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New tools for whole-genome analysis of DNA replication timing and fork elongation in saccharomyces cerevisiae
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New tools for whole-genome analysis of DNA replication timing and fork elongation in saccharomyces cerevisiae
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NEW TOOLS FOR WHOLE-GENOME ANALYSIS OF DNA REPLICATION
TIMING AND FORK ELONGATION IN SACCHAROMYCES CEREVISIAE
by
Christopher John Viggiani
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)
August 2007
Copyright 2007 Christopher John Viggiani
ii
Dedication
I dedicate this work to my loving grandparents who have never ceased to inspire
me…
To Richard and Mary Lou Hoefer who, by example, taught me to work hard and be
proud; to be strong and dependable; to respect those who have earned it; to live fully
yet simply; to have faith in family and friends; to always be loving and accepting;
and to live life as PapaHoef quite simply advised: “Do good things out there.”
To Nick and Rose Marie Viggiani for being larger-than-life; for teaching me to trust
my instincts and be spontaneous; for their combined warmth; for second helpings
“just for the flavor”; for overcoming great tragedy and emerging at the center of a
very rambunctious family; for teaching me love, faith and compassion; and most
importantly, for teaching me that our actions here matter, for we’ll all be
remembered.
iii
Acknowledgements
I would like to thank Oscar Aparicio for his guidance and patience
throughout the course of this work. I’ve been fortunate to know Oscar as an
inspiring scientist, a remarkable academic, a patient and motivating mentor, and now
as a colleague and friend. He was the ever-present guiding force behind my
intellectual development, yet the independence he granted me demonstrated a
confidence and trust, for which I am deeply thankful. He’s also a force to be
reckoned with on the basketball court and behind the grill. I owe a very special
thanks to Laura Sanders for her constant love and support. Her laughter and smile
have helped make even the hardest days worthwhile and I can’t imagine having done
this without her. I’ve counted on her in many ways, not least of all to remind me
when I begin to take myself too seriously!
I thank my entire family for their love and support over the years. In
particular to my parents, John and Nancy, who taught me kindness and respect; and
importantly, the value of hard work (though it may not have been appreciated at the
time!). They somehow found the fine distinction between expecting success, and
demanding it, which gave me the courage to think big and risk failure without the
fear of ever losing their love and respect. I owe a very special thanks to my sister
Katie, who has been a great friend and neighbor. Her confidence, strength, and
balance are simply amazing, and her quiet, steady resolve is inspiring, and in stark
contrast to my own resolve, which is rarely steady and anything but quiet. I should
also confess here that the vast majority of my funny jokes, comedic observations, or
iv
sarcastic quips (for which I’ve become basically legendary) have originated with, or
been outright stolen from Katie. Sorry.
I’d like to thank the entire Aparicio Lab, past and present - Jen, Dan,
Fangfang, Shawn, Yuan, Yan, and our undergrads - for making the lab a great place
to spend six years. Their helpful discussions, reagents and humor are greatly
appreciated. Particularly, I want to thank Jennifer Aparicio and Shawn Szyjka.
Jen’s guidance and friendship were instrumental to my success. She was the driving
force behind the early Rpd3 work and played an important advisory role for most of
my work. Her hard work developing the microarrays was monumental, and
commendable, and made much of my work possible. Shawn initiated the Mrc1 work
and subsequently has generated a staggering amount of high-quality data. His
consistent determination and perseverance was a great motivator to me.
Thanks to my dissertation committee – Steve Finkel, John Tower, Steve
Goodman, and Judd Rice for their time, their helpful suggestions, and the gift of
humility. Additional thanks to Steve Finkel and Susan Forsburg for the close role
they’ve played in my intellectual development as well as the guidance they’ve
provided me as I move on to new things. I also thank Michelle Arbeitman and her
lab members - Laura, Tom, Matt, Justin, Saori - not only for sharing their lab space
and equipment, but also for the delight (and volume) of their constant laughter.
Thanks to Michael Stallcup and the GMCB Training Grant for its role in my
intellectual development and for financial support. Thanks to Gurinder Shahi and
my BioBusiness classmates for expanding my mind.
v
Thanks to the labs of Myron Goodman, Norm Arnheim, Susan Forsburg and
John Tower for the use of equipment. Thanks to C. Cass, K. Struhl, and E. Schwob
for plasmids. Thanks to GPSS and BGSA for the work they do supporting graduate
student life. A warm thanks to Bill Trusten, Eleni Yokas, Linda Bazilian, Chris
Yokas, and Christina Tasulis for their nuanced understanding of the USC
bureaucracy. When the university periodically decided to shut me down, lock me
out, block my registration or withhold my stipend, they were there to cut through the
red tape and fix my problems. Thanks to my friends in the Forsburg and Finkel Labs
and throughout the Department, as well as those outside the department, including
the League of Ordinary Gentlemen and all who help me find balance…and rock
music.
vi
Table of Contents
Dedication ii
Acknowledgements iii
List of Tables vii
List of Figures viii
Abstract x
Chapter 1: Introduction 1
Chapter 2: New vectors for simplified construction of BrdU
incorporating strains of Saccharomyces cerevisiae
11
Chapter 3: The Sin3-Rpd3 HDAC complex regulates the initiation
timing of a subset of late origins prior to entry into S
phase
18
Chapter 4: BrdU incorporation demonstrates a central role for
Mrc1 in replication fork progression and provides a
new tool for studying fork movement across
chromosomes
40
Chapter 5: Materials and methods 63
References 73
Appendix A: BrdU pools are depleted over time 82
vii
List of Tables
Table 1: Integration and confirmation of BrdU-Inc vectors 14
Table 2: Primer sequences for confirmation of BrdU-Inc constructions 64
Table 3: Strain table 65
viii
List of Figures
Figure 1: The temporal pattern of DNA replication origin initiation 2
Figure 2: The BrdU-Inc plasmids 12
Figure 3: Integration of a BrdU-Inc vector and diagnostic PCR
confirmation scheme
13
Figure 4: PCR and immunoblot confirmation of BrdU-Inc strains 15
Figure 5: Quantitative immunoblot analysis demonstrates effective BrdU
incorporation
16
Figure 6: The Sin3-Rpd3 complex is not significantly enriched at
replication origins
20
Figure 7: Replication origins are hyperacetylated in the absence of Rpd3 22
Figure 8: Late origins escape HU-induced checkpoint inhibition in
rpd3Δ strains as measured by the incorporation of BrdU
24
Figure 9: Destruction of Rpd3-td protein for several generations results
in late origin deregulation in HU
26
Figure 10: Rpd3 function prior to S phase entry regulates the initiation
timing of late origins
29
Figure 11: Identification of early replicating regions of chromosome VI
in rpd3Δ cells
32
Figure 12: Mrc1 is required for normal progression of DNA synthesis 43
Figure 13: Slower progression of Cdc45 in mrc1Δ cells 46
Figure 14: Slowed DNA synthesis across chromosome VI in mrc1Δ cells 47
ix
Figure 15: BrdU pulses label genomic regions that replicate during
discrete windows of time
52
Figure 16: Replication forks recover slowly after stalling in HU 54
Figure 17: Replication forks proceed across a damaged DNA template in
the presence of MMS
59
Figure 18: BrdU incorporation at late replicating sequences is less
efficient than early replicating sequences
92
x
Abstract
The faithful duplication of the genome during S phase is critical to a eukaryotic
cell’s healthy proliferation. Since errors in DNA replication can lead to disastrous
cellular consequences, this process has evolved to be highly regulated throughout
species. In Saccharomyces cerevisiae, DNA replication initiates from multiple
discrete chromosomal loci, termed origins of replication. DNA synthesis proceeds
bidirectionally away from origins to replicate the entire genome prior to mitosis and
cell division. Although many of the events regulating entry into S phase have been
characterized, the regulation of events occurring within S phase are more obscure.
For instance, replication origins initiate according to a temporal program that often
correlates with transcriptional activity and appears to be regulated, in part, by their
surrounding chromatin structure. To examine replication timing and replication fork
elongation on a genome-wide scale I have developed a series of integrating plasmid
vectors that enable the one-step construction of yeast strains that can incorporate the
thymidine analogue BrdU into replicating DNA. By immunoprecipitating BrdU-
labeled DNA I determine that the histone deacetylase complex Sin3-Rpd3 functions
prior to S phase to properly delay the initiation timing of a subset of late origins,
likely by creating a repressive chromatin structure around origins. I also use BrdU
incorporation to help demonstrate that Mrc1, in addition to its checkpoint function,
plays a central and constitutive role at normal replication forks. Finally, I provide a
novel BrdU-pulse approach, coupled with microarrays, to examine how replication
forks respond to, and recover from, replication stress and DNA damage.
1
CHAPTER 1
Introduction
DNA replication initiates according to a temporal program
The faithful duplication of the genome prior to mitosis and cell division is critical to
a eukaryotic cell’s healthy proliferation. Since errors in DNA replication can lead to
disastrous cellular consequences, this process has evolved to be highly regulated
throughout species. In the budding yeast Saccharomyces cerevisiae, DNA
replication initiates at discrete origins termed autonomously replicating sequences
(ARS’s) that contain a conserved ARS consensus sequence (ACS), which is bound
throughout the cell cycle by the origin recognition complex (ORC) (Bell and Dutta,
2002). ARS elements also contain less-conserved B elements that are important for
unwinding and loading additional replication factors (Bell, 1995). ORC-bound
origins become poised for initiation in G1 phase with the ordered assembly of Cdc6
and Cdt1, which in turn load the ring-shaped MCM2-7 putative helicase proteins,
completing the formation of the pre-replicative complex (pre-RC). In S. cerevisiae,
activation of the pre-RC requires the activity of two kinases, CDK and DDK, which
regulate the assembly of a number of replication factors including Cdc45, Mcm10
and the GINS complex. Ultimately, origin DNA is unwound and stabilized by RPA,
and replicative DNA polymerases are loaded, forming two bidirectional replication
forks where DNA synthesis occurs (Machida, et al., 2005).
2
While the cell cycle events controlling entry into S phase are well
characterized, the events occurring within S phase remain obscure. Though yeast
replication origins share sequence similarities and the same requirements for
initiation, a chromosomal view of DNA replication shows origins behave quite
differently, firing in a temporally controlled manner (Figure 1). Each replication
origin fires only once during a given cell cycle but different origins initiate at
stereotypic times throughout S phase leading to their functional characterization as
early- or late-firing. Early origins tend to be clustered and more centromere-
proximal whereas late origins are often found toward the middle and ends of
chromosome arms (Machida, et al., 2005; Raghuraman, et al., 2001; Weinreich, et
al., 2004).
Figure 1. The temporal pattern of DNA replication origin initiation. Origins (ARS’s) near
centromeres (CEN) tend to fire earlier in S phase, followed by more telomere-proximal late origins.
CEN
CEN
CEN
ARS ARS ARS ARS ARS ARS ARS
3
Additionally, in yeast over 400 origins have been identified to bind ORC and form
pre-RC’s in vivo, but only a small subset become activated in a typical S phase
(Feng, et al., 2006; Newlon and Theis, 2002; Raghuraman, et al., 2001; Wyrick, et
al., 2001; Xu, et al., 2006). The precise mechanisms controlling origin usage and
initiation timing have remained elusive but it has become apparent that chromatin
structure surrounding origins is a contributing factor.
A clear relationship between gene expression and DNA replication timing
has been described with actively transcribed, euchromatic chromosomal regions
replicating early, and silent, heterochromatic regions replicating later in S phase
(Gilbert, 2002; Goren and Cedar, 2003). This correlation holds in higher eukaryotes
such as humans and Drosophila but is less strict in yeast (Cimbora, et al., 2000;
MacAlpine, et al., 2004; Raghuraman, et al., 2001). Nevertheless, several lines of
evidence demonstrate that sequences flanking the origin, not the origin itself, play a
role in the temporal replication program in yeast as well. Many late origins replicate
early when placed on a circular plasmid, but flanking the plasmid-borne origin with
its native chromosomal DNA tends to restore late replication timing (Ferguson and
Fangman, 1992; Friedman, et al., 1996). Also, initiation of early origins is delayed
when they are moved to late replicating chromosomal regions, such as telomeres
(Ferguson and Fangman, 1992). These findings demonstrate that the chromatin
context surrounding ARS’s at different chromosomal locations plays an important
role in initiation timing.
Using a recombinase-based approach to release an origin and some flanking
sequences at different cell cycle times, Raghuraman and colleagues have found that
4
the sequences conferring late initiation to telomere-proximal origins are set up
between M phase and the G1/S phase transition. When this origin was released just
prior to the G1/S phase transition, the episomal DNA fragment replicated late,
suggesting the appropriate chromatin context had been assembled. Conversely,
when the origin was released prior to G1 phase (in M phase), the episomal DNA
reverted to early firing, demonstrating that the flanking sequences conferring late
timing had not yet been established (Raghuraman, et al., 1997). Work using
Xenopus egg extracts and mammalian cells also suggests replication timing is
determined in early G1 phase, suggesting that the timing decision point (TDP) may
be a conserved cell cycle feature in both yeast and animals (Dimitrova and Gilbert,
1999; Li, et al., 2001; Li, et al., 2003).
Protein factors have been shown to directly regulate initiation timing as well.
Deletion of SIR proteins, which are required for transcriptional silencing at
telomeres, has been shown to advance the timing of telomere-proximal origins and
targeting Sir4 to ARS305 was sufficient to convert it from an early to late origin
(Stevenson and Gottschling, 1999; Zappulla, et al., 2002). These findings reveal that
proteins that modify chromatin in order to silence transcription also play a role in
establishing late replicating origins.
Our lab, and others, have shown that the transcriptional silencing complex
Sin3-Rpd3 also regulates the temporal replication program (Aparicio, et al., 2004;
Vogelauer, et al., 2002). Sin3-Rpd3 is a histone deacetylase (HDAC) complex that
is targeted to promoters by Ume6 where it deacetylates several lysine residues on the
tails of histones H3 and H4 resulting in chromatin compaction and transcriptional
5
repression of various genes important for meiosis, metabolism and cell cycle
regulation (Bernstein, et al., 2000; Burgess, et al., 1999; Kadosh and Struhl, 1997;
Rundlett, et al., 1998). Under stress conditions Sin3-Rpd3 appears to be rapidly
recruited to many other promoters through alternate mechanisms where it can act as
both a repressor and activator (Bernstein, et al., 2000; De Nadal, et al., 2004;
Humphrey, et al., 2004). Recent studies have demonstrated that Sin3-Rpd3 plays a
role in resetting chromatin after transcriptional elongation as well (Carrozza, et al.,
2005; Joshi and Struhl, 2005).
Deletion of the Sin3-Rpd3 complex results in histone hyperacetylation at
specific promoters and to some extent globally (Kadosh and Struhl, 1998;
Kurdistani, et al., 2004; Vogelauer, et al., 2000). Interestingly, in yeast strains
lacking Rpd3, late origins initiate earlier, suggesting a direct relationship between
histone acetylation levels and replication timing (Aparicio, et al., 2004; Vogelauer, et
al., 2002). Indeed, targeting the histone acetyltransferase (HAT) Gcn5 to the late
origin ARS1412 was sufficient to advance its initiation timing (Vogelauer, et al.,
2002). In Drosophila follicle cells, during developmental amplification of chorion
genes, mutation of Rpd3 results in histone H4 hyperacetylation, Orc2p redistribution,
and increased DNA replication, which was suppressed by tethering a catalytically
active Rpd3 to an origin region (Aggarwal and Calvi, 2004). Similar to the studies in
yeast, tethering of a HAT elevated levels of DNA replication in the Drosophila
system.
6
Together, the observations described provide a working model for replication
timing: chromatin-modifying enzymes create hyper- and hypoacetylated regions that
flank origins and alter their compaction states, thus conferring early or late initiation
timing respectively. However, more mechanistic models are still needed. One
model suggests that compacted chromatin surrounding late replicating regions makes
origins inaccessible to critical replication factors such as Cdc45 or DDK. Another
model suggests the spatial organization of origins in the nucleus may account for
replication timing as early origins tend to localize as discrete nuclear foci and late
firing regions tend to be located around the nuclear periphery. A third model directly
links replication and transcription where replication timing is actively regulated to
correlate with the availability of critical transcriptional regulators (Gilbert, 2002).
The possibility that these models are not mutually exclusive further confounds our
understanding of the temporal replication program.
To date, two studies have examined the whole-genome binding of Sin3-Rpd3
and others have determined gene expression changes in yeast strains lacking the
complex (Bernstein, et al., 2000; Humphrey, et al., 2004; Keogh, et al., 2005;
Kurdistani, et al., 2002; Robert, et al., 2004). In this work I examine the advanced
initiation timing of late origins in rpd3Δ cells on a chromosomal scale, and provide a
method to identify advanced firing regions genome-wide. For a more complete
picture of the relationship between transcription and DNA replication, a genomic
comparison between Rpd3’s regulation over late origins with its genome-wide
binding and transcriptional functions will be useful. The precise mechanism
underlying late origin regulation by Rpd3 is also unclear. Does it bind directly to
7
origins and deacetylate adjacent histones or does it function through a less direct
targeting mechanism? In addition, while it is known that late replication timing of
the telomere-proximal origin ARS501 is established prior to S phase, it is still
unknown whether specific chromatin-modifying proteins have a cell cycle execution
point where they establish replication timing and define origins as early or late. In
chapter 3, I examine these questions and present a novel approach to identify Rpd3-
regulated origins on a genome-wide scale.
BrdU incorporation provides a useful tool for measuring DNA synthesis
To examine the questions outlined above, and others regarding replication fork
progression, we wished to measure the incorporation the thymidine analogue
bromodeoxyuridine (BrdU) into newly synthesized DNA. Thymidine analogues
have been used in many biological systems to study various aspects of cell growth
and proliferation, including events occurring within S phase such as chromosome
replication patterns and DNA repair (Dolbeare, 1996). Unfortunately, use of this
valuable tool has been limited in yeast, as fungi lack the thymidine salvage pathway
that enables efficient uptake and incorporation of extracellular thymidine or its
analogs into replicating DNA (Grivell and Jackson, 1968). Specifically, yeast lack
an appropriate nucleoside transporter for thymidine uptake, and thymidine kinase to
phosphorylate thymidine into TMP, which would similarly convert BrdU into
BrdUMP. By inhibiting the de novo thymidine synthesis pathway or isolating
thymidine uptake (tut) mutants, yeast strains that incorporate exogenous thymidine
8
or its analogues into DNA have been characterized (Leff and Lam, 1976; Sclafani
and Fangman, 1986). However, their incorporation efficiency is poor. It has also
been shown that expression of Herpes simplex virus thymidine kinase (HSV-TK) can
improve uptake of thymidine or BrdU into yeast (McNeil and Friesen, 1981).
Recently, Schwob and colleagues increased the efficiency of BrdU
incorporation into chromosomal DNA by engineering Saccharomyces cerevisiae
strains that contain several copies of HSV-TK expressed by the constitutive yeast
GPD promoter (Lengronne, et al., 2001). These TK
+
strains have enabled analysis of
nucleotide incorporation into total DNA during replication using immunocytology or
immunoblotting, into individual DNA fibers using DNA combing, and into specific
DNA sequences by immunoprecipitation (Lengronne, et al., 2001; Szyjka, et al.,
2005). This immunoprecipitation strategy may be combined with DNA microarrays
to provide a more global view of chromosomal replication (see chapters 3 and 4)
(Katou, et al., 2003).
The ability to analyze BrdU incorporation has proven a valuable asset to the
study of DNA replication in yeast. However, one obstacle to this system is the
requirement for at least seven copies of the tandemly integrated GPD-HSV-TK gene
construct (Lengronne, et al., 2001). This requirement complicates new strain
construction by reducing the frequency of the desired strain among transformants,
and requires monitoring of existing strains for copy number changes (due to
intrachromosomal or interchromatid recombination of the tandem repeats), which
can alter or eliminate BrdU incorporation and confound analysis. Because of the
large size of the chromosomal fragment containing seven copies of the GPD-HSV-
9
TK plasmid (35 kb), pulsed-field gel electrophoresis was employed for copy number
determinations in these TK
+
yeast strains (Lengronne, et al., 2001). While
quantitative PCR may also be useful for copy number determination of TK
+
strains,
either of these methods requires additional effort, equipment, time, and expense
beyond standard strain construction methods.
The expression of recombinant human equilibrative nucleoside transporter 1
(hENT1) has been shown to further increase the efficiency of thymidine uptake and
has been used in concert with HSV-TK to enhance BrdU incorporation efficiency in
S. cerevisiae (Vernis, et al., 2003; Vickers, et al., 1999), and in the fission yeast S.
pombe (Hodson, et al., 2003; Sivakumar, et al., 2004). For the S. cerevisiae system,
Vernis et al. placed each gene, HSV-TK and hENT1, under the GAL1-10 promoter
to induce thymidine uptake upon addition of galactose. With this system, a single
insertion of each gene construct is sufficient to confer efficient BrdU incorporation,
thus obviating the need to identify strains with many integrated copies of HSV-TK
and to routinely re-confirm these strains as described above. However, two
integration and confirmation steps are required, and two selectable markers are
consumed. In addition, this system requires induction by addition of galactose,
precluding standard growth in glucose, which blocks rapid induction of the system.
In chapter 2 I describe the simplified construction, maintenance, and analysis
of BrdU-incorporating (BrdU-Inc) strains of S. cerevisiae (Viggiani and Aparicio,
2006). I have constructed a series of single plasmid vectors that express both HSV-
TK and hENT1 from constitutive promoters. Each vector in the set contains a
different selectable marker (HIS3, TRP1, LEU2, URA3), which also serves to target
10
integration to the corresponding genomic locus. Proper integration at single copy is
easily achieved, confirmed and maintained, and strains containing a single insertion
of a BrdU-Inc vector incorporate BrdU with similar efficiency as existing TK
+
strains.
The ability to conveniently measure BrdU incorporation into yeast genomic
DNA using commercially available antibodies provides a powerful tool that is highly
amenable to whole-genome approaches. BrdU can be immunoprecipitated from
newly-synthesized DNA, labeled with fluorescent dyes and hybridized onto DNA
microarrays, in a manner similar to ChIP-chip procedures. In chapter 3, I use a BrdU
immunoprecipitation approach to identify late-replicating regions regulated by Sin3-
Rpd3 and suggest that this HDAC complex creates condensed chromatin around
origin regions prior to S phase entry. In chapter 4, I present this approach as a tool to
examine the progression of replication forks across chromatin. In particular, I use
BrdU incorporation to help conclude that Mrc1 (mediator of replication checkpoint
protein 1), a protein required for replisome stability in response to exogenous stress,
plays a central and constitutive role at normal replication forks. Also, using a novel
BrdU-pulse approach, I measure replication fork progression more precisely and
provide a useful tool for examining how forks respond to genotoxic insults.
11
CHAPTER 2
New vectors for simplified construction of BrdU-incorporating strains of
Saccharomyces cerevisiae
OVERVIEW
The thymidine analog BrdU is a powerful tool for analyzing nucleotide incorporation
in studies of DNA replication or repair. However, S. cerevisiae lacks the thymidine
salvage pathway that enables efficient cellular uptake and incorporation of thymidine
analogues into DNA. Here, I have created a set of vectors that simplify the
construction and use of BrdU-incorporating (BrdU-Inc) strains of budding yeast.
With these BrdU-Inc vectors, one-step integration of a single copy produces yeast
that efficiently incorporate BrdU upon its addition to the medium. These vectors
ease strain construction and maintenance, thereby facilitating routine use of BrdU for
analysis in yeast. This work, and the detailed description of the plasmids’
construction described in chapter 5, was published in 2006 in the journal Yeast
(Viggiani and Aparicio, 2006).
RESULTS AND DISCUSSION
A series of single-copy BrdU incorporation vectors
We designed a set of plasmids that, when integrated in the genome at single copy,
enable budding yeast cells to incorporate BrdU constitutively (Figure 2). A BrdU-
incorporation cassette (BrdU-Inc) was built with HSV-TK under the control of the
strong constitutive GPD promoter, and hENT1 under the control of the ADH1
12
promoter, which expresses at a basal (uninduced) level under typical laboratory
growth conditions (Guarente, 1991). BrdU-Inc was inserted into the four yeast
integrating vectors, pRS403, pRS404, pRS405, and pRS306 (Sikorski and Hieter,
1989), yielding plasmids p403-BrdU-Inc, p404-BrdU-Inc, p405-BrdU-Inc, and p306-
BrdU-Inc, respectively.
Figure 2. The BrdU-Inc plasmids. ADH1-hENT1 and GPD-HSV-TK were inserted into the pRS
integrating vector series (HIS3, TRP1, LEU2, URA3) as shown and described in detail in chapter 5.
Diagram is not drawn to scale.
One-step construction of BrdU-Inc strains
We transformed wild-type cells with plasmid p403-BrdU-Inc, linearized by
restriction digestion in the HIS3 gene, which targets plasmid integration to the
endogenous his3 locus (Figure 3). Candidate transformants were screened for
successful plasmid integration by PCR analysis using primer sets that yield
~1.3 kb ~1.5 kb ~0.7 kb ~2.1 kb
HSV-TK hENT1 ADH1 GPD
13
diagnostic product sizes (Figure 3 and Table 1). Primers RS1 and RS2 were
designed with specificity to sequences in p403-BrdU-Inc (and the other BrdU-Inc
vectors). Another pair of primers was designed to flank the genomic his3 locus (a
different primer set was designed flanking each of the other three targeted loci)
(Table 1). Although this PCR analysis may be performed directly on yeast cells
without prior DNA isolation (data not shown), we prepared genomic DNA from cells
grown in the presence of BrdU for 1 hour for this analysis, which also allowed us to
confirm BrdU uptake into DNA (see below).
Figure 3. Integration of a BrdU-Inc vector and diagnostic PCR confirmation scheme.
Linearization of p403-BrdU-Inc within the HIS3 selectable marker directs plasmid integration to the
endogenous his3 locus. Diagnostic PCR reactions using different combinations of primers 5’His3,
3’His3, RS1, and RS2 confirm proper plasmid integration (see text and Table 1 for details). Primer
sequences and details of construction and verification of strains made with the other BrdU-Inc vectors
are available in Tables 1 and 2 in chapter 5. Diagram is not drawn to scale.
RS1
RS2
his3
3’His3 5’His3
RS1 RS2 5’His3 3’His3
NheI cut
HIS3 his 3 BrdU-Inc
1293 bp
1542 bp 1403 bp
14
Table 1. Integration and confirmation of BrdU-Inc vectors
Three separate PCR reactions were designed for each plasmid transformation
(Figure 3 and Table 1): Reaction 1 contains three primers (5’HIS3, 3’HIS3 and RS2)
that test the structure at one end of the integration; reaction 2 is similar, using three
primers (5’HIS3, 3’ HIS3 and RS1) that test the structure at the other end of the
integration; and reaction 3 contains two plasmid-specific primers (RS1 and RS2) that
only yield a product if multiple plasmid integrations occur in tandem, or if the
plasmid integrates at a site other than the targeted locus. As is typical with this type
of plasmid integration, the majority of transformants contain the plasmid correctly
integrated at the targeted locus, and about half of these transformants contain
multiple copies of the plasmid (Figure 4). For example, in reaction 1 (Figure 4, top
panel), transformants 1 and 2 show a 1293 bp product as observed in the
untransformed, wild-type strain, indicating these strains contain an unaltered his3
locus; however, transformants 3 through 8 show a 1542 bp product, indicating
insertion of p403-BrdU-Inc at his3. Reaction 3 reveals the presence of additional
copies of p403-BrdU-Inc in transformants 3 through 5 (Figure 4, middle panel).
PCR Confirmation
Plasmid Marker Integrate Reaction Primers
Single
integration
at target
No
integration
at target
Multiple integrations at
target or integration
away from target
p403-BrdU-Inc HIS3 NheI 1 5'His3 + 3'His3 + RS2 1542 bp 1293 bp -
2 5'His3 + 3'His3 + RS1 1403 bp 1293 bp -
3 RS1 + RS2 - - 1649 bp
p404-BrdU-Inc TRP1 XbaI 1 5'Trp1 + 3'Trp1 + RS2 1406 bp 1129 bp -
2 5'Trp1 + 3'Trp1 + RS1.1 1061 bp 1129 bp -
3 RS1.1 + RS2 - - 1338 bp
p405-BrdU-Inc LEU2 HpaI 1 5'Leu2 + 3'Leu2 + RS2 2553 bp 2277 bp -
2 5'Leu2 + 3'Leu2 + RS1 2424 bp 2277 bp -
3 RS1 + RS2 - - 2571 bp
p306-BrdU-Inc URA3 StuI 1 5'Ura3 + 3'Ura3 + RS2 1428 bp 1162 bp -
2 5'Ura3 + 3'Ura3 + RS1 1311 bp 1162 bp -
3 RS1 + RS2 - - 1577 bp
15
Strain constructions and confirmations with the other plasmids in the set were carried
out in analogous fashion and yielded comparable results (data not shown).
Figure 4. PCR and immunoblot confirmation of BrdU-Inc strains. Genomic DNA from
independent transformants was isolated and subjected to diagnostic PCR to confirm BrdU-Inc
integration at the his3 locus. Top Panel – Reaction 1, containing primers 5’His3, 3’His3 and RS2.
Middle Panel – Reaction 3, containing primers RS1 and RS2. Bottom Panel - 10 µl of genomic DNA
was spotted and detected with anti-BrdU antibody.
As a final confirmation of the strain construction, we performed a rapid assay
for BrdU incorporation. As mentioned above, small cultures of candidate
transformants were grown in the presence of BrdU for 1 hour prior to preparation of
genomic DNA. Genomic DNA from each candidate was denatured by heating and
applied to a nylon membrane. Immunoblotting with anti-BrdU antibody detected
incorporated BrdU in transformants 3 through 8, which each contained one or more
copies of BrdU-Inc (Figure 4, bottom panel).
anti-BrdU
1 2 3 4 5 6 7 8 WT
1 2 3 4 5 6 7 8 WT
1.9
1.4
1.3
0.7
2.3
1.9
1.4
1.3
M
16
Efficient BrdU incorporation with a single-copy vector
We compared the efficiency of BrdU incorporation in a single-copy BrdU-Inc strain
with a TK
+
strain harboring seven tandem copies of GPD-HSV-TK (Lengronne, et
al., 2001). We grew both strains in the presence of BrdU for 1 hour and isolated
total genomic DNA. Indicated amounts of DNA from each strain were applied to a
nylon membrane and analyzed by immunoblotting with anti-BrdU antibody.
Quantification of the chemiluminescence shows that similar levels of BrdU are
associated with equivalent amounts of genomic DNA isolated from these strains,
while analysis of DNA from each strain grown in the absence of BrdU demonstrates
the specificity of the signal (Figure 5). The results demonstrate that the BrdU-Inc
strains incorporate BrdU with very similar effectiveness as the TK
+
strains.
Consistent with this conclusion, we have recently used a BrdU-Inc strain to monitor
BrdU incorporation at specific DNA sequences by immunoprecipitation followed by
PCR detection (O'Neill B, et al., 2007). This strain proved as effective in this
analysis as a TK
+
strain used previously (Szyjka, et al., 2005).
Figure 5. Quantitative immunoblot analysis demonstrates effective BrdU incorporation. BrdU-
labeled genomic DNA was isolated from TK
+
(E1000) and BrdU-Inc (CVy43) strains. The indicated
amounts of DNA were applied to nylon and analyzed by immunoblotting. Both strains grew with
indistinguishable kinetics (data not shown).
100 50 10 5 1
Chemiluminescence
(Arbitrary Units)
5
15
25
DNA (ng)
0
5
10
15
20
25
1 2 3 4 5
E1000
CVy43
E1000
CVy43
BrdU - +
100 100 50 10 5 1
DNA (ng)
+ + + +
17
We have presented a set of integrating plasmids that allow for rapid, one-step
construction and confirmation of budding yeast strains that effectively incorporate
BrdU upon its addition to the growth medium. Because a single copy of the BrdU-
Inc vector is sufficient, strain construction is simplified and the resulting strains are
expected to be more stable and reliable than the previously described TK
+
strains.
The single-step nature of strain construction is particularly beneficial for highly
modified strains in which only a single selectable marker remains or strains that are
otherwise challenging to manipulate. This new vector set represents a valuable tool
for routine BrdU analyses in budding yeast.
18
CHAPTER 3
The Sin3-Rpd3 HDAC complex regulates the initiation timing of a subset of late
origins prior to entry into S phase
OVERVIEW
The replication of eukaryotic genomes follows a temporally staged program that is
especially pronounced in S. cerevisiae, with DNA replication initiating from distinct
origins that fire at reproducible times throughout S phase. While origins appear
similar at the DNA sequence level and require the same proteins for activation, the
events regulating their distinct initiation times within S phase remain obscure. It is
thought that the chromatin structure surrounding origins plays an important role in
regulating origin initiation timing. We have previously shown that the Sin3-Rpd3
histone deacetylase (HDAC) complex is required to properly delay the initiation
times of several late-firing origins. Data included in Figure 7 appears in that
published study (Aparicio, et al., 2004). In this chapter I expand our analysis of
Rpd3 and suggest that Sin3-Rpd3 sets up a repressive chromatin state that confers
late initiation to origins prior to S phase entry. Also, using BrdU and DNA
microarrays I identify chromosomal regions that show advanced initiation of DNA
replication in rpd3Δ cells. A better understanding of replication origin regulation by
Sin3-Rpd3 on a mechanistic and genomic level will help dissect the complex
chromosomal dynamics that regulate both transcription and DNA replication.
19
RESULTS AND DISCUSSION
Sin3-Rpd3 complex enables intra-S phase checkpoint regulation of late origins
by delaying their initiation time
We have demonstrated previously that Sin3-Rpd3 delays the initiation timing for
certain late-firing origins thus enabling late origin inhibition by the intra-S
checkpoint activator hydroxyurea (HU) (Aparicio, et al., 2004). HU causes
replication stress by depleting cellular nucleotide pools, which results in a signaling
cascade that greatly slows DNA replication and arrests cells in mid S phase by
stalling replication forks and preventing the initiation of late-firing origins until
nucleotide levels are restored. In rpd3Δ cells, however, late origin initiation is
advanced within S phase resulting in late origins that fire in the presence of the
checkpoint activator HU. The precise mechanism by which Sin3-Rpd3 properly
delays the firing of late origins is unknown and the interplay between DNA
replication and Rpd3-regulated transcriptional events is equally obscure. A working
hypothesis suggests that like its role in transcriptional silencing, Rpd3 may create a
repressive chromatin context by deacetylating histone tails and compacting
chromatin adjacent to replication origins.
We first examined whether Sin3, a vital regulatory protein in the Rpd3
HDAC complex, was bound to origins of replication using chromatin
immunoprecipitations (ChIP). Potentially, the Sin3-Rpd3 complex could exert its
function over origins through direct binding to regions near those origins. Evidence
also suggests that Rpd3 physically interacts with Cdc45, a component of the pre-RC,
further suggesting Sin3-Rpd3 may interact with origins (Gavin, et al., 2002).
20
However, Figure 6 suggests that the Sin3-Rpd3 complex regulates origins through a
mechanism other than direct, prolonged interaction. Sin3 binding was examined in
cells growing asynchronously as well as cell populations synchronized in G1 and M
phase but we did not observe significant enrichment at early or late origins above
background levels measured at the 3RXadj negative control telomeric locus. Sin3
was enriched at the INO1 and IME2 promoter regions demonstrating that our assay
detected Sin3 binding at promoter regions known to bind the HDAC complex
(Figure 6 and data not shown).
Figure 6. The Sin3-Rpd3 complex is not significantly enriched at replication origins. Cells
expressing Sin3-HA (JAy57) were grown asynchronously at 23
o
C and arrested in G1 using α factor or
M phase with nocodazole. Cells were harvested for ChIP analysis with anti-HA antibody and the
indicated sequences were analyzed for Sin3-HA binding by semi-quantitative PCR. The ratio of
immunoprecipitated to input DNA is plotted as “% Sin3 Bound.”
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0.16
0.18
0.2
1 2 3
Asynch G1 M
0.2
0.18
0.16
0.14
0.12
0.1
0.08
0.06
0.04
0.02
0
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0.16
0.18
0.2
1 2 3
INO1
3RXadj
ARS607
ARS305
ARS603
ARS501
INO1
3RXadj
ARS607
ARS305
ARS603
ARS501
% Sin3 Bound
Sin3-HA Binding
21
Although Sin3-Rpd3 origin binding was not detected the possibility remained
that the complex binds origins transiently (or at specific cell cycle times) whereby it
deacetylates histone tails resulting in a more compact chromatin state. Using ChIP
with antibodies specific to acetylated lysine residues on histone tails we examined
changes in histone acetylation at origins in rpd3Δ cells. This analysis detected
acetylation of two to three histones flanking each side of the origin sequences.
Deletion of Rpd3 resulted in a roughly two-fold increase in acetylation of H4 K5 at
many of the origins examined including the early origins ARS305 and ARS1, and
the Rpd3-regulated late origins ARS603, ARS1413, and ARS501 (Figure 7). Little
or no increase in H4 K5 acetylation was observed at the negative control loci SPS2
and 3RXadj or the very early origin ARS607 (Figure 7 and data not shown). Similar
data were observed for H2A K7 (Aparicio, et al., 2004). These data suggest that
Rpd3 deacetylates H4 K5 and H2A K7 at certain early and late origins but it does not
appear that Rpd3 preferentially deacetylates only late origins. Together these data
suggest that although it is difficult to detect Sin3-Rpd3 at origins, the complex does
deacetylate histones adjacent to origins. Moreover, this increased acetylation in the
absence of Rpd3 appears to be more than a general or global effect as acetylation
surrounding ARS607 remains largely unchanged. These data are in agreement with
other studies that suggest Rpd3 deacetylates histones surrounding origins
(Vogelauer, et al., 2002). Little detail is known about the specific acetylation states
of chromatin regions that surround origins and potentially regulate their initiation but
high resolution data of histone modifications at origins, like those generated for yeast
promoters and gene coding sequences, will be important (Liu, et al., 2005).
22
Figure 7. Replication origins are hyperacetylated in the absence of Rpd3. Wild type (DGy166)
and rpd3Δ (JAy72) strains were grown asynchronously and harvested for ChIP analysis using anti-Ac
H4 K5 antibody. Acetylated H4 K5 was measured at each locus using semi-quantitative PCR and the
ratio of immunoprecipitated to input DNA is plotted as “% Bound.”
Rpd3 functions prior to S phase entry to regulate late origins
To further elucidate the mechanism by which Sin3-Rpd3 delays the initiation times
of late origins, we wanted to examine Rpd3 function in a cell cycle and whole-
genome context. Determination of a cell cycle execution point would provide a
window in which to examine Sin3-Rpd3 binding and accompanying acetylation
changes in future studies. To observe the advanced initiation timing of late origins in
rpd3Δ cells in a way that was amenable to cell cycle and genome-wide experiments
we constructed BrdU-Inc strains and measured the incorporation of BrdU at early
and late origins in the presence of HU (Viggiani and Aparicio, 2006). Potentially,
measuring BrdU incorporation at representative origins in the presence of HU would
exacerbate the subtlety of the rpd3Δ phenotype as late origins that fire early enough
to escape the checkpoint would clearly incorporate BrdU compared to origins that
remain properly regulated and silent in wild type cells.
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
INO1
SPS2
ARS607
ARS305
ARS1
ARS603
ARS1413
ARS501
WT
Rpd3
1
0.8
0.7
0.6
0
0.1
0.2
0.3
0.4
0.5
0.9
WT
rpd3Δ
% Bound
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
INO1
SPS2
ARS607
ARS305
ARS1
ARS603
ARS1413
ARS501
H4 K5 Acetylation
INO1 SPS2 ARS607 ARS305 ARS1 ARS603 ARS1413 ARS501
23
First, we characterized origin firing in rpd3Δ cells by immunoprecipitating
BrdU-labeled DNA followed by semi-quantitative PCR amplification of select
origins in the enriched fraction. Cells were arrested in G1 phase and synchronously
released into S phase in media containing HU and BrdU (Figure 8A). In both wild
type and rpd3Δ strains the early origins ARS607, ARS305, and ARS1 incorporated
BrdU with indistinguishable kinetics upon entry into S phase (Figure 8B and data not
shown). As expected, in wild type cells the late origins ARS603 and ARS1413 were
inhibited by the presence of HU and failed to incorporate BrdU above background
levels (Figure 8B). However, in strains lacking Rpd3, we measured significant
incorporation of BrdU at late origins ARS603, ARS1413, ARS501, and ARS716
beginning 36 minutes after release into S phase and peaking at 60 minutes (Figure
8B and data not shown). These data demonstrate that measuring BrdU incorporation
in the presence of HU faithfully characterized the advanced firing times of late
origins in rpd3Δ cells.
The repressive chromatin structure that delays initiation of telomere-proximal
origins is set up between M phase and the G1/S transition (Raghuraman, et al.,
1997). However, no cell cycle studies have identified the execution point for a
specific chromatin-modifying complex required to confer late initiation timing to
origins. We wished to determine if Sin3-Rpd3 was required during S phase to
maintain a repressive chromatin context surrounding certain origins or if it functions
prior to S phase entry, as observed for telomere-proximal origins.
24
Figure 8. Late origins escape HU-induced checkpoint inhibition in rpd3Δ strains as measured
by the incorporation of BrdU. Wild type (CVy43) and rpd3Δ (CVy44) cells containing single
copies of an integrated BrdU-Inc vector were arrested in G1 and released into S phase in media
containing HU and BrdU at 23
o
C. (A) DNA content was measured by FACS analysis. (B) Cells were
harvested at the indicated times for BrdU immunoprecipitation analysis as described in the text.
Quantified PCR products were plotted as the ratio of immunoprecipitated DNA to input DNA (%
BrdU Incorporation).
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0 24 36 48 60 72 84 96
ARS1413
0 24 36 48 60 72 84 96
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
Time (min.)
0
0.05
0.1
0.15
0.2
0.25
0.3
0.35
0.4
0.45
0.5
0 24 36 48 60 72 84 96
ARS603
0
0.1
0.2
0.3
0.4
0.5
0 24 36 48 60 72 84 96
Time (min.)
% BrdU Incorporation
WT rpd3Δ
0
24
36
48
60
72
84
96
12
Time (min.)
0
0.5
1
1.5
2
2.5
3
3.5
4
0 24 36 48 60 72 84 96
ARS607
0
0.5
1
1.5
2
2.5
3
3.5
4
0 24 36 48 60 72 84 96
% BrdU Incorporation
0
0.5
1
1.5
2
2.5
3
3.5
4
0 24 36 48 60 72 84 96
WT
Rpd3
WT
rpd3Δ
A.
B.
25
To this end we constructed a heat-inducible degron strain (Rpd3-td) where an
epitope-tagged version of Rpd3 was fused to a temperature-sensitive mouse DHFR
domain (DHFR-HA-Rpd3) (Dohmen, et al., 1994). DHFR-HA-Rpd3 was placed
under the control of the inducible CUP1 promoter and expressed with the addition of
CuSO
4
to the growth media. Upon shift to a non-permissive temperature (37
o
C), the
DHFR domain unfolds, exposing an N-terminal arginine and internal lysine residues
that are subsequently ubiquitinated, leading to degradation of the entire protein. To
accelerate this process we placed UBR1, which encodes an E3 ubiquitin ligase, under
the inducible GAL1,10 promoter, which is rapidly activated in the presence of
galactose at 30
o
C (Sanchez-Diaz, et al., 2004). To determine the cell cycle execution
point of Rpd3 we could rapidly destroy DHFR-HA-Rpd3 protein in cultures
synchronized at specific points in the cell cycle, then assay for origin function by
measuring BrdU incorporation in the presence of HU.
To test whether DHFR-HA-Rpd3 functioned properly with respect to origin
regulation, the Rpd3-td strain was grown for many generations in the presence or
absence of the Rpd3 fusion protein (Figure 9A and C). Then, cells were
synchronously released from G1 into media containing HU (Figure 9A and B).
Again, the early origins ARS607 and ARS305 incorporated BrdU upon entry into S
phase both when DHFR-HA-Rpd3 was present or absent (Figure 9D). As
anticipated, the long-term presence of the DHFR-HA-Rpd3 (Rpd3-ON-O/N)
effectively inhibited late origins from firing in HU; however, when DHFR-HA-Rpd3
was destroyed for several generations (Rpd3-OFF-O/N) we measured incorporation
26
of BrdU at ARS603, ARS1413, ARS501 and ARS716 indicating advanced initiation
of the representative late origins (Figure 9D). This demonstration of the rpd3Δ
phenotype demonstrates that 1) the DHFR-HA-Rpd3 protein is functional when
present and 2) DHFR-HA-Rpd3 can be degraded to levels low enough to cause
deregulation of origin initiation.
Figure 9. Destruction of Rpd3-td protein for several generations results in late origin
deregulation in HU. (A) Rpd3-td cells (CVy71) were grown overnight at 23
o
C in raffinose and
CuSO
4
to express DHFR-HA-Rpd3 protein (Rpd3-ON-O/N) or in galactose to overexpress Ubr1-myc
(data not shown) to constantly destroy DHFR-HA-Rpd3. Cells were arrested in G1 in their respective
media and released into S phase in the presence of HU and BrdU at 23
o
C. (B) DNA content was
measured at the indicated time points after release from G1 arrest using FACS analysis. (C) DHFR-
HA-Rpd3 protein levels were monitored after overnight (O/N) growth and at indicated time points
after release into HU using immunoblot analysis with anti-HA antibodies. (*) represents a non-
specific cross-reacting band.
DHFR-HA-Rpd3
Raffinose + CuSO
4
Galactose
O/N 0 30 60 90 O/N 0 30 60 90
*
A.
B. C.
Rpd3-ON-O/N
Rpd3-OFF-O/N
HU
BrdU
(23
o
C)
Arrest in G1
aF (23
o
C)
30
0
20
40
50
60
70
80
90
HU HU
Raff + CuSO
4
Galactose
Time
(min.)
Time (min.) Time (min.)
27
Figure 9, continued. Destruction of Rpd3-td protein for several generations results in late origin
deregulation in HU. (D) BrdU immunoprecipitation analysis as described in Figure 8B.
0
0.5
1
1.5
2
2.5
3
3.5
4
0 24 36 48 60 72 84 96
WT
Rpd3
Rpd3-ON-O/N
Rpd3-OFF-O/N
D.
0
1
2
3
4
5
6
7
8
9
0 30 60 90
ARS305
0 30 60 90
0
1
2
3
4
5
6
7
8
9
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0 30 60 90
ARS1413
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0 30 60 90
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0 30 60 90
ARS716
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0 30 60 90
Time (min.)
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0.16
0.18
0.2
0 30 60 90
ARS501
0
0.04
0.08
0.12
0.16
0.2
0 30 60 90
Time (min.)
% BrdU Incorporation
0
0.05
0.1
0.15
0.2
0.25
0 30 60 90
ARS603
0
0.05
0.1
0.15
0.2
0.25
0 30 60 90
% BrdU Incorporation
0
1
2
3
4
5
6
7
8
0 30 60 90
ARS607
0
1
2
3
4
5
6
7
8
0 30 60 90
% BrdU Incorporation
28
Next, we tested whether Rpd3 was required during S phase to regulate origin
timing. The Rpd3-td strain was grown for many generations with DHFR-HA-Rpd3
present, then synchronized in G1. After the population was synchronized in G1, the
culture was split and resuspended in raffinose or galactose to maintain (Rpd3-ON-
G1), or destroy (Rpd3-OFF-G1) the protein, respectively (Figure 10A and C). By
first synchronizing Rpd3-td cells in G1, the DHFR-HA-Rpd3 protein was allowed to
exert its function during other phases of the cell cycle, prior to its destruction. Then
by releasing into S phase we specifically analyzed the Rpd3 requirement during S
phase (Figure 10B and C).
In HU, the early origins ARS607 and ARS305 incorporated BrdU 30 and 60
minutes after release into S phase whether DHFR-HA-Rpd3 was destroyed or
maintained in G1, as expected (Figure 10D). Interestingly however, the destruction
of DHFR-HA-Rpd3 in G1 phase did not advance the initiation timing of late origins
as BrdU incorporation in ARS603, ARS1413, ARS501 and ARS716 did not increase
above background levels in HU (compare Figures 10D and 9D). Despite the fact that
DHFR-HA-Rpd3 was absent throughout S phase, late origins remained silent in HU.
This suggests that Rpd3 performs its origin-regulating function prior to entry into S
phase and implies that once Rpd3 exerts such function its presence is no longer
required to delay the firing of late origins.
29
Figure 10. Rpd3 function prior to S phase entry regulates the initiation timing of late origins.
(A) Rpd3-td cells (CVy71) were grown overnight in raffinose and CuSO
4
to express DHFR-HA-Rpd3
protein. Cells were arrested with αF for 3 hours in raffinose and CuSO
4
, then the culture was split
and cells were resuspended in media containing either raffinose (Rpd3-ON-G1) or galactose (Rpd3-
OFF-G1) as well as αF to maintain a G1 arrest. Both cultures were shifted to 30
o
C, then 37
o
C to
rapidly induce Ubr1-myc and destroy DHFR-HA-Rpd3 in the galactose arm. Cells were released into
S phase in the presence of HU and BrdU at 23
o
C. (B) DNA content was measured by FACS and (C)
induction of Ubr1-myc and destruction of DHFR-HA-Rpd3 were monitored by immunoblot analysis
using anti-myc and anti-HA antibodies respectively.
Rpd3-td
Rpd3-ON
(25
o
C)
Arrest in G1
Rpd3-ON
(25
o
C)
Rpd3-OFF-G1
G1 arrest
(30
o
C)
Rpd3-ON-G1
G1 arrest
(30
o
C)
Rpd3-OFF-G1
HU, BrdU
(23
o
C)
Rpd3-ON-G1
HU, BrdU
(23
o
)
3 hrs 25
o
C
45 min 30
o
C
45 min 37
o
C
45 min 30
o
C
45 min 37
o
C
HU 23
o
C HU 23
o
C
30 60 90 30 60 90
Raff + CuSO
4
Galactose
Ubr1-myc
DHFR-HA-Rpd3
Rpd3-OFF-G1
G1 arrest
(37
o
C)
Rpd3-OFF-G1
G1 arrest
(37
o
C)
A.
B. C.
G1 30
o
C
0
20
30
50
60
80
90
Time (min.)
Raff + CuSO
4
Galactose
G1 G1 G1 G1 G1
Time (min.)
Time (min.)
30
Figure 10, continued. Rpd3 function prior to S phase entry regulates the initiation timing of late
origins. (D) BrdU incorporation was measured at the indicated origins after release into HU as
described.
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0.16
0.18
0.2
0 30 60 90
0
0.5
1
1.5
2
2.5
3
0 30 60 90
0
0.2
0.4
0.6
0.8
1
1.2
1.4
1.6
0 30 60 90
ARS607 ARS305
0
0.05
0.1
0.15
0.2
0.25
0 30 60 90
ARS603
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0 30 60 90
ARS1413
ARS501
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0 30 60 90
ARS716
0
0.5
1
1.5
2
2.5
3
3.5
4
0 24 36 48 60 72 84 96
WT
Rpd3
Rpd3-ON-G1
Rpd3-OFF-G1
0
0.4
0.8
1.2
1.6
0
0.5
1
1.5
2
2.5
3
0
0.05
0.1
0.15
0.2
0.25
0
0.02
0.04
0.06
0.1
0.14
0.08
0.12
0
0.04
0.08
0.12
0.16
0.2
0
0.02
0.04
0.06
0.1
0.14
0.08
0.12
0 30 60 90
D.
0 30 60 90
0 30 60 90 0 30 60 90
0 30 60 90 0 30 60 90
Time (min.) Time (min.)
% BrdU Incorporation % BrdU Incorporation % BrdU Incorporation
31
Large-scale identification of late origins regulated by Rpd3
Next, we wanted to examine how widespread the advanced firing of late origins is in
rpd3Δ cells. We coupled BrdU immunoprecipitation analysis with customized DNA
microarrays representing over 500 known and predicted ARS’s, the entire
chromosome VI and a tiled region of chromosome III to generate replication profiles
for wild type and rpd3Δ cells after 60 minutes in HU (more on array design and
normalization in chapters 4 and 5). Figure 11A shows BrdU incorporation across
chromosome VI. The early origins initiated in both wild type and rpd3Δ cells and
replication forks moved approximately 5 to 10 kb away from each origin, in
agreement with other studies examining fork progression in HU (Katou, et al., 2003;
Tourriere, et al., 2005). Interestingly, ARS608, ARS603 and a recently identified
late origin ARS602.5, incorporate substantially more BrdU in rpd3Δ cells than these
sequences in the wild type strain, where these late origins remained largely (but not
completely) silent (Figure 11A). Similar to early origins, replication forks emanating
from the deregulated late origins in rpd3Δ cells traveled roughly 5 to 10 kb,
suggesting they fired at similar times as the early ARS’s. Replication profiles of
chromosome III for wild type and rpd3Δ were identical, with the early origins
ARS305 and ARS306 firing in both and a cluster of late origins remaining silent,
demonstrating that Rpd3 regulates the timing of some, but not all late origins (data
not shown). In addition, the late origins ARS609, ARS601/2 and the telomere-
32
proximal origin ARS610 also remain silent in cells lacking Rpd3 suggesting that
other factors contribute to the temporal replication program (Figure 11A and data not
shown).
Figure 11. Identification of early replicating regions of chromosome VI in rpd3Δ cells. Wild
type (CVy43) and rpd3Δ (CVy44) cells were grown and measured for BrdU incorporation in the
presence of HU as in Figure 8. (A) Replication profiles of wild type and rpd3Δ cells after 60 minutes
in HU were generated by hybridizing immunoprecipitated BrdU-labeled DNA, coupled with Cy5,
onto DNA microarrays. Input DNA from the 60-minute time point that had not been
immunoprecipitated was labeled with Cy3 and co-hybridized. Samples were amplified and Cy-labeled
using the WGA method described in chapter 5. Arrays were normalized using hybridization
intensities of Drosophila DNA to control spots as described in chapters 4 and 5. (B) BrdU-labeled
regions in rpd3Δ and wild type cells were compared directly by co-hybridizing immunoprecipitated
fractions from each strain. Dye swaps were conducted and used for normalization and the data were
scaled such that log intensities above 0.5 were considered positive. The x-axis represents
chromosome VI coordinates (in kb) with ARS’s labeled in red. Spot fluorescence intensity is plotted
on the y-axis as the log ratio of M/A. The MET10 locus (see text) is at 215,000 kb and marked as (*).
607 608 609 606 605 604 603.5 603 602.5 601/2
A.
B.
607 608 609 606 605 604 603.5 603 602.5 601/2
*
*
WT
rpd3Δ
rpd3Δ
WT
33
To focus on regions that replicate early in cells lacking Rpd3, we directly
compared BrdU-containing DNA from rpd3Δ and wild type cells on the same array.
BrdU-labeled DNA immunoprecipitated from rpd3Δ cells after 60 minutes in HU
was labeled with Cy5 and co-hybridized onto the arrays with Cy3-labeled DNA
immunoprecipitated from wild type. By co-hybridizing the fractions enriched for
BrdU from rpd3Δ and wild type cells, we reasoned that regions that had replicated in
both (ie – early origins) should hybridize in a 1:1 ratio on the array. But regions that
replicated in rpd3Δ but not wild type (ie – early-firing late origins) would show
substantial enrichment.
The rpd3Δ/wild type direct comparison array shows that regions surrounding
ARS602.5, ARS603 and ARS608 show significant enrichment of BrdU, successfully
identifying the same regions of advanced replication timing detected by the
rpd3Δ replication profile (compare Figure 11A and B). The successful identification
of advanced-firing late origins demonstrates the utility of this direct comparison
approach, and importantly, such an approach can easily be expanded to whole-
genome arrays. Interestingly, Rpd3 has been shown to bind the promoter of MET10,
a gene under 5 kb from ARS608 (Robert, et al., 2004). However, Sin3-Rpd3 binding
near ARS603 or ARS602.5 has not been demonstrated. These findings suggest that
while some late origins are regulated by Rpd3 binding to adjacent promoters, late
firing may be conferred to other origins by less direct, or as yet unknown Rpd3-
targeting mechanisms. It will be interesting to determine how widespread late origin
deregulation is in rpd3Δ cells and make genome-wide comparisons between Rpd3’s
DNA replication function and its function as a regulator of transcription.
34
How does Sin3-Rpd3 delay the initiation timing of late origins?
While it has been demonstrated that Sin3-Rpd3 properly delays the initiation timing
of certain late origins, the precise mechanism by which the HDAC complex
accomplishes this is still largely unknown. A working model posits that Sin3-Rpd3
might be targeted to late origins where it would deacetylate histones creating a more
compact chromatin state that delays origin initiation. Here, however, I have
demonstrated that origins are not bound directly by the Sin3-Rpd3 complex (Figure
6). These findings are in agreement with ChIP-chip studies examining Sin3-Rpd3
binding genome-wide, however, these reports tend not to focus on replication origins
(Kurdistani, et al., 2002; Robert, et al., 2004). Still, even without the apparent
binding of Sin3-Rpd3, many origins are characterized by hyperacetylation in cells
where Rpd3 has been deleted, indicating less direct mechanisms may contribute to
the chromatin structures that impact origin initiation timing (Figure 7) (Vogelauer, et
al., 2002).
The data presented here also demonstrate that late origins remain silent in HU
when DHFR-HA-Rpd3 is destroyed at the G1/S transition indicating Rpd3 is not
required during S phase to delay origin timing (Figure 10). Rather, this suggests that
prior to entry into S phase, Sin3-Rpd3 deacetylates histones surrounding origins and
that this compact chromatin state is maintained in the absence of the HDAC
complex, whose chromatin interaction is likely transient. The idea that origins are
“defined” by Sin3-Rpd3 as early or late prior to entry into S phase is in agreement
with studies examining telomere-proximal origins in yeast as well as replication
timing using Xenopus egg extracts, which identify a critical timing decision point
35
(TDP) after mitosis, but prior to S phase (Li, et al., 2001; Raghuraman, et al., 1997).
It is likely that as cells pass through mitosis and enter G1 of a new cell cycle, they
begin to reset critical chromatin structures for transcriptional regulation, and in doing
so, also establish the temporal pattern of DNA replication. Strikingly, the TDP
appears in G1 phase concomitant with the nuclear reorganization that typifies
replication timing: early origins replicate from discrete nuclear foci while late
origins replicate along the nuclear periphery (Li, et al., 2003). Though acetylation
states surrounding origins clearly impinge on origin activity, there is still no direct
evidence demonstrating a causal relationship between chromatin acetylation, nuclear
reorganization and replication timing. Examination of changes in nuclear
organization resulting from RPD3 deletion or overexpression may be useful in
shedding light on this obscure, but important relationship.
The link between replication timing and transcription: at the promoter or
during elongation?
Recent work has identified two Rpd3 complexes that each contain an HDAC core
consisting of Rpd3, Sin3 and Ume1, but associate with different regulatory proteins
that make the complexes functionally distinct (Carrozza, et al., 2005; Joshi and
Struhl, 2005; Keogh, et al., 2005). The large complex, Rpd3C(L), appears to be the
classic silencing complex that is targeted to promoter elements by Ume6 to repress
gene expression. Interestingly, the small complex, Rpd3C(S), which in addition to
the HDAC core contains Eaf3 and Rco1, has been shown to play a role in
remodeling chromatin after transcriptional elongation. As genes are transcribed by
36
RNA polymerase II (RNApII) chromatin is continuously dismantled and
reassembled. To prevent spurious intragenic transcription initiation, chromatin is re-
compacted within coding regions following elongation by RNApII through a
recently described mechanism: The histone methyltransferase, Set2, interacts with a
phosphorylated C-terminal of RNApII and methylates H3 K36, which further
recruits Rpd3C(S) via a chromodomain on Eaf3. By deacetylating histones within
ORF’s, Rpd3C(S) erases the histone acetylation associated with transcription
elongation that can lead to spurious initiation from cryptic start sites (Carrozza, et al.,
2005).
The identification of two functionally distinct Rpd3 complexes presents the
possibility that one complex may function more prominently in the regulation of
replication timing. Indeed, the regulation of some origins appears to be independent
of Rpd3C(L), as deletion of Ume6, which prevents the targeting of Rpd3C(L) but has
no affect on Rpd3C(S), does not advance the initiation timing of ARS603 (Aparicio,
et al., 2004). This implies that a model where delayed origin initiation is the result of
local histone deacetylation due to Rpd3 targeting to an adjacent promoter is unlikely
or incomplete. We have also found no overrepresentation of conserved Ume6
binding sites near late origins, further discrediting the involvement of Rpd3C(L) in
replication timing (Jennifer Aparicio and Oscar Aparicio unpublished observations).
The intriguing possibility remains that DNA replication timing may be linked
to transcriptional elongation through Rpd3C(S). Such a model, where Rpd3C(S)
transiently binds a moving transcription elongation complex, is consistent with data
suggesting that Sin3 binding at origins is difficult to detect, but Rpd3-dependent
37
origin deacetylation is still apparent (Figures 6 and 7). Deletion of Eaf3, which
completely destabilizes Rpd3C(S), results in elevated levels of H3 and H4
acetylation throughout coding regions that is especially pronounced at the 3’ ends of
many genes (Joshi and Struhl, 2005). Therefore, deletion of Rpd3C(S) results in
large patches of hyperacetylation (as opposed to localized hyperacetylation confined
to promoters) that may potentially act over greater distances to advance the initiation
timing of certain late origins. Studies using Drosophila genomic arrays have shown
that replication timing correlates most strongly with transcriptional activity over
large distances (up to 180 kb) as opposed to the transcriptional activity (or changes
in activity) of any single gene (MacAlpine, et al., 2004). Similarly, it is conceivable
that large, intragenic patches of Rpd3C(S)-dependent histone deacetylation may
mark chromosomal regions for late replication timing. These models still require
testing and it is quite possible that both complexes may play some role in the
regulation of specific origins. It is also possible that Rpd3 confers late initiation
timing through an alternative targeting mechanism still yet to be identified.
A genomic approach to Rpd3 origin regulation
The possibility that Rpd3 regulates late origin timing in a more fluid,
Rpd3C(S)-dependent mechanism, means that a search for a classic, causal regulatory
mechanism underlying the temporal replication program may be unsuccessful.
Instead, the relationship between DNA replication, transcriptional regulation and
histone acetylation may be better examined using a genomic approach. Two studies
have examined the genome-wide binding of Sin3 and Rpd3 (Kurdistani, et al., 2002;
38
Robert, et al., 2004). Others have examined gene expression changes resulting from
deletions of Sin3 and Rpd3, as well as disruption of Rpd3C(L) and Rpd3C(S); and
focused analyses have identified dozens of genes regulated by either Rpd3C(L) or
Rpd3C(S) (Bernstein, et al., 2000; Humphrey, et al., 2004; Joshi and Struhl, 2005;
Keogh, et al., 2005). Many acetylation changes in strains lacking Rpd3 have also
been described (Kurdistani, et al., 2004; Vogelauer, et al., 2000). However, our use
of BrdU to successfully identify chromosomal regions under the temporal control of
Rpd3 is the first attempt to map more comprehensively the DNA replication defects
resulting from deletion of an HDAC. While the data presented here are limited to
chromosome VI and a portion of chromosome III, expanding this analysis to include
the entire yeast genome is quite feasible. By taking a genomic view of Rpd3’s
temporal regulation of replication we can begin to compare the role Rpd3 plays in
replication with its functions at other regions in the genome. These studies will be
invaluable in dissecting the link that unites gene expression and replication timing.
Sin3-Rpd3 is one protein complex shown to regulate replication timing, but it is
likely that other factors contribute to the temporal program as well, depending on
gene expression requirements at different chromosomal locations. Notably, SIR
proteins silence telomeric genes and delay initiation timing of telomeric origins
(Stevenson and Gottschling, 1999). Sin3-Rpd3 itself may regulate different origins
through different mechanisms as evidenced by the reported binding of Rpd3 near
ARS608 but not near ARS603 or ARS602.5 (Figure 11).
39
That the temporal replication pattern for different origins is regulated by
different mechanisms is in line with a view where origins themselves are rather fluid
and must rapidly adapt in order to fully replicate the genome in response to changing
cellular environments such as genotoxic stress, failure to initiate early origins, and
altered transcription patters that accompany development or the cellular stress
response. A better understanding of this temporal pattern and its mechanistic
relationship with gene expression is important to understanding the chromosomal
dynamics associated with the critical processes of DNA replication and transcription.
40
CHAPTER 4
BrdU incorporation demonstrates a central role for Mrc1 in replication fork
progression and provides a new tool for studying fork movement across
chromosomes
OVERVIEW
After the initiation of S phase, new DNA is synthesized at replication forks that
travel bidirectionally away from origins by a large replisome complex comprised of
a helicase, DNA polymerases and numerous other regulatory proteins. Faithful
replication of the genome depends on the ability of replication forks to rapidly and
accurately incorporate nucleotides as well as detect and respond to DNA damage. In
this chapter I present evidence that Mrc1, a protein required for replisome integrity
in the presence of exogenous stress, also plays a central, and constitutive role in
replisome function in the absence of replication stress. The elucidation of a general
role for Mrc1 at replication forks provides insight into the mechanisms that promote
genome stability during DNA replication. Further, I provide a novel technique to
analyze replication fork progression at high resolution across chromosomes using
BrdU and DNA microarrays. This powerful procedure holds promise in more
precisely determining how replication forks encounter and repair DNA damage.
Figures 12 and 13 were included in our published Mrc1 study (Szyjka, et al., 2005).
Figure 14 also appears in Weihong Xu’s Ph.D. dissertation, along with a thorough
discussion of normalization procedures (Xu, 2006).
41
RESULTS AND DISCUSSION
Mrc1 is required for the normal rate of replication fork progression throughout
chromatin
The mediator of the replication checkpoint 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 and Elledge, 2003; Tanaka and Russell, 2001). However, cells
lacking Mrc1 progress slowly through an unperturbed S phase as well (Figure 12B)
(Alcasabas, et al., 2001). This prolonged S phase in mrc1Δ cells could be a result of
either 1) a defect in origin initiating or 2) slower moving replication forks. In an
effort to better understand the replication defect in cells lacking Mrc1 we examined
the progression of replication forks by measuring BrdU incorporation along
chromosome VI in wild type and mrc1Δ cells. In addition, we examined a strain
harboring the mrc1-AQ allele, in which hypothetical Mec1 phosphorylation sites
were mutated (Osborn and Elledge, 2003). This allele is defective in checkpoint
signaling to activate Rad53 but appears to retain its function in DNA replication,
based on its approximately normal rate of bulk DNA synthesis (Figure 12B) (Osborn
and Elledge, 2003).
We constructed strains where ARS608 was deleted to allow analysis of a
single replication fork emanating from the early origin ARS607 (Figure 12A). Upon
release into S phase, the incorporation of BrdU at ARS607 in wild type, mrc1Δ, and
mrc1-AQ cells was very similar, occurring primarily between 24 and 36 minutes,
42
suggesting origin initiation was not defective in either mutant (Figure 12C).
However, incorporation of BrdU at sequences 5, 15, and 20 kb from ARS607 was
delayed significantly in mrc1Δ (Figure 12C). In wild type cells, BrdU-labeled DNA
was detected 20 kb distal to ARS607 between 48 and 60 minutes, however in the
mrc1Δ cells, BrdU was not incorporated into this region until 72 minutes (Figure
12C). The kinetics of BrdU incorporation into these sequences was indistinguishable
between wild type and mrc1-AQ cells. This demonstrates that Mrc1, but not its
checkpoint function, is required for replication forks to travel at the proper rate away
from origins. These data (along with two-dimensional gel data, not shown here)
demonstrate that the replication defect observed in mrc1Δ cells is due to slower
progression of replication forks rather than an origin initiation defect (Szyjka, et al.,
2005). Interestingly, this replication defect is independent of Mrc1’s checkpoint
function.
43
Figure 12. Mrc1 is required for normal progression of DNA synthesis. Wild type (CVy39),
mrc1-AQ (CVy40), and mrc1Δ (CVy29) cells with ARS608 deleted were blocked in G1 with α factor
at 23
o
C and released synchronously into S phase at 20
o
C into media containing BrdU. (A) DNA
content analysis was measured by FACS. (B) Cells were collected at the indicated times for analysis
of BrdU incorporation.
ARS607 ARS608
Region A Region B Region C
0 +5kb +15kb +20kb
B.
A.
WT
Time
(min.)
0
24
48
72
96
mrc1Δ
Time
(min.)
0
24
48
72
96
mrc1-AQ
44
Figure 12, continued. Mrc1 is required for normal progression of DNA synthesis.
Immunoprecipitated DNA sequences containing BrdU at ARS607 and distances 5, 15, and 20 kb
away were detected by semi-quantitative PCR amplification. (C) PCR products of
immunoprecipitated BrdU-labeled DNA and input DNA (data not shown) were quantified and plotted
as a ratio of immunoprecipitated DNA to input DNA (% BrdU incorporation).
0.1
1
10
100
24 36 48 60 72 84 96
0.1
1
10
100
24 36 48 60 72 84 96
0.1
1
10
100
24 36 48 60 72 84 96
0.1
1
10
100
24 36 48 60 72 84 96
WT
mrc1-AQ
mrc1
Time (min) Time (min)
ARS607 5 kb
15 kb 20 kb
% BrdU Incorporation
% BrdU Incorporation
Time (min) Time (min)
% BrdU Incorporation
% BrdU Incorporation
Δ
ARS607
5 kb
15 kb
20 kb
ARS607
5 kb
15 kb
20 kb
24 36 48 60 72 84 96 24 36 48 60 72 84 96 24 36 48 60 72 84 96
WT mrc1-AQ mrc1Δ
C.
45
In the presence of replication stress by HU, Mrc1 is required to couple
replication fork components to the site of DNA synthesis (Katou, et al., 2003). We
examined whether, in an unperturbed S phase, Cdc45 progression across chromatin
was also delayed, or whether it might uncouple and move rapidly ahead of slow-
moving replication forks. We examined Cdc45 binding to chromatin using ChIP
after cells were synchronously released into S phase at 16
o
C to enhance the
resolution of the assay. We observed Cdc45 maximally associated with ARS607 at
36 minutes in both wild type and mrc1Δ cells, which supports our conclusion that
deletion of Mrc1 does not impact origin initiation timing (Figure 13). In wild type
cells, Cdc45 traveled away from origins rapidly, associating maximally with regions
5, 15 and 20 kb away from the origin at 72, 72, and 84 minutes, respectively. In
mrc1Δ cells, Cdc45 moved much more slowly, associating maximally with
sequences 5, 15 and 20 kb from ARS607 at 72, 108, and 120 minutes. Therefore,
progression of Cdc45, like that of the replication fork, is slowed in the absence of
Mrc1. This is consistent with Cdc45 remaining coupled to the replication fork even
in the absence of Mrc1 during unperturbed S phase; however, we cannot exclude that
uncoupling from the site of DNA synthesis does occur but is only detected by this
type of analysis when DNA synthesis is greatly slowed by HU.
46
Figure 13. Slower progression of Cdc45 in mrc1Δ cells. Wild type (CVy31) and mrc1Δ (CVy30)
cells, containing Cdc45-HA and ARS608 deleted were blocked in G1 with α factor at 23
o
C and
released synchronously into S phase at 16
o
C. Cells were collected at the indicated times for ChIP
analysis with anti-HA antibody. DNA sequences enriched for Cdc45 binding at ARS607 and at
distances 5, 15 and 20 kb were quantified by PCR amplification. The ratio of immunoprecipitated to
input DNA is plotted as “% Cdc45 Bound.”
To examine replication fork progression in wild type and mrc1Δ cells in a
global manner we examined BrdU incorporation across chromosome VI in strains
lacking the late origins ARS608 and ARS609 using DNA microarrays. BrdU-
labeled regions of chromosome VI were examined in wild type and mrc1Δ cells 36
and 60 minutes after synchronous release into S phase (Figure 14A and B). The
replication profiles clearly show that replication forks move slowly
% Cdc45 Bound
% Cdc45 Bound
Time (min) Time (min)
% Cdc45 Bound
% Cdc45 Bound
Time (min) Time (min)
ARS607 5 kb
15 kb 20 kb
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
24 36 48 60 72 84 96 108 120 132
WT
mrc1
0.02
0.04
0.06
0.08
0.1
0.12
0.14
24 36 48 60 72 84 96 108 120 132
0.02
0.04
0.06
0.08
0.1
24 36 48 60 72 84 96 108 120 132
0.02
0.04
0.06
0.08
0.1
0.12
24 36 48 60 72 84 96 108 120 132
Δ
47
across the entire chromosome in mrc1Δ cells. In wild type cells, after 36 minutes,
BrdU-labeled DNA was detected over 10 kb away from the early origins ARS603.5,
ARS605, ARS606, and ARS607 (Figure 14A, DS normalization). In mrc1Δ cells,
after 36 minutes, these same origins had initiated, but BrdU was only incorporated
into regions about 5 kb away, demonstrating that replication fork progression was
indeed impaired at all forks along the chromosome (Figure 14B, DS normalization).
Figure 14 . Slowed DNA synthesis across chromosome VI in mrc1Δ cells. Wild type (CVy37) and
mrc1Δ (CVy38) cells with ARS608 and ARS609 deleted were blocked in G1 and released into S
phase at 20
o
C in media containing BrdU. BrdU-labeled DNA in (A) wild type and (B) mrc1Δ cells
was immunoprecipitated 36 and 60 minutes after release. Immunoprecipitated and input DNA were
PCR amplified, labeled with Cy dyes and hybridized to DNA microarrays. Data were normalized
using dye swaps (DS) and fluorescence intensity plotted as log M/A. The Hidden Markov model
(HMM) was used to generate plots where probes were determined to incorporate (logM/A = 2.00) or
not incorporate (logM/A = 1.00) BrdU (Xu, 2006).
A.
B.
Wild type
mrc1Δ
36 min.
60 min.
36 min.
60 min.
HMM
HMM
HMM
HMM
DS
DS
DS
DS
48
As expected, delayed fork progression in mrc1Δ cells was also observed at 60
minutes when compared to wild type. In cells lacking Mrc1, forks traveled slowly
away from the early origins (ie- ARS603.5, ARS607), incorporating BrdU
maximally into sequences 10 to 20 kb away (Figure 14B). After 60 minutes, early-
replicating regions in the wild type cells, such as sequences between ARS606 and
ARS607, incorporated BrdU more completely, reflecting more rapidly traveling
replication forks (Figure 14A). Additionally, forks emanating from ARS607 travel
through the ARS608-deleted region. (Figure 14A). In mrc1Δ cells, however, the
region surrounding ARS608 remains largely unreplicated demonstrating replication
forks from ARS607 are impaired (Figure 14B).
Initial limitations of BrdU microarray experiments make replication forks
difficult to track
Interestingly, the data for the 60-minute time points described above are somewhat
less clear than the 36-minute time points. In particular, it was difficult to resolve
fork progression in wild type cells after 60 minutes where BrdU had incorporated
into a greater percentage of the genome (Figure 14A). In this experiment, cells were
released into S phase in media containing BrdU. Under this experimental design,
BrdU was transported into cells and incorporated first into early-replicating
sequences from the large pool of available BrdU. As S phase proceeded, more of the
genome was duplicated and more BrdU was incorporated. This decreased cellular
BrdU pools relative to endogenous dTTP, which likely began to out-compete BrdU
for incorporation into nascent DNA. Indeed, this appears to be the case, as early-
49
replicating sequences show greater BrdU-enrichment than late-replicating sequences
in DNA analyzed from cells allowed to fully complete S phase in the presence of
BrdU (Figure 18 in Appendix A).
Normalization of the data in Figure 14 also presented a challenge and made
accurate analysis of fork progression difficult (see Xu, 2006 for more). Initially, we
normalized each array to the average probe intensity of chromosome VI (data not
shown). However, since the proportion of enriched sequences on chromosome VI
changed across the time course (as more sequences become replicated), this type of
normalization was unhelpful. Coupled with depleting BrdU pools, this analysis
became confounding. Instead, samples were self-normalized using dye swaps.
However, even with this normalization, the baseline signal was not adjusted properly
to zero, yielding many false positives (which appear as a high percentage of enriched
probes across the chromosome). Using the Hidden Markov model (HMM)
individual probes were compared with probes immediately adjacent and then
determined to be either replicated, or not replicated (Xu, 2006). This analysis
significantly clarified the data and greatly reduced false positives (Figure 14,
compare HMM with DS plots). However, false negatives were introduced as large
regions that were likely replicated, such as regions flanking ARS603 (see Figure 14,
WT, 60 minutes), but scored as unreplicated. Again, this was further complicated in
this experiment as late-replicating sequences incorporate BrdU less efficiently and
fail to become significantly enriched above background levels, as determined by this
type of dye swap normalization.
50
BrdU pulse experiments provide improved analysis of the replication fork and
its checkpoint response
We addressed the initial limitations described above by 1) re-designing microarray
slides to contain better internal control probes for normalization and 2) developing a
procedure where BrdU is pulsed into the cultures. The newly-designed microarray
slides contain more potential origins, better coverage of chromosome VI (every ~500
bp), high-resolution coverage (every ~300 bp) of the left arm of chromosome III, and
importantly, several unique probes specific to Drosophila sequences to be used for
normalization (see chapter 5 for more). Briefly, known concentrations of the
Drosophila sequences are amplified along with the experimental DNA from a BrdU
immunoprecipitation (IP) experiment. Together with IP or input DNA, the
Drosophila DNA is labeled with Cy dyes and hybridized to the arrays. During a
BrdU time course experiment the proportion of BrdU-labeled DNA in the genome
changes but the ratio of Drosophila DNA hybridized to the array (and their measured
Cy5/Cy3 fluorescence) remains unchanged. The Drosophila clones account for PCR
amplification biases, Cy-labeling biases and importantly, as they are theoretically in
a 1:1 Cy5 to Cy3 ratio, allow us to set the fluorescence intensity baseline to zero (log
1:1 = 0).
To account for depleting BrdU pools, which occurs as cells incorporate BrdU
into nascent DNA strands during S phase, we developed a BrdU pulse technique.
Using this approach, cells are arrested in G1 and released into media in the absence
of BrdU. As the cells move synchronously into S phase, small cultures are removed
and grown in the presence of BrdU for a short “pulse.” Next, these cells are
51
harvested and genomic DNA is prepared for a BrdU immunoprecipitation. By
pulsing cells with BrdU for short periods of time, only the regions of the genome
replicating in that small window are labeled with BrdU (Figure 15). Using short
pulses of BrdU provides technical advantages by minimizing depletion of BrdU
pools and avoiding potential harmful effects due to prolonged BrdU exposure. It
also offers a superior experimental design by allowing the more precise
determination of when specific sequences are replicated during S phase.
52
Figure 15. BrdU pulses label genomic regions that replicate during discrete windows of time.
Cells are arrested in G1 and synchronously released into S phase in media lacking BrdU. Small
cultures are removed from the main culture at various time points and allowed to grow in BrdU for
short lengths of time (above, 20 minutes). After the short pulse of BrdU, cells are harvested for BrdU
immunoprecipitation analysis. This procedure limits exposure to BrdU, minimizes depletion of BrdU
pools and selectively labels regions of DNA synthesis during a given window of time. BrdU-labeled
regions are indicated in red.
Release in
absence of
BrdU
Early origins fire
BrdU
Late origins fire Forks from early origins proceed
BrdU BrdU
Arrest
in G1
20 40 60 80 100 120 0
BrdU BrdU
53
We employed the new microarrays and the BrdU pulse approach to examine
how replication forks respond differently to 1) replication stress and 2) DNA
damage. First, we examined how forks respond to, and recover from, hydroxyurea
(HU). HU causes replication stress by depleting cellular nucleotide pools, which
results in the activation of the intra-S checkpoint through activation of Rad53 by an
Mrc1-dependent signaling cascade. This cascade greatly slows DNA replication and
arrests cells in mid S phase by stalling replication forks and preventing the initiation
of late-firing origins until nucleotide levels are restored (Melo and Toczyski, 2002).
Wild type strains were arrested in G1 and released into S phase in media containing
HU. Cells were held in HU for 60 minutes and then released into fresh media
without HU and DNA synthesis was monitored using 20-minute BrdU pulses. This
approach allowed us to observe BrdU incorporation during 20-minute windows, both
during the arrest in HU and during the recovery phase as cells were removed from
HU.
Addition of HU greatly slowed progression through S phase as measured
by bulk DNA content (Figure 16A). After 60 minutes in HU, FACS analysis
measured the vast majority of cells still contained 1C DNA content but S phase
proceeded, albeit slowly, after HU was removed. DNA replication was complete
100 minutes after HU release (Figure 16A). As expected, early origins on
chromosome III and chromosome VI initiated replication in the presence of HU
during the 20 to 40-minute pulse, and to a slightly lesser extent during the 40 to 60-
minute pulse (Figure 16B and C). In agreement with previous studies, we observed
54
Figure 16. Replication forks recover slowly after stalling in HU. OAy1026 (ura3:BrdU-
Inc::ars305) cells were blocked in G1 and released into media containing HU for 60 minutes, then
released into fresh media in the absence of HU. (A) DNA content analysis was measured by FACS.
25 mL cultures were removed at the indicated time points and grown in the presence of BrdU for 20
minutes, then collected for BrdU immunoprecipitation analysis. Regions enriched for BrdU-labeled
DNA and input DNA (genomic DNA at t = 0 min.) were PCR amplified, labeled with fluorescent Cy
dyes and hybridized to DNA microarrays. BrdU incorporation was measured across (B) chromosome
III and (C) chromosome VI. Chromosomal coordinates are represented in kilobases and ARS’s are
indicated in red. Fluorescence intensity (log M/A) is plotted on the y-axis.
HU
Release
20 - 40
40 - 60
0 - 20
20 - 40
40 - 60
60 - 80
80 - 100
100 - 120
B. A.
C.
Time
(min.)
HU
Release
20 - 40
40 - 60
0 - 20
20 - 40
40 - 60
60 - 80
80 - 100
100 - 120
Time
(min.) 607 608 609 606 605 604 603.5 603 602.5 601/2
0
120
100
80
60
40
20
60/0
40 HU
Release
Time (min.)
306 305 304 301/2/20
55
that forks from the early origins ARS603.5, ARS605, ARS606, and ARS607 travel
up to about 10 kb in the presence of HU (Katou, et al., 2003; Tourriere, et al., 2005).
In general, late origins remained silent as a result of the activated checkpoint in the
presence of HU (Figure 16B and C). There was little or no incorporation of BrdU at
ARS601/2, ARS603, ARS604, or ARS609 and minimal incorporation at ARS602.5
and ARS608. Using a strain where the highly efficient early origin ARS305 was
deleted, we were able to focus on DNA synthesis at ARS306 and a cluster of late-
firing inefficient origins. As anticipated, in the presence of HU forks stalled
approximately 10 kb from ARS306 and there was limited, although some, BrdU
incorporation at the cluster of late origins and ARS304 (Figure 16B).
When cells were released from HU arrest into fresh media, stalled forks from
early origins resumed elongation almost immediately, but progressed slowly. The 0
to 20-minute pulse, upon release, shows that DNA synthesis resumed rapidly as there
was no longer BrdU incorporation at origins, but sequences flanking the origins
began to incorporate BrdU more robustly (Figure 16B and C). However, nascent
DNA synthesis progressed slowly. Using the high resolution data for chromosome
III, we see that 40 minutes after release from HU (20 to 40-minute pulse), BrdU was
detected maximally 25 kb away from ARS306, suggesting it had traveled 15 kb after
the HU arrest, where forks traveled 10 kb (Figure 16B). This suggests a fork rate of
less than 500 bp/min during this 40-minute period, which is much slower than the
estimated rate (1 to 3 kb/min) of fork progression in unperturbed cells (Szyjka, et al.,
2005; Tercero and Diffley, 2001). During the 40 to 60-minute window, forks from
56
early origins appear to travel more rapidly, but an accurate calculation for fork rate is
difficult as cells begin to lose synchrony and forks from adjacent origins begin to
collide. In addition, during the 40 to 60-minute window, the late origins ARS601/2,
ARS602.5, ARS603, and ARS609 as well as the cluster of late origins on
chromosome III begin incorporating BrdU as those origins initiate after removal of
the checkpoint signal. It is likely that immediately after release from HU, nucleotide
pools are quite low, but as nucleotides are replenished, DNA synthesis is resumed at
higher rates and late origins are able to initiate.
Interestingly, we observed BrdU incorporation at some late origins
(ARS602.5, ARS608, ARS304) even in the presence of HU (Figure 16B and C).
These late origins display fairly robust incorporation and may demonstrate bona fide
origin initiation, which could be tested by examining Cdc45 binding or two-
dimensional electrophoresis. The precise times of replication (Trep) for these origins
are still somewhat obscure but their initiation may be less synchronous throughout a
cell population with the origins firing slightly earlier in some cells but later in others.
A small population of cells may initiate these origins early enough to escape HU-
induced checkpoint regulation. However, replication forks emanating from
ARS602.5, ARS608 and ARS304 are not apparent after release from HU, as would
be expected had they truly initiated (Figure 16B and C). It is unclear from this
analysis whether these data represent non-specific background, low-level initiation
followed by fork collapse, or undetectable levels of actual fork elongation.
However, these data clearly demonstrate the ability to better examine replication fork
progression using high-resolution microarrays and BrdU pulses.
57
Next, we examined the effects of the DNA-damaging agent methyl
methanesulfonate (MMS) on replication fork progression using the same approach.
MMS alkylates bases within DNA, eliciting a DNA damage checkpoint response.
The intra-S phase damage checkpoint is activated through a Rad9-dependent
signaling cascade which, like the HU response, slows the cell cycle by inhibiting late
origin firing and slowing replication forks. Additionally, DNA damage response
genes are upregulated (Melo and Toczyski, 2002). Like the HU experiment
described above, cells were synchronously released into S phase in the presence of
MMS for 1 hour, then released into fresh media and BrdU pulses were used to
examine fork response and recovery. Unlike the stalled S phase observed in HU, we
found that cells proceeded through S phase in the presence of MMS (compare
Figures 16A and 17A). After 60 minutes in MMS, cells progressed substantially into
S phase as measure by bulk DNA content (Figure 17A). This observation is in
agreement with studies demonstrating that forks indeed progress in MMS, and cells
eventually complete S phase (Tercero and Diffley, 2001). Examination of BrdU
incorporation during the 20 to 40-minute window, in the presence of MMS,
demonstrates that early origins on chromosome VI and ARS306 initiated firing
(Figure 17B and C). However, during the 40 to 60-minute pulse, forks continued to
progress, incorporating BrdU up to 25 kb away from recently fired origins, even in
the presence of MMS. Late origins remained largely silent in response to the MMS-
induced checkpoint.
58
Upon release into fresh media, forks continued to proceed and during the 20
to 40-minute pulse, late origin sequences (ARS602.5, ARS603, ARS609, and the
chromosome III cluster) began to incorporate BrdU, indicating their initiation upon
removal of MMS (Figure 17B and C). After 60 minutes, replication of chromosome
VI and the left arm of chromosome III was largely complete, and during the 60 to 80
minute pulse almost no nascent synthesis was detected. This correlates with the
accumulation of 2C bulk DNA content at the same time points (Figure 17A). These
data illustrate that the depletion of nucleotides by HU and DNA damage caused by
MMS exposure elicit two different checkpoint responses. While the lack of
nucleotides in the presence of HU forces forks to arrest and recover slowly, in MMS,
DNA synthesis proceeds, albeit, using a damaged template.
Our data also show that BrdU incorporation is less focused in the presence of
MMS. Rather, the BrdU signal is spread across more sequences, notably the entire
region between ARS603 and ARS603.5, during the 0 to 20 minute release window
(Figure 17C). This may be a result of more rapidly moving forks, resulting in
asynchrony within the population. However, DNA synthesized after MMS insult
contains alkylated bases and possibly lesions on the template that must be repaired.
Therefore, it is possible that the observed spreading out of BrdU signal represents
forks encountering damage or post-replication repair synthesis.
59
Figure 17. Replication forks proceed across a damaged DNA template in the presence of MMS.
OAy1026 (ura3:BrdU-Inc::ars305) cells were blocked in G1 and released into media containing
MMS for 1 hour, then released into fresh media in the absence of MMS. (A) DNA content analysis
was measured by flow cytometry. BrdU was pulsed for 20-minute windows and incorporation was
measured across (B) chromosome III and (C) chromosome VI using microarrays. Chromosome
coordinates are represented in kilobases and ARS’s are represented in red. Fluorescence intensity (log
M/A) is plotted on the y-axis.
MMS
Release
20-40
40-60
0-20
20-40
40-60
60-80
B. A.
C.
Time
(min.) 306 305 304 301/2/20
MMS
Release
20-40
40-60
0-20
20-40
40-60
60-80
Time
(min.)
607 608 609 606 605 604 603.5 603 602.5 601/2
0
120
100
80
60
40
20
60/0
40 MMS
Release
Time (min.)
60
A central role for Mrc1 at replication forks
Previous analysis of mrc1Δ 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 has
been shown to physically interact with Cdc45, Mcm2 and Mcm3 (Katou, et al., 2003;
Nedelcheva, et al., 2005). S. pombe Mrc1, and its human homologue Claspin, have
been shown to bind branched DNA structure in vitro (Sar, et al., 2004; Zhao and
Russell, 2004). These data, together with ours, suggest that Mrc1 links Cdc45 and/or
Mcm proteins to replication forks in vivo.
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 on their physical association (Dong, et al., 1996; Kim, et al.,
1996). If the loss of Mrc1 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
61
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 elucidation of a general role of Mrc1 in the function
of replication forks provides new insight into the mechanisms that promote genome
stability during DNA replication.
High-resolution BrdU pulse experiments effectively analyze replication forks
Initial microarray analysis of BrdU-labeled regions of chromosome VI was sufficient
to characterize slow moving replication forks in mrc1Δ cells. However, by using
BrdU pulse experiments in combination with microarrays we were able to more
clearly examine replication forks as they responded to HU and MMS treatments.
This technique, in conjunction with standard ChIP-chip analysis will be an important
tool in better defining protein dynamics at replication forks. In particular, how do
replication forks respond to damage in vivo? Initial work in E. coli suggested that
UV-induced damage on DNA templates was bypassed by replication forks, leaving
gaps that were subsequently repaired (Rupp and Howard-Flanders, 1968). While
DNA polymerases known to carry out translesion synthesis (TLS) have been
discovered, filling these gaps would require re-priming events that were thought to
be acceptable on the discontinuously synthesized lagging strand, but inconceivable
on the continuously synthesized leading strand. Therefore, most current models of
repair suggest that TLS takes place at replication forks (Langston and O'Donnell,
62
2006; Lehmann and Fuchs, 2006). However, growing evidence suggests that leading
strands can indeed be re-primed and that leading strand synthesis may in fact, be
partially discontinuous (Amado and Kuzminov, 2006; Heller and Marians, 2006). In
addition, using electron microscopy Lopes et al. have identified gaps behind
replication forks in UV-irradiated S. cerevisiae suggesting a damage bypass
mechanism (Lopes, et al., 2006). To gain better insight into the mechanism of DNA
repair in vivo, it will be useful to more accurately map where important replication
fork proteins, such as Cdc45, MCM proteins, and RPA, are located in the presence of
damaging agents. Additionally, the examination of the localization of TLS
polymerase subunits Rad30, Rev1 and Rev3, in combination with a BrdU pulse
approach may help illuminate where DNA repair occurs relative to the replication
fork.
63
CHAPTER 5
Materials and methods
Yeast methods and strain construction
Strains were derived from W303 and are described in Table 3. LiOAc
transformations were used for genomic integrations of linearized plasmids or PCR-
amplified cassettes for gene deletions or epitope taggings. Strains were routinely
confirmed using PCR directed at specific loci or immunoblot analysis to detect
tagged proteins. Chromosomal integrations of the pBrdU-Inc constructs were tested
as described in chapter 2 using primer sequences listed in Table 2 and standard PCR
amplification conditions as follows: an initial 3 minute 95
o
C denaturation, followed
by 30 cycles of 30 seconds at 95
o
C, 30 seconds annealing at 59
o
C, and a 1 minute
and 45 second extension at 72
o
C, all followed by a final 4 minute extension step at
72
o
C. Primers RS1 and RS2 were used in combination with the specific HIS3, LEU2
and URA3 primers to test integrations at those loci. Primer RS1.1 was designed to
test p404-BrdU-Inc integrations at the trp1 locus and was used in place of primer
RS1.
64
Yeast strains were generally cultured in selective media and overnight prior
to an experiment in YEP with 2% of the appropriate sugar source. Cells were
synchronized in G1 phase using the mating pheromone α factor (1:10,000) typically
at 23
o
C for ~4 hours. Rpd3-td strains were grown in YEP raffinose plus CuSO
4
or
galactose where indicated and manipulated as described in Figures 9 and 10.
Overnight growth of Rpd3-td in galactose at 23
o
C sufficiently destroyed DHFR-HA-
Rpd3 while it was stable overnight in raffinose and CuSO
4
. When indicated, cells
were grown in 200 mM hydroxyrea and 0.033% MMS.
Table 2. Primer sequences for confirmation of BrdU-Inc constructions
Primer Sequence (5' to 3')
RS1 tgaaaacctctgacacatgcag
RS1.1* ggcatcagagcagattgtactgag
RS2 cttgattagggtgatggttcacg
5'His3 ctactattgctttgctgtggg
3'His3 gccacctatcaccacaactaac
5'Trp1 ctaaaagagctgacagggaaatgg
3'Trp1 cttgcttttcaaaaggcctgc
5'Leu2 gcagattcccttttatggattcc
3'Leu2 ggtagatttagtactgaagaggaggtcg
5'Ura3 ggctgtggtttcagggtccataaagc
3'Ura3 gtcattatagaaatcattacgaccgagattccc
*RS1.1 should be used to test the BrdU-Inc integration at the trp locus
65
Table 3. Strain table
Strain Genotype Source
E1000 MATa ade2-1, trp1-1, can1-100, leu2-
3,112, his3-11,15, ura3::GPD-TK(7x)
Lengronne et al. 2001
OAy470 MATa ade2-1, trp1-1, can1-100, leu2-
3,112, his3-11,15, bar1 Δ:: hisG
Aparicio et al. 2004
JAy22 OAy470 + rpd3 Δ ::HIS5 Aparicio et al. 2004
JAy57 OAy470 + Sin3-HA3::KanMX Aparicio et al. 2004
SSy161 MATa ade2-1 ura3-1 his3-11,15 trp1-1
leu2-3,112 can1-100 bar1 Δ ::hisG
Szyjka et al. 2005
CVy29 E1000 + mrc1 Δ ::KanMX ,
ars608 Δ ::his3 , bar1 Δ ::leu2
Szyjka et al. 2005
CVy30 E1000 + mrc1D ::KanMX ,
ars608 Δ ::his3 , bar1 Δ ::leu2 , Cdc45-
HA3::trp1
Szyjka et al. 2005
CVy31 E1000 + ars608 Δ ::his3 ,
bar1 Δ ::leu2 , Cdc45-HA3::trp1
Szyjka et al. 2005
CVy39 E1000 + ars608 ::his3 , bar1 Δ ::trp1 Szyjka et al. 2005
CVy40 CVy39 + mrc1-AQ::leu2:mrc1 Δ :his5 Szyjka et al. 2005
CVy43 SSy161 + ura3 ::BrdU-Inc Viggiani et al. 2006
CVy44 CVy43 + rpd3 Δ ::KanMX This study
CVy59 SSy161 + his3 ::BrdU-Inc Viggiani et al. 2006
CVy61 SSy161 + trp1 ::BrdU-Inc Viggiani et al. 2006
CVy63 SSy161 + leu2 ::BrdU-Inc Viggiani et al. 2006
CVy71 SSy161 + leu2 ::BrdU-Inc, Gal1,10-
Ubr1-myc ::his5 , CuP-Ub-DHFR-HA-
Rpd3 ::ura3
This study
OAy1026 SSy161 + ura3:BrdU-Inc::ars305 This study
66
Construction of BrdU-Inc plasmids
HSV-TK, under the control of the strong constitutive GPD promoter, and hENT1,
under the control of the ADH1 promoter, were placed in a back-to-back orientation
in each of the four integrating pRS vectors (Sikorski and Hieter, 1989). hENT1 was
amplified from pYhENT1 (from C. Cass) using the Expand High-Fidelity PCR
System (Roche) and the following primers (Vickers, et al., 1999): 5’-
GTAGATCTATGACAACCAGTCACCAGCCT, and 5’-
GCTGCGGCCGCCTCGAGTCACACAATTGCCCGGAA. The hENT1 PCR
product was cut with BglII and NotI and ligated into pRS306 cut with BamHI and
NotI. The ADH1 promoter was excised from Yeplac112-Rpd3-Flag (from K. Struhl)
using SalI and ClaI and placed in front of hENT1 using those same enzymes to
create p306-ADH1-hENT1. The GPD-HSV-TK fragment was excised from pGPD-
TK (from E. Schwob) using SalI and KpnI and inserted into this new plasmid with
the same enzymes, resulting in p306-BrdU-Inc (Lengronne, et al., 2001). This entire
BrdU incorporation cassette (BrdU-Inc) was excised with KpnI and NotI and inserted
into pRS404 cut with KpnI and NotI. The resulting plasmid was subjected to partial
digestion with XbaI followed by fill-in and ligation to eliminate the XbaI site in the
polylinker and leave a single XbaI site in the TRP1 gene for genomic integration,
yielding p404-BrdU-Inc.
A three-way ligation was used to construct the LEU2 and HIS3 vectors. The
SalI-NotI ADH1-hENT1 fragment and the SalI-SacI GPD-HSV-TK fragment were
simultaneously ligated into pRS405 and pRS403 each cut with SacI and NotI,
67
creating p405-BrdU-Inc and p403-BrdU-Inc, respectively. Plasmid constructions
were confirmed with PCR analysis, restriction mapping, and DNA sequencing of the
BrdU-Inc cassette (data not shown).
Immunoblot analysis
To detect BrdU-labeled DNA using immunoblotting (Figures 4 and 5), 5 ml cultures
were grown into log-phase in selectable media at 30°C. Cells were harvested and
resuspended in YEPD containing 400 µg/ml of BrdU for 1 hour and genomic DNA
was isolated using standard glass bead and phenol chloroform (PCI) extraction
methods and resuspended in 100 µl TE. Samples were quantified on an agarose gel
or by UV-spectroscopy (Nanodrop). The indicated amounts of DNA were diluted to
500 µl with 5xSSC and denatured at 95°C for 10 minutes. Samples were snap-
cooled in an ice water bath, applied to a nylon membrane (Hybond N+, GE
Healthcare) equilibrated in 5xSSC using a slot blot Minifold II (Schleicher and
Schuell), and UV-crosslinked (Stratalinker, 1.2x10
5
µJoules). The membrane was
blocked with 5% milk in TBS containing 0.1% Tween-20 for 20 minutes, and probed
sequentially with anti-BrdU antibody (Oxford, 1:1000) for 30 minutes and, following
washes, with HRP-conjugated anti-rat secondary antibody for 30 minutes (Santa
Cruz, 1:5000). Chemiluminescence was detected (SuperSignal, Pierce) using a
Chemidoc (BioRad) and quantified using Quantity One software.
Immunoblot analysis of tagged proteins was conducted similarly. Proteins
were prepared using TCA precipitations and separated by 10% SDS-PAGE. Proteins
were transferred to nitrocellulose using TransferBlot SD Semi-Dry Transfer Cell
68
(BioRad) at a constant voltage of 20 V for 30 minutes (up to 1 hour for the large
Ubr1-myc protein). The membrane was blocked in milk as above and probed for at
least 1 hour using the appropriate antibody. DHFR-HA-Rpd3 was detected using
anti-HA (16B12 at 1:1000). Ubr1-myc was detected using anti-myc (9e10 at
1:1000). Anti-mouse secondary was used at 1:5000.
ChIP and BrdU immunoprecipitations
ChIP analyses of HA-tagged proteins and acetylated H4 K5 and H2A K7 were
performed as previously described (Aparicio, et al., 2004; Szyjka, et al., 2005).
Percent bound is the ratio of immunoprecipitated to total DNA based on the
quantification of the respective PCR products and appropriate sample dilution
factors.
For BrdU immunoprecipitations, cells were grown in media containing 400
µg/mL of BrdU or pulsed with 400 µg/mL BrdU as discussed in chapter 4. 20 mL of
cells were harvested in 0.1% NaN
3
, washed in ice cold PBS and resuspended in 500
µl of genomic lysis buffer (100 mM Tris pH=8, 50 mM EDTA, 1% SDS). Cell
lysates were prepared by vigorous bead beating using a FastPrep FP120 (two 45
second pulses at power 5.5 yielded over 90% cell breakage). Genomic DNA was
sheared for 45 seconds then purified by a PCI extraction followed by RNase and
Proteinase K treatment, followed by a second PCI extraction and finally cleaned over
a QiaQuick PCR Purification Column (QiaGen) and resuspended in 50 µL TE. 40
µL of this DNA (typically 1 to 5 µg) was combined with 20 µg of sheared salmon
sperm DNA and denatured for 15 minutes at 95
o
C. This denatured DNA was snap
69
cooled in an ice water bath and ice cold PBS/0.1% Triton was added to a total
volume of 200 µL and incubated with anti-BrdU antibody (GE Healthcare, 0.5 µl
antibody in 200 µl total reaction volume) overnight at 4
o
C, followed by G sepharose
beads for 1 hour. Beads were washed twice in PBS/0.1% Triton and once in TE,
then eluted in TE/1%SDS at 65
o
C for 15 minutes. Immunoprecipitated BrdU-labeled
DNA was purified (QiaQuick Column) and amplified using 22 to 24 cycles of PCR.
Input DNA that was not immunoprecipitated was diluted 1:100 and subject to the
same PCR analysis. For Figures 8, 9, and 10, input DNA was un-
immunoprecipitated genomic DNA at each time point. For microarray experiments,
unless otherwise noted, input DNA for each time point was un-immunoprecipitated
genomic DNA from the t = 0 minute time point. Percent BrdU incorporated is the
ratio of the immunoprecipitated to input DNA based on quantification of respective
PCR products.
DNA microarray sample amplification and labeling, array design and analysis
BrdU-labeled DNA was immunoprecipitated, amplified and labeled with fluorescent
dyes for microarrays using ligation-mediated PCR initially as described by DeRisi’s
Protocol (Figures 14, 16, 17, 18), and subsequently using Sigma’s GenomePlex
Whole Genome Amplification kit (WGA) (Figure 11). Following the DeRisi
protocol, samples were subjected to a series of PCR amplification steps; Round A, B,
and C. In Round A, primers that will serve as a template in subsequent reactions,
were annealed using Sequenase. This was conducted on ~100 ng (in 7 µl) of IP
DNA and ~50 ng input DNA. In Round B a specific primer amplifies the Round A
70
templates. In Round C amino-allyl dUTP was incorporated. We currently prefer the
WGA method. Following the WGA protocol, ~10 ng of IP or input DNA was
subjected to the Library Preparation step followed by a 16 cycle amplification round
resulting in ~2 to 6 µg of DNA. Re-amplification steps were conducted as needed
and as outlined in the protocol (15 ng from first amplification was re-amplified with
14 cycles). 3 µg of amplified DNA was subjected to incorporation of aa-dUTP using
a Klenow extension reaction and random nonamers at 37
o
C for 4 hours, typically
doubling the amount of DNA. Using both methods, 0.125 µl of Drosophila DNA
was added to the initial amplification steps and amplified along with the sample
DNA.
All samples were coupled with Cy3 and Cy5 dyes identically, regardless of
how they were amplified. Samples were first cleaned using a QiaQuick PCR
purification column (Qiagen) without Tris buffers. Buffer PB was used as usual but
the column was washed with 80% ethanol and the sample was eluted in water then
vacuum dried and resuspended in 10 µl milli-Q dH
2
O and 1 µl of freshly-made 1M
sodium bicarbonate, pH 9.0. This buffered DNA solution was vigorously mixed
with the appropriate dried Cy-dye aliquot and incubated for 1 hour at room
temperature. Un-coupled material was removed using a QiaQuick PCR purification
column (Qiagen) with the following modifications: 35 µl of 3M NaOAc was added
to the coupling reaction, followed by 250 µl of Buffer PB and two sequential washes
with 750 µl of Buffer PE. The sample was eluted in 40 µl of Buffer EB, pre-warmed
to 55
o
C. Sample concentrations and FOI’s were calculated using spectrophotometry
(NanoDrop) prior to their appropriate pooling and hybridization.
71
In all cases 1 µg of IP and input DNA were co-hybridized together, overnight
at 50
o
C. The appropriate amounts of Cy-labeled IP and input DNA were combined
and vacuum-dried, then resuspended in 5 µl of 10 mM EDTA and denatured at 95
o
C
for 1 minute. Then, 50 µl of pre-warmed (68
o
C) Hybridization Buffer (30%
formamide, 5X SSC, 0.1% SDS, 100 µg/ml ssssDNA) was added to the sample and
it was denatured again for 2 minutes prior to its addition to the array. After the
hybridization was complete, slides were washed of non-specific background through
a series of sequential washes: a 5-minute wash in 45
o
C 1X SSC, 0.1%SDS, 1 mM
DTT, a dip in room temperature 0.2X SSC, 1mM DTT, and two 3-minute washes in
room temperature 0.1X SSC, 1 mM DTT. Slides were dried using a centrifuge and
scanned with a GenePix 4100A (Axon Instruments).
DNA microarray slides contained approximately 2100 isothermal probes (60
bp in length) representing the S. cerevisiae genome, each spotted 5 times. The arrays
represent the following sequences: 588 probes for confirmed and predicted ARS
elements, 894 probes tiling chromosome VI approximately every 270 bp, 607 probes
tiling the left arm of chromosome III approximately every 100 bp, 8 negative control
probes for pbluescript, and 50 probes representing unique D. melanogaster DNA for
normalization. Other probes represent non-origins sequences. Probes tiling
chromosomes III and VI were designed using OligArray software.
Array normalization is discussed extensively elsewhere (Xu, 2006; Xu, et al.,
2006). Briefly, the Cy5 and Cy3 foreground signals were converted to log ratio of
enrichment defined as M = log
2
Cy5 – log
2
Cy3 to log intensity defined as A =
(log
2
Cy5 + log
2
Cy3)/2 for each spot. D. melanogaster clones (DC) were amplified
72
and labeled along with enriched and input DNA as described in chapter 4. Using this
approach BrdU-labeled DNA was immunoprecipitated, amplified, and labeled with
Cy5, whereas input DNA was labeled with Cy3. After hybridization to the arrays,
probes with high Cy5 intensities represent sequences containing BrdU and the log
ratio of the Cy5/Cy3 intensity is positive. Probes with high Cy3 intensities were not
enriched for BrdU-labeled DNA relative to the input, and therefore not replicated.
These sequences yield a negative Cy5/Cy3 log ratio, placing them below the baseline
generated by the Drosophila clones (which are theoretically present in a 1:1 ratio).
73
REFERENCES
Aggarwal, B. D. and Calvi, B. R. (2004). Chromatin regulates origin activity in
Drosophila follicle cells. Nature 430, 372-6.
Alcasabas, A. A., Osborn, A. J., Bachant, J., Hu, F., Werler, P. J., Bousset, K.,
Furuya, K., Diffley, J. F., Carr, A. M. and Elledge, S. J. (2001). Mrc1
transduces signals of DNA replication stress to activate Rad53. Nat Cell Biol
3, 958-65.
Amado, L. and Kuzminov, A. (2006). The replication intermediates in Escherichia
coli are not the product of DNA processing or uracil excision. J Biol Chem
281, 22635-46.
Aparicio, J. G., Viggiani, C. J., Gibson, D. G. and Aparicio, O. M. (2004). The
Rpd3-Sin3 histone deacetylase regulates replication timing and enables intra-
S origin control in Saccharomyces cerevisiae. Mol Cell Biol 24, 4769-80.
Bell, S. P. (1995). Eukaryotic replicators and associated protein complexes. Curr
Opin Genet Dev 5, 162-7.
Bell, S. P. and Dutta, A. (2002). DNA replication in eukaryotic cells. Annu Rev
Biochem 71, 333-74.
Bernstein, B. E., Tong, J. K. and Schreiber, S. L. (2000). Genomewide studies of
histone deacetylase function in yeast. Proc Natl Acad Sci U S A 97, 13708-
13.
Burgess, S. M., Ajimura, M. and Kleckner, N. (1999). GCN5-dependent histone H3
acetylation and RPD3-dependent histone H4 deacetylation have distinct,
opposing effects on IME2 transcription, during meiosis and during vegetative
growth, in budding yeast. Proc Natl Acad Sci U S A 96, 6835-40.
Carrozza, M. J., Li, B., Florens, L., Suganuma, T., Swanson, S. K., Lee, K. K., Shia,
W. J., Anderson, S., Yates, J., Washburn, M. P. and Workman, J. L. (2005).
Histone H3 methylation by Set2 directs deacetylation of coding regions by
Rpd3S to suppress spurious intragenic transcription. Cell 123, 581-92.
74
Cimbora, D. M., Schubeler, D., Reik, A., Hamilton, J., Francastel, C., Epner, E. M.
and Groudine, M. (2000). Long-distance control of origin choice and
replication timing in the human beta-globin locus are independent of the
locus control region. Mol Cell Biol 20, 5581-91.
De Nadal, E., Zapater, M., Alepuz, P. M., Sumoy, L., Mas, G. and Posas, F. (2004).
The MAPK Hog1 recruits Rpd3 histone deacetylase to activate
osmoresponsive genes. Nature 427, 370-4.
Dimitrova, D. S. and Gilbert, D. M. (1999). The spatial position and replication
timing of chromosomal domains are both established in early G1 phase. Mol
Cell 4, 983-93.
Dohmen, R. J., Wu, P. and Varshavsky, A. (1994). Heat-inducible degron: a method
for constructing temperature-sensitive mutants. Science 263, 1273-6.
Dolbeare, F. (1996). Bromodeoxyuridine: a diagnostic tool in biology and medicine,
Part III. Proliferation in normal, injured and diseased tissue, growth factors,
differentiation, DNA replication sites and in situ hybridization. Histochem J
28, 531-75.
Dong, F., Weitzel, S. E. and von Hippel, P. H. (1996). A coupled complex of T4
DNA replication helicase (gp41) and polymerase (gp43) can perform rapid
and processive DNA strand-displacement synthesis. Proc Natl Acad Sci U S
A 93, 14456-61.
Feng, W., Collingwood, D., Boeck, M. E., Fox, L. A., Alvino, G. M., Fangman, W.
L., Raghuraman, M. K. and Brewer, B. J. (2006). Genomic mapping of
single-stranded DNA in hydroxyurea-challenged yeasts identifies origins of
replication. Nat Cell Biol 8, 148-55.
Ferguson, B. M. and Fangman, W. L. (1992). A position effect on the time of
replication origin activation in yeast. Cell 68, 333-9.
Friedman, K. L., Diller, J. D., Ferguson, B. M., Nyland, S. V., Brewer, B. J. and
Fangman, W. L. (1996). Multiple determinants controlling activation of yeast
replication origins late in S phase. Genes Dev 10, 1595-607.
75
Gavin, A. C., Bosche, M., Krause, R., Grandi, P., Marzioch, M., Bauer, A., Schultz,
J., Rick, J. M., Michon, A. M., Cruciat, C. M., Remor, M., Hofert, C.,
Schelder, M., Brajenovic, M., Ruffner, H., Merino, A., Klein, K., Hudak, M.,
Dickson, D., Rudi, T., Gnau, V., Bauch, A., Bastuck, S., Huhse, B.,
Leutwein, C., Heurtier, M. A., Copley, R. R., Edelmann, A., Querfurth, E.,
Rybin, V., Drewes, G., Raida, M., Bouwmeester, T., Bork, P., Seraphin, B.,
Kuster, B., Neubauer, G. and Superti-Furga, G. (2002). Functional
organization of the yeast proteome by systematic analysis of protein
complexes. Nature 415, 141-7.
Gilbert, D. M. (2002). Replication timing and transcriptional control: beyond cause
and effect. Curr Opin Cell Biol 14, 377-83.
Goren, A. and Cedar, H. (2003). Replicating by the clock. Nat Rev Mol Cell Biol 4,
25-32.
Grivell, A. R. and Jackson, J. F. (1968). Thymidine kinase: evidence for its absence
from Neurospora crassa and some other micro-organisms, and the relevance
of this to the specific labelling of deoxyribonucleic acid. J Gen Microbiol 54,
307-17.
Guarente, L., Schneider J.C. (1991). Vectors for expression of cloned genes in yeast:
regulation, overproduction, and underproduction. In Methods in Enzymology,
Fink, G.R., and Guthrie, C. (ed.). Academic Press: New York.
Heller, R. C. and Marians, K. J. (2006). Replication fork reactivation downstream of
a blocked nascent leading strand. Nature 439, 557-62.
Hodson, J. A., Bailis, J. M. and Forsburg, S. L. (2003). Efficient labeling of fission
yeast Schizosaccharomyces pombe with thymidine and BUdR. Nucleic Acids
Res 31, e134.
Humphrey, E. L., Shamji, A. F., Bernstein, B. E. and Schreiber, S. L. (2004). Rpd3p
relocation mediates a transcriptional response to rapamycin in yeast. Chem
Biol 11, 295-9.
Joshi, A. A. and Struhl, K. (2005). Eaf3 chromodomain interaction with methylated
H3-K36 links histone deacetylation to Pol II elongation. Mol Cell 20, 971-8.
76
Kadosh, D. and Struhl, K. (1997). Repression by Ume6 involves recruitment of a
complex containing Sin3 corepressor and Rpd3 histone deacetylase to target
promoters. Cell 89, 365-71.
Kadosh, D. and Struhl, K. (1998). Targeted recruitment of the Sin3-Rpd3 histone
deacetylase complex generates a highly localized domain of repressed
chromatin in vivo. Mol Cell Biol 18, 5121-7.
Katou, Y., Kanoh, Y., Bando, M., Noguchi, H., Tanaka, H., Ashikari, T., Sugimoto,
K. and Shirahige, K. (2003). S-phase checkpoint proteins Tof1 and Mrc1
form a stable replication-pausing complex. Nature 424, 1078-83.
Keogh, M. C., Kurdistani, S. K., Morris, S. A., Ahn, S. H., Podolny, V., Collins, S.
R., Schuldiner, M., Chin, K., Punna, T., Thompson, N. J., Boone, C., Emili,
A., Weissman, J. S., Hughes, T. R., Strahl, B. D., Grunstein, M., Greenblatt,
J. F., Buratowski, S. and Krogan, N. J. (2005). Cotranscriptional set2
methylation of histone H3 lysine 36 recruits a repressive Rpd3 complex. Cell
123, 593-605.
Kim, S., Dallmann, H. G., McHenry, C. S. and Marians, K. J. (1996). Coupling of a
replicative polymerase and helicase: a tau-DnaB interaction mediates rapid
replication fork movement. Cell 84, 643-50.
Kurdistani, S. K., Robyr, D., Tavazoie, S. and Grunstein, M. (2002). Genome-wide
binding map of the histone deacetylase Rpd3 in yeast. Nat Genet 31, 248-54.
Kurdistani, S. K., Tavazoie, S. and Grunstein, M. (2004). Mapping global histone
acetylation patterns to gene expression. Cell 117, 721-33.
Langston, L. D. and O'Donnell, M. (2006). DNA replication: keep moving and don't
mind the gap. Mol Cell 23, 155-60.
Leff, J. and Lam, K. B. (1976). Bromodeoxyuridine 5'-monophosphate incorporation
into yeast nuclear and mitochondrial deoxyribonucleic acid. J Bacteriol 127,
354-61.
Lehmann, A. R. and Fuchs, R. P. (2006). Gaps and forks in DNA replication:
Rediscovering old models. DNA Repair (Amst) 5, 1495-8.
77
Lengronne, A., Pasero, P., Bensimon, A. and Schwob, E. (2001). Monitoring S phase
progression globally and locally using BrdU incorporation in TK(+) yeast
strains. Nucleic Acids Res 29, 1433-42.
Li, F., Chen, J., Izumi, M., Butler, M. C., Keezer, S. M. and Gilbert, D. M. (2001).
The replication timing program of the Chinese hamster beta-globin locus is
established coincident with its repositioning near peripheral heterochromatin
in early G1 phase. J Cell Biol 154, 283-92.
Li, F., Chen, J., Solessio, E. and Gilbert, D. M. (2003). Spatial distribution and
specification of mammalian replication origins during G1 phase. J Cell Biol
161, 257-66.
Liu, C. L., Kaplan, T., Kim, M., Buratowski, S., Schreiber, S. L., Friedman, N. and
Rando, O. J. (2005). Single-nucleosome mapping of histone modifications in
S. cerevisiae. PLoS Biol 3, e328.
Lopes, M., Foiani, M. and Sogo, J. M. (2006). Multiple mechanisms control
chromosome integrity after replication fork uncoupling and restart at
irreparable UV lesions. Mol Cell 21, 15-27.
MacAlpine, D. M., Rodriguez, H. K. and Bell, S. P. (2004). Coordination of
replication and transcription along a Drosophila chromosome. Genes Dev 18,
3094-105.
Machida, Y. J., Hamlin, J. L. and Dutta, A. (2005). Right place, right time, and only
once: replication initiation in metazoans. Cell 123, 13-24.
McNeil, J. B. and Friesen, J. D. (1981). Expression of the Herpes simplex virus
thymidine kinase gene in Saccharomyces cerevisiae. Mol Gen Genet 184,
386-93.
Melo, J. and Toczyski, D. (2002). A unified view of the DNA-damage checkpoint.
Curr Opin Cell Biol 14, 237-45.
Nedelcheva, M. N., Roguev, A., Dolapchiev, L. B., Shevchenko, A., Taskov, H. B.,
Stewart, A. F. and Stoynov, S. S. (2005). Uncoupling of unwinding from
DNA synthesis implies regulation of MCM helicase by Tof1/Mrc1/Csm3
checkpoint complex. J Mol Biol 347, 509-21.
78
Newlon, C. S. and Theis, J. F. (2002). DNA replication joins the revolution: whole-
genome views of DNA replication in budding yeast. Bioessays 24, 300-4.
O'Neill B, M., Szyjka, S. J., Lis, E. T., Bailey, A. O., Yates, J. R., 3rd, Aparicio, O.
M. and Romesberg, F. E. (2007). Pph3-Psy2 is a phosphatase complex
required for Rad53 dephosphorylation and replication fork restart during
recovery from DNA damage. Proc Natl Acad Sci U S A.
Osborn, A. J. and Elledge, S. J. (2003). Mrc1 is a replication fork component whose
phosphorylation in response to DNA replication stress activates Rad53.
Genes Dev 17, 1755-67.
Raghuraman, M. K., Brewer, B. J. and Fangman, W. L. (1997). Cell cycle-dependent
establishment of a late replication program. Science 276, 806-9.
Raghuraman, M. K., Winzeler, E. A., Collingwood, D., Hunt, S., Wodicka, L.,
Conway, A., Lockhart, D. J., Davis, R. W., Brewer, B. J. and Fangman, W. L.
(2001). Replication dynamics of the yeast genome. Science 294, 115-21.
Robert, F., Pokholok, D. K., Hannett, N. M., Rinaldi, N. J., Chandy, M., Rolfe, A.,
Workman, J. L., Gifford, D. K. and Young, R. A. (2004). Global position and
recruitment of HATs and HDACs in the yeast genome. Mol Cell 16, 199-209.
Rundlett, S. E., Carmen, A. A., Suka, N., Turner, B. M. and Grunstein, M. (1998).
Transcriptional repression by UME6 involves deacetylation of lysine 5 of
histone H4 by RPD3. Nature 392, 831-5.
Rupp, W. D. and Howard-Flanders, P. (1968). Discontinuities in the DNA
synthesized in an excision-defective strain of Escherichia coli following
ultraviolet irradiation. J Mol Biol 31, 291-304.
Sanchez-Diaz, A., Kanemaki, M., Marchesi, V. and Labib, K. (2004). Rapid
depletion of budding yeast proteins by fusion to a heat-inducible degron. Sci
STKE 2004, PL8.
Sar, F., Lindsey-Boltz, L. A., Subramanian, D., Croteau, D. L., Hutsell, S. Q.,
Griffith, J. D. and Sancar, A. (2004). Human claspin is a ring-shaped DNA-
binding protein with high affinity to branched DNA structures. J Biol Chem
279, 39289-95.
79
Sclafani, R. A. and Fangman, W. L. (1986). Thymidine utilization by tut mutants and
facile cloning of mutant alleles by plasmid conversion in S. cerevisiae.
Genetics 114, 753-67.
Sikorski, R. S. and Hieter, P. (1989). A system of shuttle vectors and yeast host
strains designed for efficient manipulation of DNA in Saccharomyces
cerevisiae. Genetics 122, 19-27.
Sivakumar, S., Porter-Goff, M., Patel, P. K., Benoit, K. and Rhind, N. (2004). In
vivo labeling of fission yeast DNA with thymidine and thymidine analogs.
Methods 33, 213-9.
Stevenson, J. B. and Gottschling, D. E. (1999). Telomeric chromatin modulates
replication timing near chromosome ends. Genes Dev 13, 146-51.
Szyjka, S. J., Viggiani, C. J. and Aparicio, O. M. (2005). Mrc1 is required for normal
progression of replication forks throughout chromatin in S. cerevisiae. Mol
Cell 19, 691-7.
Tanaka, K. and Russell, P. (2001). Mrc1 channels the DNA replication arrest signal
to checkpoint kinase Cds1. Nat Cell Biol 3, 966-72.
Tercero, J. A. and Diffley, J. F. (2001). Regulation of DNA replication fork
progression through damaged DNA by the Mec1/Rad53 checkpoint. Nature
412, 553-7.
Tourriere, H., Versini, G., Cordon-Preciado, V., Alabert, C. and Pasero, P. (2005).
Mrc1 and Tof1 promote replication fork progression and recovery
independently of Rad53. Mol Cell 19, 699-706.
Vernis, L., Piskur, J. and Diffley, J. F. (2003). Reconstitution of an efficient
thymidine salvage pathway in Saccharomyces cerevisiae. Nucleic Acids Res
31, e120.
80
Vickers, M. F., Mani, R. S., Sundaram, M., Hogue, D. L., Young, J. D., Baldwin, S.
A. and Cass, C. E. (1999). Functional production and reconstitution of the
human equilibrative nucleoside transporter (hENT1) in Saccharomyces
cerevisiae. Interaction of inhibitors of nucleoside transport with recombinant
hENT1 and a glycosylation-defective derivative (hENT1/N48Q). Biochem J
339 ( Pt 1), 21-32.
Viggiani, C. J. and Aparicio, O. M. (2006). New vectors for simplified construction
of BrdU-Incorporating strains of Saccharomyces cerevisiae. Yeast 23, 1045-
51.
Vogelauer, M., Rubbi, L., Lucas, I., Brewer, B. J. and Grunstein, M. (2002). Histone
acetylation regulates the time of replication origin firing. Mol Cell 10, 1223-
33.
Vogelauer, M., Wu, J., Suka, N. and Grunstein, M. (2000). Global histone
acetylation and deacetylation in yeast. Nature 408, 495-8.
Weinreich, M., Palacios DeBeer, M. A. and Fox, C. A. (2004). The activities of
eukaryotic replication origins in chromatin. Biochim Biophys Acta 1677, 142-
57.
Wyrick, J. J., Aparicio, J. G., Chen, T., Barnett, J. D., Jennings, E. G., Young, R. A.,
Bell, S. P. and Aparicio, O. M. (2001). Genome-wide distribution of ORC
and MCM proteins in S. cerevisiae: high-resolution mapping of replication
origins. Science 294, 2357-60.
Xu, H., Boone, C. and Klein, H. L. (2004). Mrc1 is required for sister chromatid
cohesion to aid in recombination repair of spontaneous damage. Mol Cell
Biol 24, 7082-90.
Xu, W. (2006). Searching For and Beyond Origins, Molecular Biology, University of
Southern California, pp. 212.
Xu, W., Aparicio, J. G., Aparicio, O. M. and Tavare, S. (2006). Genome-wide
mapping of ORC and Mcm2p binding sites on tiling arrays and identification
of essential ARS consensus sequences in S. cerevisiae. BMC Genomics 7,
276.
81
Zappulla, D. C., Sternglanz, R. and Leatherwood, J. (2002). Control of replication
timing by a transcriptional silencer. Curr Biol 12, 869-75.
Zhao, H. and Russell, P. (2004). DNA binding domain in the replication checkpoint
protein Mrc1 of Schizosaccharomyces pombe. J Biol Chem 279, 53023-7.
82
Appendix A: BrdU pools are depleted over time
Figure 18. BrdU incorporation at late replicating sequences is less efficient than at early
replicating sequences. CVy43 (BrdU-Inc::URA3) cells were arrested in G1 and released into S
phase in media containing 400 µg/ml BrdU. BrdU-labeled DNA was immunoprecipitated after 120
minutes. At this time point S phase was complete as measured by FACS analysis (data not shown)
and all sequences had replicated in the presence of BrdU. BrdU incorporation along chromosome VI
(in kb) was measured as (A) un-normalized raw data or (B) normalized using Drosophila clones
(DC). Input DNA (that had not been immunoprecipitated) from the 120-minute time point was co-
hybridized. This analysis demonstrates that early replicating sequences incorporate BrdU more
efficiently than late replicating ones, suggesting that BrdU pools are depleted significantly during a
typical time course.
Raw data
DC Normalization
A.
B.
607 608 609 606 605 604 603.5 603 602.5 601/2
607 608 609 606 605 604 603.5 603 602.5 601/2
Abstract (if available)
Abstract
The faithful duplication of the genome during S phase is critical to a eukaryotic cell's healthy proliferation. Since errors in DNA replication can lead to disastrous cellular consequences, this process has evolved to be highly regulated throughout species. In Saccharomyces cerevisiae, DNA replication initiates from multiple discrete chromosomal loci, termed origins of replication. DNA synthesis proceeds bidirectionally away from origins to replicate the entire genome prior to mitosis and cell division. Although many of the events regulating entry into S phase have been characterized, the regulation of events occurring within S phase are more obscure. For instance, replication origins initiate according to a temporal program that often correlates with transcriptional activity and appears to be regulated, in part, by their surrounding chromatin structure. To examine replication timing and replication fork elongation on a genome-wide scale I have developed a series of integrating plasmid vectors that enable the one-step construction of yeast strains that can incorporate the thymidine analogue BrdU into replicating DNA. By immunoprecipitating BrdU-labeled DNA I determine that the histone deacetylase complex Sin3-Rpd3 functions prior to S phase to properly delay the initiation timing of a subset of late origins, likely by creating a repressive chromatin structure around origins. I also use BrdU incorporation to help demonstrate that Mrc1, in addition to its checkpoint function, plays a central and constitutive role at normal replication forks. Finally, I provide a novel BrdU-pulse approach, coupled with microarrays, to examine how replication forks respond to, and recover from, replication stress and DNA damage.
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Asset Metadata
Creator
Viggiani, Christopher John
(author)
Core Title
New tools for whole-genome analysis of DNA replication timing and fork elongation in saccharomyces cerevisiae
School
College of Letters, Arts and Sciences
Degree
Doctor of Philosophy
Degree Program
Molecular Biology
Degree Conferral Date
2007-08
Publication Date
07/05/2007
Defense Date
06/08/2007
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
BrdU,chromatin structure,DNA replication,OAI-PMH Harvest,replication forks,replication origin initiation,Sin3-Rpd3
Language
English
Advisor
Aparicio, Oscar (
committee chair
), Finkel, Steven E. (
committee member
), Goodman, Steven (
committee member
), Rice, Judd C. (
committee member
), Tower, John G. (
committee member
)
Creator Email
cviggiani@gmail.com
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https://doi.org/10.25549/usctheses-m585
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Viggiani, Christopher John
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texts
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(contributing entity),
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Repository Email
cisadmin@lib.usc.edu
Tags
BrdU
chromatin structure
DNA replication
replication forks
replication origin initiation
Sin3-Rpd3