks progressing through the origin. Interestingly, ARS603 firing can be
detected in pph3  cells, which may be due to slow fork movement, which allows more time for origin
firing. Alternatively, the firing of ARS603 may reflect the action of another phosphatase such as Ptc2,
which may dephosphorylate Rad53 on specific residues, and result in derepression of late origin firing.
In line with the latter hypothesis, only a small amount of replication intermediates are detected at
ARS603 in ptc2  pph3  cells, indicating that time alone is not sufficient to fire the late origin,
ARS603.
156

Figure 40.

Asset Metadata

Creator Szyjka, Shawn Joseph (author)

Core Title The intra-S phase checkpoint and its effect on replication fork dynamics in saccharomyces cerevisiae

Contributor Electronically uploaded by the author (provenance)

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publication Date 10/21/2008

Defense Date 07/25/2008

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag 2-D gel electro,BrdU,cell cycle checkpoint,Claspin,DNA damage,DNA repair,DNA replication,microarray,Mrc1,OAI-PMH Harvest,phosphatase,Pph3,Rad53,replication fork,replication fork progression,Rrm3

Language English

Advisor Aparicio, Oscar (committee chair), Finkel, Steven E. (committee member), Forsburg, Susan (committee member), Qin, Peter Z. (committee member), Rice, Judd C. (committee member)

Creator Email hammonds77@gmail.com,sszyjka@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m1688

Unique identifier UC1230529

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Legacy Identifier etd-Szyjka-2341.pdf

Dmrecord 119913

Document Type Dissertation

Rights Szyjka, Shawn Joseph

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email uscdl@usc.edu

Abstract (if available)

Abstract Duplication of a cell's genetic material is of paramount importance. This task must be completed accurately, efficiently and occur once and only once within any given cell cycle. Origin "firing", which occurs according to a temporal program, results in the establishment of bidirectional replication forks. As replication forks traverse the chromosomal landscape, their stability is threatened by endogenous "obstacles" such as heavily transcribed tRNAs, protein-DNA complexes and "replication slow zones". In addition, fork stability can also be threatened by genotoxic agents. During S phase, in response to genotoxin-induced stress, the cell activates the intra-S phase checkpoint. The intra-S phase checkpoint is comprised of three major groups of proteins: sensors, adaptors and effectors. Sensor proteins detect problems at the replication fork and elicit a phosphorylation cascade that is mediated by adaptor proteins and results in the activation of effector kinases. Together, these proteins act to inhibit late origin firing, slow cell cycle progression, upregulate DNA repair genes, and stabilize replication forks until the stress has been alleviated.