Chromatin Reassembly following a DNA Double-Strand Break Repair: The Ctf18-complex and Ctf4 work
in concert with H3K56 Acetylation
by
Harshika Seepany
A thesis submitted in conformity with the requirements for the degree of Masters
Department of Molecular Genetics University of Toronto
© Copyright by Harshika Seepany 2011
ii
Chromatin Reassembly following a DNA Double-Strand Break
Repair: The Ctf18-complex and Ctf4 work in concert with
H3K56 Acetylation
Harshika Seepany
Masters
Department of Molecular Genetics
University of Toronto
2011
Abstract
The budding yeast, Saccharomyces cerevisiae, serves as an excellent model for
identifying fundamental mechanisms of DNA repair. A Local Coherence Detection (LCD)
algorithm that uses biclustering to assign genes to multiple functional sub-groups was applied
on the chromosome E-MAP containing genetic interactions among genes involved in nuclear
processes. Using this method, we found that Asf1 and Rtt109, genes that are together
required for histone H3K56 acetylation, cluster together with Ctf4, Ctf18, Ctf8 and Dcc1,
genes important for efficient sister chromatid cohesion. It is known that H3K56 acetylation is
required for post-repair chromatin reassembly at sites of DNA double-strand breaks (DSBs).
The cohesion genes were previously implicated in the repair of some DNA DSBs, but the
nature of their involvement has not been reported. The experimental data in my thesis work
suggest that Ctf4, Ctf8, Ctf18 and Dcc1 function in the post-repair chromatin reassembly
pathway.
iii
Acknowledgments
I am indebted to my supervisors, Dr. Jack Greenblatt and Dr. Shoshana Wodak for
their guidance in my project. Their brilliance and foresight is what guided me throughout my
graduate studies. I am thankful to our many scholarly discussions through the past few years.
They both have inspired me in many ways and helped me gain invaluable skills. These are
skills that are important not just in science research, but in other career paths as well. I have
no doubt that my graduate school experience in the Greenblatt and Wodak Lab has prepared
me well for the next phase in my life.
I would also like to thank everyone in my supervisory committee and exam
committee, especially my supervisory committee members Dr. Gary Bader (past member),
Dr. Brigitte Lavoie and Dr. Daniel Durocher for their time and patience. They have given me
incredible feedback that has helped me become an independent thinker and problem solver.
I want to thank all the member of the Greenblatt Lab, with whom I have spent almost
every day of my last three years. I have learnt so much from every one of them. I am
especially grateful to Dr. Jeffrey Fillingham, who have trained me and taught me much of the
experimental techniques. His perspective and brilliance is incredible, and I am happy that I
got the opportunity to work so closely with him.
Finally, I would like to thank my family and friends for their moral support in every
academic decision I have made and during my time at the graduate school. I am especially
thankful to Harsh Jain, Gaurav Jain and my nephew Maanav Jalan for being a source of
constant encouragement.
iv
Table of Contents
Contents
Acknowledgments.................................................................................................................... iii
Table of Contents ..................................................................................................................... iv
List of Tables .......................................................................................................................... vii
List of Figures ........................................................................................................................ viii
List of Abbreviations .................................................................................................................x
Chapter 1 Introduction ...............................................................................................................1
1 Introduction ..................................................................................................................2
1.1 Yeast as a model organism to study human gene function ..........................3
1.2 An Epistatic MiniArray Profile (E-MAP) of chromosome-related genes ...................4
1.3 Local Coherence Detection Algorithm ........................................................................7
1.4 Repair of DNA double-strand breaks ..........................................................................9
1.5 Signaling the presence of DNA DSBs .......................................................................14
1.6 Histone H3K56 acetylation and the histone code at the site of a DNA DSB ............15
1.7 Sister chromatid cohesion and DNA repair ...............................................................18
1.8 Galactose induction of a double-strand break ...........................................................21
1.9 Conservation of DSB repair pathways in higher organisms ....................................23
1.10 Thesis Rationale .........................................................................................................24
Chapter 2 Materials and Methods ..............................................................................................2
2 Material and Methods ................................................................................................28
2.1 Yeast transformations ................................................................................................28
2.2 Yeast strains and strain construction .........................................................................29
2.2.1 Construction of single and double mutants ......................................................33
v
2.2.2 Construction of tagged strains .........................................................................36
2.2.3 Confirmation of the mutant/tagged strains ......................................................38
2.3 Growth sensitivity assays ..........................................................................................38
2.4 Analysis of DNA DSB repair by SSA .......................................................................39
2.5 Whole-cell extraction and western blotting ...............................................................41
2.6 Chromatin fractionation .............................................................................................43
2.7 Chromatin immunoprecipitation (ChIP) ....................................................................44
2.8 One step TAP-Tag purification .................................................................................50
Chapter 3 Results .....................................................................................................................28
3 Results........................................................................................................................53
3.1 Epistasis relationships between genes required for efficient cohesion and those
required for histone H3K56 acetylation ..............................................................................53
3.2 Effect of MMS on the association of Ctf4p and Ctf18p with the chromatin fraction
of the cell .............................................................................................................................58
3.3 Sensitivity of cells lacking cohesion promoting genes to the presence of a single
double-strand break .............................................................................................................62
3.4 The Ctf18p complex and Ctf4p are not required for DSB repair by SSA .................63
3.5 Rad53 hyperphosphorylation in the absence of cohesion promoting genes ..............68
3.6 Chromatin reassembly at the site of the DNA double-strand break ..........................70
3.7 Kinetics of the appearance of Ctf18p and Ctf4p around the site of the DSB ............73
3.8 Influence of histone H3K56 acetylation on the recruitment of Ctf18p and Ctf4p
around the site of a DSB ......................................................................................................78
3.9 Physical interactions of Asf1p with Ctf18p and Ctf4p ..............................................82
Chapter 4 Discussion and Future Experiments ........................................................................53
4 Summary ....................................................................................................................86
4.1 Cellular response to DNA damage ............................................................................90
4.2 Chromatin reassembly around the site of a DNA DSB .............................................95
vi
4.3 Future experiments ....................................................................................................98
4.3.1 What are the defects in DNA DSB repair in the absence of CTF18 and
CTF4: DNA replication or chromatin reassembly? .........................................98
4.3.2 Where does Ctf18p function in this pathway? .................................................99
4.3.3 Why does the presence of Ctf4p depend on Asf1p/Rtt109p? ........................100
4.3.4 Is there a role for other histone chaperones in DSB repair? ..........................101
Chapter 5 References .............................................................................................................102
Copyright Acknowledgements...............................................................................................116
vii
List of Tables
Table 1 – List of strains used in this study 29
Table 2 – List of PCR primers used for deletion and conformation 34
Table 3 – List of PCR primers used for tagging 36
Table 4 – List of PCR primers around the DBS used for analysis of DNA repair by SSA
(from Keogh et. al., 2006) 40
Table 5 – List of PCR primers spanning 20 kb around the site of DSB used for ChIP 47
viii
List of Figures
Figure 1 – Results obtained after applying the LCD algorithm to the chromosome function
E-MAP dataset 8
Figure 2 – A bicluster containing genes involved in DNA repair together with cohesion
promoting genes obtained using the LCD Algorithm 10
Figure 3 – Three different modes of DNA DSB repair by homologous recombination 12
Figure 4 – Galactose-inducible HO system to study the repair of a DSB by HR mediated by
SSA in the YMV2 strain 22
Figure 5 – Epistasis analysis for various mutations in the presence of 0.1% MMS 55
Figure 6 – Epistasis analysis for various mutations in the presence of 0.1% MMS 56
Figure 7 – Effects of deleting cohesion promoting genes on histone modifications 57
Figure 8 – Effect of MMS on the association of Ctf18p with the chromatin fraction of the
cell 60
Figure 9 – Effect of MMS on the association of Ctf4p with the chromatin fraction of the
cell 61
Figure 10 – Effects of deletions of cohesion promoting genes on the growth of a strain with a
single inducible DSB 64
Figure 11 – Effects of deletions of cohesion promoting genes on the growth of a strain with a
single inducible DSB 65
Figure 12 – PCR analysis to assess the effects of mutations in cohesion promoting genes on
DSB formation and its repair 67
Figure 13 – Western blotting to assess the effect of various mutations in cohesion promoting
genes on Rad53p hyperphosphorylation as an index of checkpoint activation and relief 69
ix
Figure 14 – Effects of various mutations in cohesion promoting genes on nucleosome
occupancy at the DSB 71
Figure 15 – Relative enrichment of histone H3 at the site of a DNA DSB 72
Figure 16 – Occupancy of Ctf18p at various positions around the site of the DSB at various
time points 75
Figure 17 – Occupancy of Ctf4p at various positions around the site of the DSB at various
time points 76
Figure 18 – Relative enrichment of Ctf18p (at 2 hours) and Ctf4p (at 6 hours) for about 20kb
on either side of the DSB 77
Figure 19 – The enrichment of Ctf18p around the site of the break in the absence of ASF1 or
RTT109 2 hours after DSB induction 79
Figure 20 – The enrichment of Ctf4p around the site of the break in the absence of ASF1 or
RTT109 6 hours after DSB induction 80
Figure 21 – Influence of CTF18 on the enrichment of Ctf4p around the site of the break 6 hours
after DSB induction 81
Figure 22 – Physical interactions of Asf1p with Ctf18p and Ctf4p 83
Figure 23 – Repair of a DNA DSB by single-strand annealing 89
Figure 24 – Final Model for the role of Ctf18-complex and Ctf4 during the process of DNA
DSB 91
x
List of Abbreviations
ASF – Anti-silencing factor
BIR – Break induced repair
BSA – Bovine serum albumin
CAR – Cohesin associated region
ChIP – Chromatin immunoprecipitation
CPT – Campothecin
CTF – Chromosome transmission fidelity
DDR – DNA damage response
DMSO – Dimethyl sulfoxide
DNA – Deoxyribonucleic acid
DSB – Double-strand break
DTT – Dithiothreitol
EDTA – Ethylenediaminetetraacetic acid
E-MAP – Epistatic miniarray profile
EtBr – Ethidium bromide
GC – Gene conversion
HAT – Histone acetyltransferase
HCl – Hydrochloric acid
HDAC – Histone deacetylase
HEPES – 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid
HO – Homothallic
HR – Homologous recombination
xi
HU – Hydroxyurea
IP – Immunoprecipitation
KAN – Kanamycin
KAT – Lysine acetyltransferase
kb – Kilo bases
KCl – Potassium chloride
KOH – Potassium hydroxide
LCD – Local coherence detection
LiCl – Lithium chloride
MgCl2 – Magnesium chloride
MMS – Methyl methane sulfonate
NaAz – Sodium Azide
NaCl – Sodium chloride
NAT – Nourseothricin
NER – Nucleotide excision repair
NHEJ – Non-homolohous end joining
OD – Optical density
PAGE – Polyacrylamide gel electrophoresis
PCNA – Proliferating cell nuclear antigen
PCR – Polymerase chain reaction
PEG – Polyethylene glycol
PTM – Post-translational modification
RAD – Radiation sensitive
xii
RFC – Replication factor C
rpm – Rotations per minute
RTT – Regulator of ty1 transposition
SCC – Sister chromatid cohesion
SDS – Sodium dodecyl sulfate
SGA – Synthetic genetic array
SSA – Single-strand annealing
ssDNA – Single-stranded DNA
TAP – Tandem affinity purification
TCA – Trichloroacetic acid
WCE – Whole cell extract
WT – Wild type
YPD – Yeast peptone dextrose
1
Chapter 1
Introduction
2
1 Introduction
Chromatin consists of nucleosomes, short stretches of DNA wrapped around histone
octamers composed of two dimers of core histones H2A and H2B (H2A-H2B) connected to a
tetramer of core histones H3 and H4 (H3-H4)2 [1, 2]. The nucleosome represents the first
level of DNA compaction, and this structure stabilized by the linker histone H1 subsequently
folds around itself in several higher levels of organization, thereby enabling the DNA to be
packaged within the confined space of the nucleus. Formation of nucleosomes occurs
reversibly during the life cycle of the cell. Nucleosome assembly and disassembly are
influenced by histone modifications, by energy-dependent nucleosome repositioning, or
replacement of core histones by various variants, all of which can lead to the active
recruitment of other regulatory proteins [3]. With increased understanding of these processes,
it is clear that the dynamic mechanisms of nucleosome assembly and disassembly are
functionally important and control many important events in the nucleus, including
transcription, DNA replication, chromosome condensation, telomeric silencing and DNA
repair [4-6].
Motivation for the research described in this thesis was provided by analysis of yeast
genetic interaction data suggesting that several genes involved in sister chromatid cohesion
could function together with histone H3K56 acetylation in the repair of DSBs in DNA.
During the past several years there have been numerous reports linking chromatin
remodeling to the execution of specific steps in DSB repair pathways [7]. Chromatin
remodeling leading to specific alterations of chromatin around a DSB is required for the
subsequent recruitment or stabilization of repair factors at DSBs. Various kinds of
3
misregulations in chromatin remodeling can potentially lead to mutation and transform a
normal epigenetic landscape into a cancerous one. For example, failure to repair DSBs, or
misrepair, can result in cell death or large-scale chromosomal changes that enhance
chromosome instability. Chromatin remodeling factors are also becoming major drug targets
for an increasing number of diseases, and the importance of understanding the genes
involved in this complex process is clear [8, 9].
1.1 Yeast as a model organism to study human gene
function
The yeast Saccharomyces cerevisiae serves as an excellent model system to
understand the molecular mechanisms of basic processes in eukaryotic cells [10]. The
Saccharomyces cerevisiae genome encodes about 6000 genes, many of which have
counterparts in higher organisms. Its ease of genetic manipulation and propagation, the
availability of genomic resources and the conservation of basic cellular processes in
eukaryotes makes yeast an attractive model system. The proteins encoded by the human
homologs of certain yeast genes are so similar that replacing the yeast gene by its human
counterpart can often restore the function of the gene [11]. For example, two human genes
required for mismatch repair, MSH2 and MSH1, and the human DNA helicase SGS1, which
have been shown to be linked to colorectal cancer and Werner’s syndrome (a disease related
to premature aging), respectively, can be studied in yeast [12-15]. This also makes yeast an
attractive model for studying the function of disease-related genes. In fact, much of our
4
knowledge about cell cycle regulators, cell cycle checkpoints, and DNA repair came from
studies performed in yeast [16].
1.2 An Epistatic MiniArray Profile (E-MAP) of
chromosome-related genes
A genetic interaction is said to occur when a mutation in one gene gives an
unexpected outcome when combined with a mutation in another gene. When growth is
measured as a phenotype, an aggravating (synthetic sick/lethal) genetic interaction is a type
of genetic interaction in which the double mutant grows more slowly than expected given the
growth rates of the individual single mutants, whereas an alleviating (or suppressing)
interaction is one in which the double mutant is not sicker, or indeed grows better, than
expected, given the growth rates of the single mutants [17, 18]. A genetic interaction between
two genes is said to be epistatic when both the genes work in a common pathway, such that a
deletion of a second gene in addition to the first one does not cause any additional growth
sensitivity. The extent of a genetic interaction can be assessed by calculating the size of the
mutant colony, as a proxy for growth rate. The most commonly used model for calculating
the extent of a yeast genetic interaction is the neutral model, which assumes that genetic
interactions are rare and favors a multiplicative scoring system in which a null (or neutral)
interaction is one for which the growth rate of the double mutant is equal to the product of
the growth rates of the two single mutants [19-21].
Genetic interaction studies have been very helpful in understanding the relationships
between yeast genes and the pathways in which they work, since most of the yeast genes are
5
non-essential, and their deletions are well tolerated by the cell [20]. The study of yeast
genetic interactions was revolutionized by the development of SGA (synthetic genetic array)
technology, in which a query mutation in a haploid strain of one mating type is crossed with
deletions of all the non-essential genes in a haploid yeast deletion collection of the opposite
mating type (about 4700 yeast deletion strains in the original work). Following sporulation of
the resulting heterozygous diploids and selection for double mutant haploids, the growth of
the double mutants is compared to the growth of single mutants. This enabled the high
fidelity systematic assessment of synthetic sick/lethal interactions.
Many large and small scale studies have used SGA technology for the identification
of genes involved in cell polarity, DNA synthesis, DNA repair, transcription and secretion
[19, 22-29]. Most recently, considerable effort has been made to study all yeast genes in this
way [29]. These studies have resulted in the cataloguing of millions of individual genetic
interactions, but the confounding task lies in the interpretation of the data [30]. Many studies
have put forward different ideas and methods to extract meaningful information from these
genetic interaction data. Similar to these large scale genetic interaction studies, there are also
many large scale studies on physical interactions between proteins [31-34]. Some studies
have also integrated genetic and physical interaction data for a better and more complete
understanding of the relationships between proteins and the pathways in which they work
[35, 36].
To facilitate better understanding of protein function, a high-throughput variation of
SGA, the E-MAP (Epistatic Miniarray Profile), was created which involved the elucidation
of genetic interactions among a subset of genes working in a single cellular process (e.g. the
early sceretory pathway or chromosome function) [21, 22, 37]. Confining the experimental
6
work to a single cellular process is beneficial, as including unrelated genes increases the size
and cost of the dataset exponentially without adding much meaningful data (for every n
genes that are studied, the size of the dataset is n2) [19, 22]. Working with a smaller dataset
also improves the signal-to-noise ratio, so that the genetic interactions in an E-MAP are more
easily made quantitative.
The information-rich E-MAP on chromosome-related genes was particularly
interesting for my purposes, as it included genes involved in DNA synthesis, DNA repair,
chromosome segregation and transcription [22]. The E-MAP involved genetic interactions
among about 750 genes, mostly deletions of non-essential genes but also including some
hypomorphic alleles of essential genes. This E-MAP included assessment of both
aggravating and alleviating interactions based on colony sizes. The growth analysis was done
on a continuous scale, thus differentiating the extents of epistasis. Hierarchical clustering was
used to sort the functionally related gene groups, with genes having the most similar patterns
of genetic interactions being linked most closely. The resulting clusters were essentially
modular, with the basic cellular processes roughly separated. While physically interacting
protein complexes often corresponded to functional sub-modules, epistasis groups of genes
otherwise not involved in physical interactions were also obtained. Thus, the E-MAP
emerged as a powerful tool to discover genes working together in distinct, compensatory or
common pathways [22]. Similar SGA and E-MAP-like techniques have also been used for
other unicellular organisms (e.g. S.pombe, E.coli) and for drug screening using chemical
genetics approaches [38-41].
7
1.3 Local Coherence Detection Algorithm
E-MAP datasets contain a plethora of information, much of which is dependent on
computational analysis of the data [42]. While hierarchical clustering of E-MAP data on
chromatin-related genes successfully grouped genes into functional modules, it does not
represent the functional versatility of a protein. With growing understanding of protein
function, it is evident that certain proteins can interact with different sets of proteins to
accomplish different functions, and many protein complexes share subunits. For example,
two histone acetyltransferases, Gcn5p and Rtt109p, acetylate histone H3 on Lysine 9, but
they also acetylate other histone lysine residues independently as well [43, 44]. Similarly, the
histone chaperone Asf1p is important for DNA synthesis, transcriptional silencing and DNA
repair, and it interacts with different proteins to accomplish its different functions [45-48].
Thus, if there were a way to group genes into overlapping epistasis groups, it would help in
discovering multiple functions of many chromatin remodeling genes.
The Local Coherence Detection (LCD) algorithm is a biclustering algorithm that can
group a gene into multiple clusters, in which each cluster contains functionally related genes
[35]. In effect, the LCD finds subsets of library genes for which certain query genes share
very similar genetic interaction patterns. Each cluster is also assigned a score that reflects
confidence in the existence of that cluster. This method was validated and applied to other
similar datasets as well, e.g. the E-MAP on the early secretory pathway and a microarray
study.
Figure 1 represents the functional connections predicted by the LCD for complexes
and pathways involved in chromatin-related functions. Many as yet unknown and
8
Figure 1. Results obtained after applying the LCD algorithm to the chromosome function E-
MAP. The green nodes represent protein complexes/epistasis groups and the pink lines
connecting the nodes represent predicted functional connections. The thickness of the edges
is proportional to the confidence in the functional connection between the two connected
nodes. Figure taken from Pu, S., Ronen, K., Vlasblom, J., Greenblatt, J., and Wodak, S. J.
(2008). Local coherence in genetic interaction patterns reveals prevalent functional
versatility. Bioinformatics 24, 2376-2383.
a. Known functional connected between Set2-complex and Rpd3S complex b. Predicted
functional connection between Swr1-complex and Rpd3L complex c. Predicted functional
connection between DNA damage epistasis group, Ctf18-complex and checkpoint genes.
9
uncharacterized functional connections were predicted (represented by the pink lines in
Figure 1) in addition to the ones that had already emerged from simple hierarchical
clustering. Thus, LCD emerged as a powerful tool to predict functional connections among
genes and assign new functions to genes. One of the new functional connections predicted by
LCD involved the proteins in the histone H3K56 acetylation pathway, i.e. Asf1p and
Rtt109p, together with several non-essential proteins required for efficient sister chromatid
cohesion, i.e. Ctf18p, Ctf8p, Dcc1p and Ctf4p, as shown in Figure 2.
1.4 Repair of DNA double-strand breaks
Cells are constantly assaulted by both endogenous factors and exogenous agents that
can cause lesions in their DNA. If left unrepaired, these lesions can lead to abnormal cell
growth and sometimes even cell death. Hence, damaged DNA must be repaired. In the event
of DNA damage, eukaryotic cells activate DNA damage checkpoints to ensure that the
damaged DNA is repaired before the cell progresses into another cell cycle [49]. Activation
of the checkpoint not only gains time for the cell to react to the DNA damage by arresting
cell cycle progression but also turns on a signaling cascade leading to the recruitment of
repair proteins to the sites of DNA damage. Such checkpoints can act at three stages of the
cell cycle, at the G1/S boundary, during progression through S phase and at the G2/M
transition.
The problems faced by cells following DNA damage depend on the stage of the cell
cycle. During the G1 resting state, the cells mostly accumulate oxidative damage to DNA
that must be repaired prior to entering into S phase, whereas during S phase the cells must
10
A: Histone H3K56 Acetylation Pathway B: MMS complex
C: Cohesion promoting genes D: MRX complex
Figure 2. A bicluster containing genes involved in DNA repair together with cohesion
promoting genes obtained using the LCD Algorithm. The scale indicates the color scheme
used for the scoring system, where green represents aggravating interactions and red
represents alleviating interactions. Blank/white boxes indicate that the genetic interaction
information is not available.
Scale
A B C D
11
ensure complete DNA replication and correct nucleotide incorporation, and during mitosis
the cells must ensure proper segregation of the sister chromatids. Although there are
checkpoint proteins common to the various stages of the cell cycle, the G1/S checkpoint is
mostly sensitive to the concentration of the nucleotide pool and single-stranded gaps
remaining after excision repair, whereas the G2/M arrest is very sensitive to DNA double-
strand breaks (DSBs) [50]. It is also thought that certain types of repair can only take place at
a particular stage of the cell cycle, and thus the checkpoint becomes very important to
monitor the repair before the cell cycle progresses [50]. Furthermore, depending on the
nature of the damage, the cell activates different repair mechanisms [51].
DNA DSBs are the most lethal form of DNA damage and, if not repaired properly,
can cause genomic instability and lead to gross chromosomal loss. There are two major
pathways for repairing DNA DSBs, Homologous Recombination (HR) and Non-
Homologous End Joining (NEHJ), sometimes called error-free and error-prone repair,
respectively. Whereas HR requires homology to the broken ends of the DNA and usually
uses the sister chromatid as a template for repair, NHEJ simply involves religation of the
broken ends [52, 53].
Broadly speaking, there are 3 different types of DNA repair by HR: break-induced
repair [54], gene conversion (GC) and single-strand annealing (SSA) (Figure 3). When both
ends of the DSB share homology with the sister chromatid, a homologous chromosome or an
ectopically located donor, the break is usually repaired by gene conversion (GC); when there
is homology to only one end of the DSB, then the repair occurs by break-induced repair [54];
and when the break is flanked by direct repeats, the repair occurs by single-strand annealing
(SSA). Repair by HR always involves extensive 5’ to 3’ resection of the broken DNA ends,
12
Figure 3. Three different modes of DSB repair by homologous recombination: break-
induced repair (BIR), gene conversion (GC) and single-strand annealing (SSA).
Gene Conversion Single Strand
Annealing
Break-Induced
Repair
13
leaving single-stranded DNA 3’ extensions. Whereas the initial step of homology search and
strand invasion by the 3’ extensions occurs with almost equal efficiency and kinetics in all
three processes, the initiation of new DNA synthesis is regulated differentially among these
processes [51].
During SSA, the broken ends are resected till the two homologous ends become
single-stranded and anneal, leading to the loss of the intervening DNA and deletion of one of
the repeats [55, 56]. Since the process of SSA is dependent on resection for exposing the
homologous sequences, the distance between the homologous regions governs the kinetics of
repair, whereas the efficiency of the repair process itself is dependent on the length of the
repeats, the extent of sequence identity between the repeats and the presence of a third repeat
[56-59].
Once the break is repaired, the DNA damage checkpoint is turned off to allow normal
cell cycle progression, leading to checkpoint recovery. However, when the cells are unable to
repair the damaged DNA, there is yet another process by which the cells can turn off the
checkpoint and resume the cell cycle. This process is termed adaptation and appears to be a
highly regulated process. Many key proteins involved in homologous recombination and the
processing of DSBs have been linked to adaptation, including Cdc5p, Ckb1p, Ckb2p, Tid1p,
Yku70p, Yku80p and Srs2p [58, 60-63]. Sometimes, cells with a single HO-induced DSB
can adapt, whereas those with two DSBs cannot [60]. However, adaptation is not dependent
on the number of DSBs, but rather on the extent of single-stranded DNA (ssDNA) that is
created as a result of the DSB [60].
14
1.5 Signaling the presence of DNA DSBs
In Saccharomyces cerevisiae DNA DSBs cause cells to arrest in the G2/M phase of
the cell cycle. Moreover, a single DSB is sufficient to activate the DNA damage checkpoint
and arrest the cell cycle until the repair is complete [64]. The Mec1p and Tel1p protein
kinases, whose human counterparts are ATR and ATM, respectively, are initially activated
along with the Mec1p partner Dcc2p [64]. The interaction between Mec1p and Dcc2p is
required for the recruitment of Mec1p to the site of DNA damage, where Mec1p and Dcc2p
recognize the ssDNA generated by resection of DNA DSBs. While both Mec1p and Tel1p
are required to phosphorylate histone H2A on serine 129 (Serine 139 in human H2A.X),
Mec1p principally phosphorylates the checkpoint signal transducer kinases, Chk1p and
Rad53p (the yeast orthologue of human CHK2), as well a checkpoint mediator protein Rad9p
[64-67]. Mec1p is itself activated by the Rad9p kinase. Once Rad53p protein is
phosphorylated by Mec1p, it autophosphorylates in the presence of Rad9p [64].
Hyperphosphorylated Rad53p then modulates the downstream effectors of DNA repair and
cell cycle progression [68].
Upstream of Mec1p are two independent monitors of DNA damage, one comprised of
Rad24p with the trio of Rad17p/Mec3p/Ddc1p, the other being Rad9p [69-73]. Thus, there is
a tight regulation of the kinases and their activities to prevent spurious activation/deactivation
of the DNA damage signal. Permanent cell cycle arrest depends on active maintenance of the
checkpoint kinase cascade through the phosphorylation of at least two proteins, Rad53p and
Chk1p. Intriguingly, this phenomenon of checkpoint arrest is only specific to G2/M. When
15
cells are arrested in G1 and DNA damage is induced, there is no activation of the Rad53p
kinase [60].
Once the break is repaired, the Rad53p phosphorylation is reversed by its
dephosphorylation [74]. The dephosphorylation of Rad53p is achieved by two independent
phosphatase complexes: one is the PP4-type phosphatase complex, Pph3p-Psy2p,
independent of its partner, Psy4p; and the other is the PP2C-type phosphatase, Ptc2-Ptc3 [75-
77]. Additionally, Pph3p is required for the dephosphorylation of γH2AX [78]. These
dephosphorylation events finally lead to switching off of the cell cycle checkpoint and
resumption of cell cycle progression.
One commonality of all DSBs is resection for the generation of ssDNA. In budding
yeast, a trio of three proteins, Mre11p-Rad50p-Xrs2p (MRX), assisted by Sae2p and Exo1p,
is required for performing resection at DSBs [79-81].
Many proteins have well characterized roles in the events that follow a DNA DSB.
While the checkpoint activation and the subsequent recruitment of proteins to the DSB are
dependent on the type of repair, there are some proteins that have common roles in all
mechanisms of DSB repair (e.g. H2AX, Rad53p and Chk1p phosphorylation). Furthermore,
presumably for tighter regulation, there are also redundant pathways and feedback loops.
1.6 Histone H3K56 acetylation and the histone code at the
site of a DNA DSB
There are many types of histone post-translational modifications (PTMs) that alter
chromatin structure and function. Histone PTMs include lysine acetylation, lysine and
16
arginine methylation, serine and threonine phosphorylation, lysine ubiquitination, lysine
sumoylation and arginine and glutamate ADP-ribosylation [3]. Most of these modifications
occur on the long histone N-terminal tails and play fundamental roles in regulating
chromosome function by altering the accessibility of the DNA to various binding factors
and/or by creating binding sites for recruitment of specific protein complexes to their sites of
action [82]. Post-translational modification of the histone tails has been linked to different
chromatin states that regulate transcription, DNA replication, repair and recombination [83-
85]. Many of these histone modifications are also associated with DNA DSB repair [86-90].
The first histone modification that was found to be associated with DNA repair was
phosphorylation of Serine 129 on the C-terminal tail of histone H2A [78, 89].
Histone lysine acetylation involves the transfer of an acetyl group from acetyl-
coenzyme A to a lysine residue on a histone. This reaction is carried out by specialized
enzymes called histone acetyltransferases (formerly known as HATs and now known as
KATs). The transfer of the acetyl group to the lysine neutralizes its positive charge and
decreases the histone-DNA interaction, leading to a more relaxed conformation of the
chromatin and making the DNA more accessible. This also leads to the recruitment of
bromodomain-containing proteins which recognize the acetylated lysine residue. This relaxed
conformation can be reversed by histone deacetylation, i.e. removal of the acetyl group by a
histone deacetylase (HDAC) [91, 92]. Both histone acetylation and deacetylation are
important for the viability of cells following induction of a DSB [93]. In particular, the
histone H3 and H4 N-terminal tails and their acetylatable lysine residues are required for
growth following exposure to a HO endonuclease-induced DSB that is repaired by HR.
17
Histones can also be reversibly modified in their globular domains [83]. Unlike
deletion of the N-terminal tail of a histone, deletion of the core domain of a histone leads to
inviability [94, 95]. Lysine 56 acetylation in the core domain of histone H3 is required for
maximum transcription of several genes, including the histones themselves [96]. This
acetylation has also emerged as an important mark for histone deposition accompanying
DNA replication and DNA repair [84, 97]. As an indication of histone deposition behind the
replication fork, the level of H3K56 acetylation peaks during the S phase of the cell cycle.
This acetylation occurs predominantly on the newly synthesized histones [84]. Two proteins
were identified that were important for maintaining cellular levels of H3K56 acetylation, the
histone chaperone Asf1p and the HAT Rtt109p [44, 98]. Rtt109p together with Asf1p
acetylates newly synthesized histones on K56. H3K56 acetylation is reversed by members of
the Sir family of NAD+-dependent deacetylases. While this deacetylation is mainly carried
out by two Sir-family deacetylases, Hst3p and Hst4p, Sir2p deacetylates H3K56 in telomeric
regions [96, 99, 100].
Deletions of genes encoding proteins related to the H3K56 acetylation pathway,
including Asf1p, Rtt109p, Hst3p and Hst4p, as well as point mutations of K56 itself, cause
sensitivity to various DNA damaging agents, including HU, MMS and CPT, highlighting the
importance of this pathway during DNA replication and DNA damage/repair. There is tight
control over the levels of H3K56 acetylation, as both the acetyltransferase and the
deacetylase for H3K56 acetylation have strict cell cycle regulated patterns of expression:
where the Rtt109p level peaks during S phase [44], Hst3p and Hst4p levels drop during S
phase when H3K56 acetylation levels are at their peak [100]. As well, there is a Mec1p-
dependent proteolytic degradation of Hst3p/Hst4p in response to DNA damage, which
18
promotes subsequent high levels of H3K56 acetylation [101], possibly for active chromatin
reassembly after the repair of damaged DNA. Interestingly, both hyper-acetylated and under-
acetylated forms of H3K56 are deleterious for the cells in the presence of DNA damage.
In cells suffering a single HO endonuclease-induced DSB, which can be repaired by
HR following SSA, Asf1p- and Rtt109p-dependent histone acetylation of K56 is required for
the cells to exit the DNA damage checkpoint [97, 102]. While the DNA is repaired
efficiently in the absence of Asf1p and Rtt109p, the checkpoint protein Rad53p remains
hyperphosphorylated. When the lysine of H3K56 is replaced by glutamine, mimicking the
acetylated form, the need for Asf1p and Rtt109p for checkpoint relief is abrogated, indicating
that K56 acetylation is required for an event which leads to checkpoint relief. In fact, H3K56
acetylation is required for chromatin reassembly at the site of the DSB following the repair of
the DSB [97, 102].
1.7 Sister chromatid cohesion and DNA repair
During replication, the two copies of the replicated DNA have to be held together to
facilitate the proper segregation of the sister chromosomes during mitosis. This is achieved
by the protein complex cohesin. In yeast, cohesin is comprised of four proteins, Scc1p,
Scc3p, Smc1p and Smc3p, which form a ring-like structure that holds the two sister
chromatids together [103-108]. The cohesin complex associates with the DNA shortly before
S phase and dissociates at the metaphase to anaphase transition [107]. The binding of cohesin
to chromosomes is not sequence-specific but occurs preferentially at the centromeres and
along chromosome arm regions with high AT content, called cohesin-associated regions
19
(CARs) [109-112]. The loading of cohesin onto DNA is carried out by Scc2p and Scc4p
[113, 114]. Ctf7, another S-phase-specific essential protein required for cohesion, is a lysine
acetyltransferase that modifies Smc3p and Scc1p. Ctf7 is required for the establishment of
cohesion but it is dispensable for its maintenance [113, 115, 116]. In anaphase during
mitosis, the removal of the cohesin complex to liberate the two sister chromosomes is carried
out by separase (Esp1p). Separase is under the control of its inhibitor, securin (Psd5p), which
is degraded by the anaphase promoting complex (APC) to release Esp1p [117, 118].
In addition to cohesin’s role in holding the two sister chromatids together, it has also
been shown to be important for DNA repair and recombination, regulation of transcription, a
function at the mitotic spindle and specific roles in meiosis [119-123]. In particular, cohesin
plays an important role during the repair of DNA DSBs [124-127]. In the presence of a DSB,
cohesin is targeted to the regions flanking the break, leading to its de novo loading onto a
region of about 50Kb on either side of the break site in a sequence-independent manner.
Similar to cohesin loading prior to S-phase, cohesin at a DSB is loaded in a Scc2p- and
Scc4p-dependent manner [126]. However, DSB-induced cohesin loading depends on other
proteins as well, including the Mec1p and Tel1p kinases via the phosphorylation of histone
H2A S129, Rad53p and Mre11p [127]. Thus, the loading of cohesin is highly regulated
during DSB repair as well.
Ctf18p, Ctf8p and Ctf4p were initially identified as genes whose mutation leads to a
decrease in the fidelity of chromosome transmission [128]. These proteins are also important
for the proper establishment of sister chromatid cohesion, although they are not essential for
it. Cells lacking Ctf18p or Ctf4p have increased chromosome instability and show a strong
pre-anaphase delay, inducing the spindle assembly checkpoint [129-132]. Null mutants for
20
CTF18 and CTF4 show cohesion failure in 25-30% of cells that are held at metaphase in the
absence of microtubules [129].
Ctf18p shares sequence similarity with the large subunit of the RFC (Replication
factor C) complex, Rfc1. RFC is a five-subunit DNA-binding protein complex, which is
comprised of Rfc1p, Rfc2p, Rfc3p, Rfc4p and Rfc5p, recognizes the primer-template
junction and catalyzes the loading of the clamp, PCNA, during replication [133-135]. Ctf18p,
together with Ctf8p and Dcc1p, forms a complex with the small subunits of the RFC
complex, Rfc2-5 [129, 136, 137]. Two other RFC complexes have been identified in yeast,
Elg1p-RFC and Rad24-RFC [138-141]. Ctf18-RFC interacts with the PCNA and can both
load and unload PCNA from the DNA [137, 142, 143]. This function of Ctf18p suggests a
role in polymerase switching during DNA replication, possibly at sites where cohesin is
present, and thus leading to the efficient establishment of sister chromatid cohesion.
However, a direct link of Ctf18p, Ctf8p and Dcc1p to cohesin or establishment of cohesion
has not been uncovered. Moreover, Ctf18p, Ctf8p and Dcc1p are not required for either
association of cohesin with chromosomes or for protecting the Scc1 subunit of cohesin from
cleavage by separase during meiosis [144].
Ctf4p, on the other hand, was originally identified as a DNA polymerase interacting
protein [132, 145], but is now known for its interaction with multiple complexes and proteins
involved in DNA replication and repair, including the GINS and MCM complexes, Cdc45p,
Mrc1p, Tof1p, Csm3 and the histone chaperone FACT [146-148]. These interactions of
Ctf4p with components of the DNA replication fork make it an interesting candidate for
studying the interplay among different processes in the cell.
21
Most of the genes involved in the establishment of sister chromatid cohesion,
including cohesin, Ctf18p, Ctf8p, Dcc1p and Ctf4p, have been implicated in the process of
DNA DSB repair by HR [149, 150]. While the presence of cohesin at the site of DSB repair
is necessary for proper repair, the nature of the involvement of Ctf18p, Ctf8p, Dcc1p and
Ctf4p is still a matter of speculation.
1.8 Galactose induction of a double-strand break
S.cerevisiae has been used for many decades to study recombinational repair, and
many physical assays have been established in yeast to study the events that occur after a
DNA DSB. The yeast genome contains two site-specific endonucleases, HO and I-Sec1,
which cause DSBs at known loci. The homothallic (HO) endonuclease gene is part of a
tightly regulated system for the cells to switch from one mating type to another. Once a DSB
is created by HO at the MAT locus, DNA is copied into the MAT locus from either the HML
or HMR locus through HR by gene conversion [151]. Normally, the HO endonuclease is
expressed in cells that have previously divided and only at one time in the cell cycle [152].
By placing the HO endonuclease gene under the control of a galactose-inducible promoter,
the endonuclease can be expressed within minutes after treating cells with galactose at any
time during the cell cycle [153]. All aspects of DSB induction and DSB repair, including
resection, repair, checkpoint recovery and adaptation, have been studied using budding yeast
cells having either repairable or non-reparable DSBs created by the HO endonuclease [58,
60, 62].
22
Figure 4. Galactose-inducible HO system to study the repair of a DSB by HR mediated by
SSA in the YMV2 strain from Vase et. al. (2002). HO endonuclease is placed under the
control of a galactose-inducible promoter, which gets activated in the presence of galactose.
P1, P2, P3 and P4 represent primers used for assessing DSB induction and its subsequent
repair by PCR.
23
A recent study has shown the involvement of Asf1p and Rtt109p in chromatin
reassembly after a DNA DSB is repaired by SSA [97]. In this yeast cell background, the
MAT, HML and HMR loci are deleted and the HO endonuclease is placed under the control
of a galactose-inducible promoter. The recognition site of the HO endonuclease was inserted
at the leucine (leu2) gene locus, and a region homologous to the break site was placed about
30 kb (25 kb and 5 kb strains were also used in the original work) [58], as shown in Figure 4.
Once the endonuclease is induced, it creates a single DSB at the leu2 locus, and the DNA at
the site of the cut is resected till the two homologous sequences become single-stranded. For
a 30kb separation of the homologous regions it takes about 8 hours for the resection and
subsequent repair of the DSB by SSA, since the rate of resection is about 4 kb/hr [55, 58, 97].
Due to the long resection time, Rad53p becomes hyperphosphorylated and the cells
accumulate in G2/M [58]. By deleting genes in this background, one can study genes
required for repair by SSA. If a gene is required for this process, its deletion will cause
sensitivity to growth in the presence of galactose, which induces the HO endonuclease.
Moreover, various assays can be performed to assess the function, if any, of a particular
protein during the time course of DSB induction, resection, repair, and checkpoint recovery.
1.9 Conservation of DSB repair pathways in higher
organisms
Studies on yeast chromatin remodelers and other proteins associated with DNA repair
have gained momentum due to the conservation of these processes in higher organisms.
Although yeast is single-celled, the basic cellular process and response to induction of a
24
DNA DSB is highly conserved. The first line of defense, carried out by the checkpoint
proteins, is very similar between yeast and mammalian cells. Most of the proteins involved in
signal transduction following induction of a DNA DSB in yeast have homologs in
mammalian cells. Even the pattern of DDR, which involves Rad53p and γH2AX
phosphorylation, is similar between the two distantly related species. Hence, using this
relatively simple organism, much can be understood about cells’ reactions to a DSB and the
events that follow, even though recombinational repair is only a minor repair pathway in
mammalian cells. The consequences of inefficient repair can be very deleterious for
mammalian cells, because erroneous DNA damage responses can cause failure in the
transmission of the genome from one generation to the next, resulting in genomic instability,
a hallmark of cancer [154].
1.10 Thesis Rationale
The LCD algorithm applied to the E-MAP data for genes involved in chromosome
biology suggested many uncharacterized functional connections. Histone chaperone Asf1p
and its major functional partner, the HAT Rtt109p, has been of immense interest to the
Greenblatt laboratory [22, 43, 155]. A functional connection was predicted by the LCD
algorithm among genes involved in the histone H3K56 acetylation pathway, i.e. Asf1p and
Rtt109p, and several genes required for efficient sister chromatid cohesion, i.e. Ctf18p,
Ctf8p, Dcc1p and Ctf4p. In other words, all these genes share similar patterns of genetic
interactions. Furthermore, the bicluster of these genes, i.e. Asf1p, Rtt109p, Ctf18p, Ctf8p,
Dcc1p and Ctf4p, also included additional proteins with similar patterns of genetic
25
interactions, including core genes involved in DNA repair and those required for DNA
resection at the DSB (the MRX complex), as shown in Figure 2. As well, the genes that
interacted with Asf1p, Rtt109p, Ctf18p, Ctf8p, Dcc1p, Ctf4p and the other genes that shared
similar patterns of genetic interactions included the RAD group of genes and many other
genes that could be classified as DNA damage repair genes, e.g. Pph3p, Psy2p and Hst4p,
suggesting their involvement in a functional DNA repair pathway. It has also become very
apparent that the process of DNA damage repair is not limited to a few proteins specialized
in this function, but also involves a number of other proteins linked to chromosome structure
and dynamics. For example, the presence of phosphorylated H2AX, a variant of histone H2A
that is inserted into chromatin surrounding the site of DNA damage, leads to the recruitment
of chromatin remodelers [156]. Also, the cohesin complex, which holds sister chromatids
together for faithful segregation of the two sister chromatids, is also present at the site of a
DNA DSB to ensure proper repair by HR [122, 124-127].
Histone H3K56 acetylation is important for genomic stability, as cells lacking either
its HAT, Rtt109p, or its chaperone, Asf1p, are highly sensitive to DNA damaging agents [44,
46, 48, 157, 158]. It has been speculated that H3K56 acetylation has an important role in
chromatin reassembly after the repair of damaged DNA, ensuring the cells turn off the
checkpoint and resume cell cycle progression [97, 102]. On the other hand, the lack of
cohesion promoting proteins, including Ctf18p, Ctf8p, Dcc1p and Ctf4p, also causes
sensitivity to DNA damaging agents, including MMS and HU [129, 146, 149, 150]. Ctf18p,
Ctf8p and Dcc1p have also been implicated in the repair of DNA DSB’s by the HR method,
and Ctf4p has been linked to the H3K56 acetylation pathway after DNA damage is induced
26
by HU [25, 149, 159]. While some of these proteins may be found at the site of DNA repair,
the nature of their involvement is not fully understood.
In spite of the wealth of knowledge about proteins required during the process of
DNA DSB repair, relatively little is known about how chromatin structure is reestablished
after repair is complete. Recent findings have indicated that Asf1p- and Rtt109p-dependent
histone acetylation on H3K56 is important for chromatin reassembly following repair. Based
on the LCD algorithm, there is the possibility that Ctf18p, Ctf8p and Dcc1p work together
with Ctf4p, Asf1p and Rtt109p in DNA repair. The studies in this thesis were designed to test
this hypothesis. In particular, I found that, although Ctf4p and the Ctf18-complex are
apparently not required for the repair of a DSB, they are required for chromatin reassembly
and checkpoint recovery after repair by SSA has taken place.
27
Chapter 2
Materials and Methods
28
2 Material and Methods
2.1 Yeast transformations
5 ml of overnight cultures of cells were grown in rich medium (YPD) to saturation. In
the morning, the cells were diluted to an OD600 of 0.2 and grown for about 4-5 hours to mid-
log phase. The cultures were harvested by centrifuging at 4,000 rpm at 4oC in a 50 ml Falcon
tube for 5 mins to separate the liquid culture medium from the cells. The pellets were then
washed with 10 ml of ice-cold sterile water and centrifuged again as described above. The
pellets were then suspended in 1ml 100 mM lithium chloride (LiCl) solution and transferred
to an Eppendorf tube. The cells were then centrifuged in a mini-centrifuge for 15 sec at
maximum speed at room temperature. The supernatant was discarded and cells were
suspended in 400 µl of 100 mM LiCl. The cells were then mixed in the LiCl solution to form
a homogeneous solution. 50 µl of cells in solution were transferred to new Eppendorf tubes,
where each tube was used for one transformation. All the tubes were then centrifuged in a
mini-centrifuge for 15 sec at maximum speed. The supernatants were discarded and to each
tube the following were added in order: 240 µl of 50% poly-ethylene glycol (PEG), 36 µl of
1M LiCl, 50 µl of 20% Salmon Sperm DNA (1:5 dilution of 10 mg/ml salmon sperm DNA
was prepared in distilled water and heated at 68oC for 5 mins) and 50 µl of purified DNA
(50ng/µl) (50 µl of distilled water for the control). The transformation mixture was then
mixed using a wooden stick and briefly vortexed for 10 sec. The cells were incubated at 30oC
for 30 mins after which 40 µl of dimethylsulfoxide (DMSO) was mixed in by gently flicking
the tubes. The mixture was then put in a water bath to heat shock at 42oC for 15 mins.
29
Finally, the contents were centrifuged at 8,000 rpm for 1 min to remove the supernatant. The
cell pellet was resuspended in 100 µl of sterile water and plated on YPD rich medium. On the
following day, the plates were replica plated on selective media.
2.2 Yeast strains and strain construction
The strains used in this study are listed in Table 1. The single deletions of yeast
genes, substituted with a KAN or NAT marker, were taken from the E-MAP strain collection
of the laboratory or purchased from Open Biosystems [22]. The YMV2 strain set up for an
inducible DSB was kindly provided by the Haber laboratory. Single mutations and gene
tagging were done in this strain.
The gene to be tagged or deleted was PCR amplified along with a small region of
homology on either side. The PCR product was run on a 0.8% agarose gel to confirm the
presence of a DNA fragment corresponding in size to the marker plus the flanking regions of
the gene. The DNA was purified using the commercially available Qiagen purification kit
before transformation.
Table 1. List of stains used in this study
Strain Genotype Source
Y3656 MATα, ura3Δ0, leu2Δ0, his3Δ1, met15Δ0, lys2Δ0,
can1ΔMFA1pr-HIS3-Mfα1pr-LEU2 Tong et al., 2001
AT131 Y3635 asf1Δ::NAT Collins et al., 2007
30
AT398 Y3635 rtt109Δ::NAT Collins et al., 2007
AT233 Y3635 rad52Δ::NAT Collins et al., 2007
AT656 Y3635 ctf18Δ::NAT Collins et al., 2007
AT669 Y3635 ctf8Δ::NAT Collins et al., 2007
AT479 Y3635 ctf4Δ::NAT Collins et al., 2007
AT54 Y3635 pph3Δ::NAT Collins et al., 2007
AT47 Y3635 rtt101::NAT Collins et al., 2007
AT355 Y3635 hir1::NAT Collins et al., 2007
AT255 Y3635 rtt106::NAT Collins et al., 2007
AT153 Y3635 hir2::NAT Collins et al., 2007
AT260 Y3635 hir3::NAT Collins et al., 2007
AT162 Y3635 hpc2::NAT Collins et al., 2007
BY4741 MATa his3Δ1 leu2Δ0 met15Δ0 ura3Δ0 Open Biosystems
YHS001 Y3635 asf1Δ::NAT, ctf18∆::KAN This study
YHS002 Y3635 asf1Δ::NAT, ctf8∆::KAN This study
YHS003 Y3635 asf1Δ::NAT, ctf4∆::KAN This study
YHS004 Y3635 rtt109Δ::NAT, ctf18∆::KAN This study
YHS005 Y3635 rtt109Δ::NAT, ctf8∆::KAN This study
YHS006 Y3635 rtt109Δ::NAT, ctf4∆::KAN This study
31
YMV2
MAT! ho hml::ADE1 mata::hisG hmr::ADE1
his4::-URA3-leu2-(Xho1to Asp718)-his4
leu2::HOcs ade3::GAL::HO ade1 lys5 ura3-52
Vase et. al., 2002
YHS007 YMV2 asf1Δ::NAT This study
YHS008 YMV2 rtt109Δ::NAT This study
YHS009 YMV2 ctf18∆::KAN This study
YHS010 YMV2 ctf8∆::KAN This study
YHS011 YMV2 dcc1::KAN This study
YHS012 YMV2 ctf4∆::KAN This study
YHS013 YMV2 rad52Δ::NAT This study
YMV80
MAT! ho hml::ADE1 mata::hisG hmr::ADE1
his4::-URA3-leu2-(Xho1to Asp718)-his4
leu2::HOcs ade3::GAL::HO ade1 lys5 ura3-52
Vase et. al., 2002
YHS015 YMV80 asf1Δ::NAT This study
YHS016 YMV80 rtt109Δ::NAT This study
YHS017 YMV80 ctf18∆::KAN This study
YHS018 YMV80 ctf4∆::KAN This study
W303-1a MATa ade2-1 can1-100 his3-11,15 leu2-3,112
trp1-1 ura3-1 Celic et. al.,2008
YHS020 W303-1a asf1Δ::NAT This study
YHS021 W303-1a rtt109Δ::NAT This study
32
YHS022 W303-1a ctf18Δ::NAT This study
YHS023 W303-1a ctf8Δ::NAT This study
YHS024 W303-1a ctf4Δ::NAT This study
YHS025 W303-1a pph3Δ::NAT This study
HMY221 MATa ade2-1 can1-100 his3-11,15 leu2-3,112
trp1-1 ura3-1 hst3D::his5+ hst4D::kanMX6 Celic et. al.,2008
YHS030 HMY221 hst3D::his5 hst4D::kanMX6
asf1Δ::NAT This study
YHS031 HMY221 hst3D::his5 hst4D::kanMX6
rtt109Δ::NAT This study
YHS032 HMY221 hst3D::his5 hst4D::kanMX6
ctf18Δ::NAT This study
YHS033 HMY221 hst3D::his5 hst4D::kanMX6
ctf8Δ::NAT This study
YHS034 HMY221 hst3D::his5 hst4D::kanMX6
ctf4Δ::NAT This study
YHS035 HMY221 hst3D::his5 hst4D::kanMX6
pph3Δ::NAT This study
YHS036 YMV2, CTF18-13MYC::KAN This study
YHS037 YMV2, CTF4-13MYC::KAN This study
YHS038 YMV2, CTF8-13MYC::KAN This study
YHS039 YMV2, ASF1-13MYC::KAN This study
33
YHS040 YMV2, RTT109-13MYC::KAN This study
YHS041 YMV2, CTF18-13MYC::KAN, asf1::NAT This study
YHS042 YMV2, CTF18-13MYC::KAN, rtt109::NAT This study
YHS043 YMV2, CTF4-13MYC::KAN, asf1::NAT This study
YHS044 YMV2, CTF4-13MYC::KAN, rtt109::NAT This study
YPR135W BY4741, CTF4-TAP::HIS3 TAP fusion library
YHS045 BY4741, CTF4-TAP::HIS3, rtt109::NAT This study
YMR078C BY4741, CTF18-TAP::HIS3 TAP fusion library
YHS046 BY4741, CTF18-TAP::HIS3, rtt109::NAT This study
BSY678 ASF1-vsv Suter et. al., 2007
YHS047 BY4741, CTF4-TAP::HIS3, ASF1-vsv This study
YHS048 BY4741, CTF18-TAP::HIS3, ASF1-vsv This study
YHS049 BY4741, DCC1-TAP::HIS3, ASF1-vsv This study
2.2.1 Construction of single and double mutants
For constructing single mutants (yeast strains YHS007-YHS025) a selectable marker
was PCR amplified and transformed using the protocol described above. For gene deletion,
primers were constructed 250 base pairs upstream and downstream of a gene, which had
been previously replaced by a selectable marker as described in Table 2. For making double
34
deletions (yeast strains YHS001-YHS006 and YHS030-YHS035), the transformation
protocol described above was used on a strain carrying the first deletion. In this case, a
different marker with gene-specific flanking regions was PCR amplified and transformed.
The cells were replica plated after the transformation on rich medium carrying one or two
selective agents and colonies were picked for testing.
Table 2: List of PCR primers used for deletion and confirmation
Name Sequence
Asf1 – F CTCATCCTAAACGCGTAAATCTGTT
Asf1 – R TTAATTCGTTGAACGTGCCGCATCC
Asf1 – conf CTGTTTTATTCCGTTCTTACATGGG
Rtt109 – F TCTTGCTTCTGAGATGCATACAATT
Rtt109 – R TCAAGTTTTAGGCAAGGCTTTAGCT
Rtt109 – conf CTGTACCTTTTAGCCTAAGCGCCAA
Ctf4 – F ACCTGGAGAATGGGTGTTTTCACTA
Ctf4 – R TTATTTCAATTGCTGTTCATATCTA
Ctf4 – conf TATGTCCAATTTGAGTGTAAAATCA
Ctf18 – F AGGAAAGGACACTTAAGACACAAAC
Ctf18 – R TTATTCCCACAGGTTATTCCAAGTC
Ctf18 – conf TATCTGAAGAATGTGAATATTGTCA
35
Ctf8 – F ATCCGATCTCCTCTTTGTATGATTT
Ctf8 – R GGTAGAGGCCTATCCTTAAAGATAA
Ctf8 – conf TGACTCAAGATGGATTGGCTTTTCT
Dcc1 – F AGGAGTAGCATACCACAGCGATATT
Dcc1 – R CTACCTGCTCGTGACCACGGTCTTT
Dcc1 – conf ACTGGCAATCGCTCAACATGACCTA
Rad52 – F TCTGCTCTTCCCGTTAGTGATTCTC
Rad52 – R TAGGCTTGCGTGCATGCAGGGGATT
Rad52 – conf ACTAAATGGTTGAATCGGGTCTTGC
NAT – F ACATGGAGGCCCAGAATACCCT
NAT – R CAGTATAGCGACCAGCATTCAC
KANB CTGCAGCGAGGAGCCGTAAT
KANC TGATTTTGATGACGAGCGTAAT
Rtt101 – F GGATGTTCAATATTCCTGATTAAGT
Rtt101 – R AGTGTAACTGTTGTATTGAACGTGG
Rtt101 – conf TGGTGCAATGTATGAGGTTTCGCTG
Rtt106 – F TTTAAAAATTTAAATATAAACGAAT
Rtt106 – R TCAATCGTATTCTACTCCGGATCCA
Rtt106 – conf GGAATTTGCCGAATGATATCAAGAG
36
2.2.2 Construction of tagged strains
A plasmid pFA6a-13myc-kanMX6 containing 13 MYC epitopes was kindly provided
by the Longtine lab [160]. Primers (70 bases) for tagging contained the 50 base sequence
upstream or downstream of the stop codon of the gene to be tagged, as well as homology to
one side of the tag and a selectable marker to ensure that the tag was inserted just before the
stop codon of the gene. Using PCR, two 50 base pair DNA sequences containing the gene-
specific flanking regions were added to the tag and the marker. The tag was inserted by
transformation. After replica plating the cells on appropriate selective medium, cells were left
to grow for a few days and colonies were picked for testing.
Table 3: List of PCR primers used for tagging genes
Tagging Primer
Name
Sequence
pCtf8-myc-F ATGTAATGAAATATAAGGTTATCTTTAAGGATAGGCCTCT
ACCTATTATGCGGATCCCCGGGTTAATTAA
pCtf8-myc-R AGTCTGCGCCAAATAACATAAACAAACAACTCCGAACAA
TAACTAAGTACGAATTCGAGCTCGTTTAAAC
pRtt109-myc-F CGATAACAATGCTAAAACCGCGTAAAAAAGCTAAAGCCT
TGCCTAAAACTCGGATCCCCGGGTTAATTAA
pRtt109-myc-R TCTAAGATCGATGCTACATACGTGTACTAAATAATAAATA
TCAATATGTAGAATTCGAGCTCGTTTAAAC
37
pCtf18-myc-F GGTTCTCTAACGCTGTCAGGAAAAATGTGACTTGGAATA
ACCTGTGGGAACGGATCCCCGGGTTAATTAA
pCtf18-myc-R CATATACAAGTATGCTTCTTAAGAGAGACTGCGTATATAT
CTTACGTCATGAATTCGAGCTCGTTTAAAC
pCtf4-myc-F TTAAAAAAATTAATAATATAAGGGAAGCTAGATATGAAC
AGCAATTGAAACGGATCCCCGGGTTAATTAA
pCtf4-myc-R TGAACAGGTATCAAATAATTGTCTCTTGCGTATATATATT
TTACATTTTTGAATTCGAGCTCGTTTAAAC
pRtt101-myc-F TACGAGACAAATTCATAACTAGGGACGAATCAACAGCAA
CTTACAAGTACCGGATCCCCGGGTTAATTAA
pRtt101-myc-R GGATTATAAACTATCTCAGTAGTTAGGTAATATATAAGAT
GGCACCAGTCGAATTCGAGCTCGTTTAAAC
pAsf1-myc-F GGTCCACGGATATTGAATCCACTCCAAAGGATGCGGCAC
GTTCAACGAATCGGATCCCCGGGTTAATTAA
pAsf1-myc-R TAAAGTGTACCTCTCTTGCAGGTACCATTAATCTTATAAC
CCATAAATTCGAATTCGAGCTCGTTTAAAC
pRtt106-myc-F ATAATGATGACGATGAAGATGATGAGGATGGATCCGGAG
TAGAATACGATCGGATCCCCGGGTTAATTAA
pRtt106-myc-R TGAACTCTTACATATGCGTATTCATGCTATATTATAATAT
CGAATCTAAGGAATTCGAGCTCGTTTAAAC
38
2.2.3 Confirmation of the mutant/tagged strains
For confirming the presence of a tag, 5 ml cultures for all 16 colonies that were
picked for each tag were grown in test tubes overnight in YPD. In the morning, the cells were
centrifuged at 4,000 rpm at 4oC for 4 mins. The supernatant was discarded and the cells were
washed once with distilled water and centrifuged again, keeping the cells on ice. The water
was discarded and 350 μl of 1X SDS loading buffer was added to the pellet. The cells were
immediately re-suspended in the buffer and boiled for 10 mins. The samples were loaded on
10% SDS-polyacrylamide gels and probed for the tag using western blotting.
For confirming deletions, about 8 single colonies were picked from each plate after 2
days, then re-streaked and re-grown. The deletion was confirmed by PCR, where one primer
hybridized about 350-400 bp upstream of the gene and the other within the replacing NAT or
KAN cassette to confirm the correct integration of the insertion cassette. The PCR product
was run on a 0.8% agarose gel to confirm the presence of DNA of the appropriate length.
2.3 Growth sensitivity assays
To assess possible growth defects, single and double mutants were tested for growth
in the presence of MMS, which causes DNA damage. MMS was added to YPD plates at
concentrations of 0.005%, 0.0075% and 0.01%. For the growth assay, 5 ml cultures of cells
were grown overnight at 30oC. In the morning, the cells were diluted to OD600 of about 0.1
and grown until mid-log phase (OD600~0.2). The cultures were then harvested and plated in
5-fold serial dilutions on YPD plates at 30oC.
39
To assess possible growth defects associated with induction of a single double-strand
break, the YMV2 strain and its derivatives were plated on medium containing galactose,
rather than glucose, as carbon source. Photos were taken periodically for up to three days.
2.4 Analysis of DNA DSB repair by SSA
The yeast YMV2 strain carries a galactose-inducible HO endonuclease, which causes
a single double-strand break. The sequence of the DNA flanking the break site is known, and
the homologous sequence was genetically engineered 30 kb away. To assess DSB formation
and its subsequent repair, primers flanking both homologous sites were used [78]. Cells were
grown in raffinose, so that galactose could be directly added to the medium for induction
without washing the cells. This induction occurs in less than 5 mins after adding galactose to
the medium. Cells were first grown in YPD for about 8-9 hours. In the evening, the cells
were washed and inoculated into fresh medium containing 2% raffinose, 0.05% glucose and
0.03% glycerol. The cells grow very slowly in this medium and hence all the mutants were
inoculated at an OD600 of 0.1, which resulted in an approximate OD600 of 0.5 after about 12-
14 hours (next morning). The HO endonuclease in the mid-log phase cells was then induced
by adding galactose to 2%. Aliquots of the cultures were harvested every 2 to 4 hours for
about 24 to 30 hours to assess the DNA damage and its repair. At each time point, samples
were also taken for repair analysis, western blotting and chromatin immunoprecipitation.
About 5 ml of the cultures were harvested for preparation of genomic DNA by the
phenol-chloroform extraction method. Cells were centrifuged at 4,000 rpm for 5 mins. The
pellets were suspended in 10 ml of ice-cold water and centrifuged again. The pellets were
40
then resuspended in 350 µl breaking buffer (2% Triton X100, 1% SDS, 100 mM NaCl, 10
mM Tris pH8.0, 1 mM EDTA) and transferred to Eppendorf tubes. 350 µl of
phenol/chloroform (25:24:1 phenol/chloroform/isoamyl alcohol) and 350 µl of glass beads
were added to each tube. The tops of the tubes were sealed with parafilm and then they were
vortexed at highest speed for 5 mins (one mins vortex, one mins on ice, repeated 5 times).
350 µl of 1X TE buffer (49.4 ml dH2O, 500 µl 1M Tris pH 8.0, 100 µl 0.5 M EDTA) were
added to each tube and they were vortexed again for 2 mins at highest speed. The tubes were
centrifuged at 12000 rpm for 5 mins at room temperature, and the supernatants were
collected. 1 ml of 100% ethanol was added to each supernatant, and the tubes were
centrifuged for 5 mins at maximum speed at room temperature. The ethanol was removed
and the pellets (seen as white precipitates along the sides of the tubes) were resuspended in
500 µl of 70% ethanol and centrifuged again. The supernatants were removed and the pellets
were left in the hood to dry for about 2 hours. Finally, 200 µl of elution buffer (5 ml 10%
SDS, 1 ml 1M Tris pH 8.0, 1 ml 0.5M EDTA, 43 ml dH2O) was added to each pellet. Since a
large number of extracts for SSA analysis had to be prepared within a relatively short time
for time course experiments, the commercially available MasterPure DNA extraction kit
(Epicenter) was used later for genomic DNA preparations. The primers around the site of the
break listed in Table 4 were used for the PCR analysis of break formation and subsequent
repair.
41
Table 4: PCR primers around the DSB used for analysis of DNA repair by SSA (from
Keogh et al., 2006)
Name Sequence
SSA P1 GCTGGGAAGCATATTTGAGAAGATGCG
SSA P2 TGGGTTGAAGGCTCTCAAGGGCATC
SSA P3 GGTGACCACGTTGGTCAAGAAATCA
SSA P4 GGTGACCACGTTGGTCAAGAAATCA
Rad3 – F GATAAGATTGCGACAAAAGAGGATA
Rad3 – R GTGGGACGAGACGTTTAGATAGTAA
2.5 Whole-cell extraction and western blotting
5ml of cells were grown overnight at 30oC. The next morning cells were diluted into
50ml of fresh medium and grown to mid-log phase at 30oC. Alternatively, aliquots were
removed from cultures used to assess DNA repair by SSA. Cultures were centrifuged at 4000
rpm for 5 mins at 4oC. Cells were then washed once with 1ml of 20% trichloroacetic acid
(TCA). Cells were then either frozen at -80oC or used immediately for protein extraction. The
pellet was thawed (if frozen) and resuspended in 250 μl of 20% TCA, then added to an equal
volume of glass beads. The mixture was chilled briefly on ice and then vortexed three times
in the cold room for 1 min each, with 1 min cooling on ice in between. The TCA-precipitated
42
lysates were then filtered into 15 ml Falcon tubes by centrifuging at 1,250 rpm for 1 min at
4oC to remove the glass beads. The beads were washed once with 300 μl of 5% TCA at the
same speed. The lysates were then transferred to Eppendorf tubes for centrifugation at 14,000
rpm for 15 mins at 4oC. All precipitates were then aspirated. At this point the pellets can
either be frozen and stored at -80oC for later use or resuspended in 500 l of fresh 1X SDS
gel sample buffer (60 mM Tris pH 6.8, 2% SDS, 10% glycerol, 100 mM DTT and a pinch of
bromophenol blue) and 40 μl 1M Tris-HCl, pH 8.0 (to neutralize the TCA). The samples
were boiled for 5 mins before loading onto a SDS polyacrylamide gel for western blotting.
The percentage of polyacrylamide gel used for western blotting depended on the
protein to be detected. For detection of Rad53p phosphorylation, 6% polyacrylamide gel was
used, for detection of Ctf18p, Dcc1, Ctf4p and Asf1p proteins, 8% polyacrylamide gel was
used, while for the detection of histones 15% polyacrylamide gel was used. 10 μl of boiled
samples were loaded onto the gel. After electrophoresis the proteins in the gels were
transferred onto nitrocellulose paper. The blots were washed twice with TBS buffer for 5
mins. The blots were blocked overnight at 4oC in blocking solution (5% milk in TBS buffer).
The blots were washed thrice with TBS buffer in the morning for 5 mins and enough primary
antibody buffer (3% BSA, 0.01%NaAz, 0.5% Tween-20, primary antibody in TBS buffer)
was added to cover the entire blot. The blot was left in the primary antibody for 1 hour, after
which the blots were washed thrice with TBS buffer. To the blots, 3% milk in TBS buffer
with the secondary antibody was added and the secondary antibody was left for 45 mins,
after which the blots were washed thrice with TBS buffer for 5 mins and made
chemiluminescent by using ECL.
43
The antibodies used in this study were α-H3 (Abcam #ab1791) used in 1:5000
dilution, α-H3K56Ac (Upstate #07-677) used in 1:10000 dilution, α-γH2AS129 (Abcam
#ab15083) used in 1:20000 dilution and α-Rad53 (Santa Cruz #sc-6749) used in 1:2000
dilution.
2.6 Chromatin fractionation
5 ml of cells were grown overnight at 30oC. The next morning cells were diluted into
50 ml of fresh medium and grown to mid-log phase at 30oC. Cells were collected by
centrifugation. Pellets were washed once with 10 ml cold PBS buffer. The pellets were then
resuspended in 1.5 ml cold CBS buffer (50mM HEPES, 150 ml NaCl, 0.8M Sorbitol) and 15
μl 1M DTT. The suspension was then incubated for 10 mins at 30oC. The suspension was
then centrifuged for 2 mins at 2,000 rpm. The pellets were resuspended in 1.5 ml cold SB
buffer (50 mM HEPES, 0.8M Sorbitol) and 15 μl 1M DTT. To the suspension, 20 μl of
10mg/ml freshly prepared zymolase was added. The suspension was again incubated for 10
mins at 30oC. The suspension was centrifuged again for 2 mins at 2,000 rpm at 4
oC. Ice cold
SB buffer was added again to the pellets, without re-suspending the pellets. The buffer was
added on the opposite side of the pellet, such that the pellet spun through the buffer. The
pellet was washed again with ice cold SWB buffer (100 mM KCl, 50 mM HEPES, 2.5 mM
MgCl2, 0.6M Sorbitol), without re-suspending the pellets. Again the buffer was added on the
opposite side of the pellet. The pellets were then resuspended thoroughly with cut-off P200
pipette tip in 100 μl ice-cold EB buffer (100 ml KCl, 50 mM HEPES, 2.5 mM MgCl2) with
protease inhibitors (50 mM NaF, 2 mM PMSF, 2 μg/ml pepstatin A, 100 mM Na o-vanadate,
44
5 μg/ml leupeptin, 5 μg/ml TLCK, 2.5 μg/ml aprotinin). 25 μl of the suspension was saved
for whole cell extract (WCE).
To the remaining suspension, 83 μl of 10% Triton-X was added and incubated on ice
for 5 mins. The suspension was mixed occasionally by inversion. The suspension was
centrifuged for 20 mins at 14,000 rpm at 4oC. About 25 μl of the supernatant, corresponding
to the nucleoplasmic fraction (or soluble fraction) was saved. The pellets were then washed
with 50 μl of EB buffer (without Triton-X) and centrifuged for 5 mins at 10,000 rpm at 4oC.
Pellets were again resuspended in 83 μl of EB buffer (without Triton-X). This fraction
corresponded to the chromatin fraction and was saved.
An equal amount of SDS loading dye was added to each fraction saved and boiled for
5 mins. About 10 μl of each fraction was run on a SDS-PAGE gel. To assess for TAP-tagged
Ctf18p or Ctf4p, 10% SDS gel was used while for histones 15% SDS gels were used. Blots
were blocked in 5% milk overnight, then washed three times with 1X TBS buffer. To the
blots, primary antibody solution (3% BSA solution, 0.01% NaAz, 0.1% Tween-20, primary
antibody in 1X TBS) was added. Blots were washed three times after 1 hour. After washing
the blots, secondary antibody solution (3% milk and secondary antibody in 1X TBS) was
added. The blots were washed after 45 mins and washed again three times. The dilution used
for TAP antibody was 1:5000 and the dilution used for histone H3 antibody was 1:10000.
2.7 Chromatin immunoprecipitation (ChIP)
The growth conditions for ChIP varied with the strain that was used. For YMV2
strains about 45ml of cells were harvested at the desired time points up to 24 hours, with the
45
first sample being taken just before the addition of galactose. Cultures were crosslinked by
adding 37% formaldehyde to a final concentration of 1% for 20 mins. 6.5ml of 3M Glycine,
20mM Tris base solution was then added to stop the crosslinking reaction. After 5 mins of
quenching at room temperature, each sample was transferred to a 50mL Falcon tube. The
cells were centrifuged at 4,000 rpm for 5 mins at 4oC. Pellets were then washed twice with
ice-cold water and centrifuged at 4000 rpm for 5 mins. The pellets were finally washed with
1 X FA buffer (50 mM Hepes-KOH, pH7.5, 150 mM NaCl, 1mM EDTA, 1% Triton X-100,
0.1% sodium deoxycholate, 0.1% SDS) and centrifuged again. Pellets were left on dry ice to
freeze and then stored at -80oC.
Frozen pellets were thawed on ice. 1ml of 1 X FA buffer containing 100 mM protease
inhibitors (2.4µg/ml Chymostatin, 1.5µg/ml Pepstatin A, 87µg/ml PMSF, 0.5µg/ml
Leupeptin, 17µg/ml Aprotinin, 310µg/ml Benzamidine) and 50 mM PMSF was added to the
pellets, which were then transferred to a 2ml screw cap tube. Enough glass beads were added
to completely fill the tube. Cells were lysed using a bead beater 16 times for 2 minutes, with
1 min in between on ice. Using a needle, holes were punched into the bottoms of the tubes to
collect the lysates and leave the glass beads behind. To accomplish this, the 2ml screw cap
tubes were placed into 5ml syringes, which were mounted on 15ml Falcon tubes. The cells
were then collected by briefly centrifuging the tubes for 1 min at 4oC. Using a 1ml pipette,
the lysates were collected and transferred to new Eppendorf tubes. Lysates were then
centrifuged for 30 mins at 14,000 rpm at 4oC. After the centrifugation, a clear jelly-like layer
of chromatin was seen on top of the cell pellets. The supernatants were then removed and
750ul of fresh 1 X FA buffer was added. Using a wooden stick, the lysates were mixed with
the buffer. The lysates were sonicated (Branson Digital Sonifier) to solubilize the chromatin
46
for 25 pulses 6 to 8 times at a setting of 30% Amp, 0.35 cycle, with 1 min in between on ice,
to yield about 300 to 400 bp fragments. Care was taken not to overheat the samples. The
lysates were then centrifuged again for 30 mins at 14,000 rpm at 4oC. The supernatants
containing the chromatin fractions were collected into new tubes and diluted with 1ml of 1 X
FA buffer to final volumes of 1.75ml. This was aliquoted into 2 tubes of 750 µl, each of
which was used for one IP. About 250 µl was stored for Input, and they were all frozen in
liquid nitrogen if not used for IP the same day.
For IP, one 750 µl sample was thawed on ice (if frozen) and about 3 to 5 µl of the
appropriate antibody was added. For myc-tagged strains 5 μl of α-myc (Santa Cruz #sc-
56634) and 3 μl of α-H3 (Abcam # ab15083) were used. The samples were rotated overnight
in the cold room at 4oC. The next morning 40 μl of 50% slurry of protein-G Sepharose
(Sigma), reconstituted with 1 X FA buffer, was added to each sample. The beads were
allowed to bind to the antibody for 2 hours with rotation in the cold room at 4oC. The
protein-G Sepharose beads were subjected to 5 rounds of washing: twice with 1 X FA buffer
(275mM NaCl); once with 1 X FA buffer (500mM NaCl); once with ChIP wash buffer
(10mM Tris-HCl pH 8.0, 0.25M LiCl, 1mM EDTA, 0.5% NP-40, 0.5% sodium
deoxycholate) and finally once with TE buffer (500mM Tris pH 8.0, 0.5mM EDTA). For
each washing, 1ml of buffer was added to the beads and the solution was left on the nutator
for 5 mins in the cold room at 4oC. To collect the beads, samples were gently centrifuged at
2000 rpm for 2 mins at 4oC. After all the washing steps, the immunoprecipitates were eluted
into 250 μl of elution buffer (5ml 10% SDS, 1ml 1M Tris pH 8.0, 1ml 0.5M EDTA in 50ml
water). The beads were then incubated for 10 mins at 65oC. The beads were briefly flicked
with the finger and incubated again for 10 mins at 65oC. The contents were then centrifuged
47
at 5000 rpm for 5 mins and the supernatants collected in PCR tubes. Another 250μl of sterile
water was added to the beads. The suspensions were mixed and centrifuged again at 5000
rpm for 5 mins, then supernatants were added to the previous samples to a total volume of
500 μl. For the Input sample, 50 μl of Input DNA sample was added to 200 μl of elution
buffer and 250 μl of sterile water in a PCR tube. The chromatin and Input samples were de-
crosslinked by adding 10 μl of 20mg/ml proteinase K (Fermentas) and incubating for 2 hours
at 42oC followed by 8 hours at 65
oC. The de-crosslinked DNA was purified using Qiagen
PCR Purification Kits and the resulting samples were analyzed using radiolabelled PCR.
Table 5: List of PCR primers spanning 20 kb around the site of DNA DSB used for
ChIP
Name Sequence
Tel VI - F GGATTTTACCAACGACTTCGTCTCA
Tel VI - R CGCTATTCCAGAAAGTAGTCCAGC
HO Cut-F CCAAATCTGATGGAAGAATGGG
HO Cut-R CCGCTGAACATACCACGTTG
HO+19.3kbF GGGCTCCTCAACCTCTCTCT
HO+19.3kbR TACTGCAGAGGCGTGTTTTG
HO+17kbF CCAAGAACCCCAAGTTGAAA
HO+17kbR ACAATCGGCAATCCACTCTC
48
HO+14.3kbF TGCTTTTGGCAGCATCATAG
HO+14.3kbR CTCCAGGCTCATCATGGAAT
HO+12.1kbF CTTCCTCGGTGCTTGTCTTC
HO+12.1kbR TGAAGCACGAGCATTTGAAC
HO+10.2kbF TCGATATGGCATCTTGGACA
HO+10.2kbR GGTACAGTGGGCGAAGTTGT
HO+2.8kbF TTCTAGCGCAGAACATGGTG
HO+2.8kbR GGACTGTCCAAGCCGTACAT
HO+0.5kbF AGGCTGAACCCGAGGATAAT
HO+0.5kbR CGGATCTCCAGATCATCGTT
HO+2.2kbF TTTTGCCCAGTCTTTTACGG
HO+2.2kbR GCGAGGCTATCATTTCAAGC
HO+9.6kbF ATGGTTCGGTTGGTGCTTAG
HO+9.6kbR TCGACTTGTTTGGGCCTATC
HO+5.6kbF ACGAGCTTTGCGCTTATGAT
HO+5.6kbR CTTTGGCCATCGATGAGTTT
HO+7.3kbF GGAAGGACGCAAAGGACATA
HO+7.3kbR GACTAGCCAAGCGTTTCCAG
HO+1.3kbF AAGCATCGCTTATTGCGACT
49
HO+1.3kbR CTCCAGGTCAACCAGGTCAT
HO-2.9kbF TGGATTCACTGTTGGACGAA
HO-2.9kbR CGTCTACCTTCACTGCACGA
HO-3.1kbF CGTGCAGTGAAGGTAGACGA
HO-3.1kbR CGTGCACCAAAAGAAGTTGA
HO-9.2kbF TTTCAACCCGTATTGCTTCC
HO-9.2kbR GTGAGAAAATCGCTGCTTCC
HO-5.3kbF GGCCAATCTGTCGCTAACAT
HO-5.3kbR TCTCGGTGACATCATTCCAA
HO-7.9kbF AAACAATAGCCGCCAAACTG
HO-7.9kbR CCTCAATTCCCTTTTGTGGA
HO-6.1kbF AGGCGCTACCATGAAGAGAA
HO-6.1kbR GCTGAAACGCAAGGATTGAT
HO-11.7kbF CTAGCGCATGGCAGTATGAA
HO-11.7kbR TTCAACAACAACGGAAACCA
HO-13.9kbF TGGATGATGGTTTTGGGTTT
HO-13.9kbR CTTCTGCATAACCACGAGCA
HO-17.1kbF ATCGGACTGTGGCGTTTTAC
HO-17.1kbR CCAGGTAACTCCGGTTTCAA
50
HO-16.0kbF CACCCATGTTGTGATCGAAG
HO-16.0kbR GTGGTAATTCGGGCTTCAAA
HO-18.5kbF GCATCCAGAGGCGTGTTATT
HO-18.5kbR CCAGCATGCCCTCGTATAAT
HO-1.3kbF AAGCATCGCTTATTGCGACT
HO-1.3kbR CTCCAGGTCAACCAGGTCAT
2.8 One step TAP-Tag purification
One liter of TAP-tagged Ctf18p, Ctf8p and Dcc1p strains were grown in YPD
medium until the OD600 reached 0.8 (exponential phase). Cells were collected by
centrifugation and successively washed with 100ml of cold distilled water and 50ml of cold
Lysis buffer (20mM Tris Cl pH 7.6, 10% Glycerol, 1mM EDTA, 200mM KoAc, 1mM DTT,
Protease inhibitor A and B). Cells were then transferred to a 50ml Flacon tube and collected
by centrifugation. Pellets were either frozen in liquid nitrogen to be stored in –80oC or used
immediately for purification. Pellets were resuspended completely in 1ml of cold Lysis
buffer in a 2ml tube and enough glass beads were added to completely fill the tube. The cells
were lysed 5 times in a bead beater 1 minute each time, with 1 minute in between on ice. The
pellets were collected by puncturing a hole in the bottom of the 2ml tube and placing it on a
5ml syringe on a 15ml falcon tube. The pellets were then centrifuged at 10,000 rpm for 10
mins at 4oC. The supernatant was collected and was resuspended in 1ml of Lysis buffer. 25
51
µl of IgG Agarose (Sigma, equilibrated in Lysis buffer) was added to each tube and tubes
were incubated overnight at 4oC with rotation.
The following morning, beads were collected by centrifugation at 1500 rpm for 2
mins at 4oC. Cells were washed three times with 1ml of ice-cold Lysis buffer (without
protease inhibitor) and beads were collected at 1500 rpm at 4oC each time. Beads were
washed once with 1ml TEV buffer (50mM Tris Cl pH 8.0, 1mM DTT, 0.5mM EDTA) and
centrifuged again at 1500 rpm for 2 mins at 4oC. Beads were collected by removing the
supernatant as much as possible.
The beads were suspended in 100 µl of 1X SDS loading buffer and boiled for 10
mins. The samples were then loaded on a 10% SDS-PAGE gel to confirm the presence of the
TAP-tagged protein and the proteins it interacts with (which was tagged with vsv for their
detection).
52
Chapter 3
Results
53
3 Results
3.1 Epistasis relationships between genes required for
efficient cohesion and those required for histone
H3K56 acetylation
Ctf18p/Ctf8p/Dcc1p and Ctf4p have been shown to function in promoting efficient
sister chromatid cohesion [129, 136, 144, 146, 148, 161] but genetic interaction studies have
implicated these genes in other processes as well [25]. Using the LCD algorithm to analyze
E-MAP data, we found that genes encoding members of the Ctf18-complex, Ctf4p and
Asf1p/Rtt109p share similar patterns of genetic interactions with other genes mostly involved
in the maintenance of genomic integrity, i.e. DNA double-strand break repair proteins, the
MMS complex, the MRX resection complex, NHEJ proteins, and excision repair proteins
(Figures 1 and 2). This finding implies that all these genes might function together in a
common pathway.
Cells lacking Ctf18p, Ctf8p, Dcc1p or Ctf4p are sensitive to MMS [24, 161-165].
Similarly, cells lacking Asf1p or Rtt109p are highly sensitive to MMS and HU [48].
Mutation of H3K56 to an unacetylatable residue (i.e. arginine) causes hyper sensitivity to
DNA damaging agents, and, since a H3K56R mutation is epistatic with asf1Δ’s sensitivity to
DNA damaging agents, it seemed that the lack of H3K56 acetylation is the basis of the DNA
damage sensitivity of asf1Δ [166].
A genetic interaction between two genes is said to be epistatic when both the genes
work in a common pathway, such that a deletion of a second gene in addition to the first one
54
does not cause any additional growth sensitivity. If these genes all work in the same DNA
damage repair pathway, then the double deletion of ASF1 or RTT109 with CTF18 or CTF4
would not be any more sensitive to DNA damaging agents than the single mutants. In fact, I
observed that the deletion of ASF1 is moderately epistatic with the deletion of CTF18 or
CTF4, as shown in Figures 5a & 6, indicating that these genes might function in the same
pathway. The moderate additional sensitivity of asf1Δ could be due to an additional function
of Asf1p beyond H3K56 acetylation in DNA repair processes [45, 46, 48]. Likewise, deletion
of RTT109 is moderately epistatic with deletion of CTF18 or CTF4, as shown in Figures 5b
& 6. Taken together, this indicates that these genes could potentially work together in the
same DNA damage repair pathway.
To see if the cohesion promoting genes work upstream of Asf1p/Rtt109, I checked
bulk H3K56 acetylation by western blotting. Absence of Asf1p and Rtt109p caused complete
loss of acetylated H3K56 in cells. Previous studies have also shown that cells lacking ASF1
or RTT109 have elevated levels of H2A S129 phosphorylation [43]. While the global levels
of H3K56 acetylation were not affected by deletion of CTF18, CTF8, DCC1 or CTF4, cells
lacking CTF18, CTF8 or DCC1 possessed elevated levels of H2A S129 phosphorylation,
indicative of spontaneous DNA damage and/or inefficiency in repairing DNA damage
(Figure 7). Asf1p and Rtt109p also share the role in acetylating histone H3K9 with another
HAT, Gcn5p. However, the deletion of the cohesion promoting genes had no effect on H3K9
acetylation. Since this acetylation has not been linked to DNA damage repair and previous
studies indicated a common role of all these genes in a DNA damage repair pathway, the
H3K9 acetylation pathway was not further pursued.
55
Figure 5. Epistasis analysis for various mutations in the presence of 0.1% MMS. 5-fold
serial dilutions were carries out. a. Single deletion mutant strains for ASF1, CTF18 and
CTF8, in comparison with double deletions for ASF1/CTF18 and ASF1/CTF8 strains. b.
Single deletion of RTT109, CTF18 and CTF8, in comparison with double deletions for
RTT109/CTF18 and Rtt1091/CTF8 strains on MMS. Plates were incubated at 30oC for 3
days.
WT
asf1Δ
ctf18Δ
ctf8Δ
asf1Δ/ctf18Δ
asf1Δ/ctf8Δ
Glucose Glucose + 0.01% MMS
WT
rtt109Δ
ctf18Δ
ctf8Δ
rtt109Δ/ctf18Δ
rtt109Δ/ctf8Δ
b
Glucose Glucose + 0.01% MMS a
56
Figure 6. Epistasis analysis for various mutations in the presence of 0.1% MMS. 5-fold
serial dilutions of single deletion mutant strains for ASF1, RTT109. and CTF4 in comparison
with double deletions for ASF1/CTF18 and ASF1/CTF8. Plates were incubated at 30oC for 3
days.
Glucose Glucose + 0.01% MMS
WT
WT
asf1Δ
ctf4Δ
asf1Δ/ctf4Δ
rtt109Δ/ctf4Δ
ctf4Δ
asrtt109f1Δ
57
Figure 7. Effects of deleting cohesion promoting genes on histone modifications. Global
levels of histone H3K9 and H3K56 acetylation and histone H2A serine 129 phosphorylation
in rtt109∆, asf1∆, ctf4∆, ctf18∆, ctf8∆ and dcc1∆ strains analyzed on a western blot. Histone
H3 levels were used as control using α-H3 (Abcam #ab1791). Antibody used here are:
H3K9Ac (Upstate #07-352), H3K56Ac (Upstate #07-677), H2AS129 (Abcam #ab15083)
α-H3K9Ac
α-H3K56Ac
α-H2AS129Ph
α-H3
58
3.2 Effect of MMS on the association of Ctf4p and Ctf18p
with the chromatin fraction of the cell
The alternative clamp loaders play an important role in DNA replication and the
DNA damage checkpoint, as the RFC complex is intricately linked to DNA replication, and
DNA damage is inevitable during DNA replication. Among the alternative RFC complexes,
including Rad24-RFC and Elg1-RFC, Rad24p appears to play an important role in DNA
replication and the DNA damage checkpoint, while Rad24p along with Elg1p and Ctf18p
play partially redundant roles in activation of the checkpoint protein Rad53p in response to
DNA damage [162, 164, 165]. Importantly, deletion of both CTF18 and RAD24 causes
sensitivity to MMS and HU and leads to a defective intra-S checkpoint [162]. Although
Rad53p phosphorylation levels are normal when CTF18 is deleted [165], deleting RAD24,
ELG1 or CTF18 causes different extents of sensitivity to MMS and HU, while combining all
three deletions leads to a total loss of Rad53p phosphorylation. Hence, all three alternative
RFC complexes seem to play redundant roles in DNA replication and the DNA repair
checkpoint [162, 164].
This data suggests that the Cf18-complex could be recruited to DNA damage sites.
Because Ctf4p interacts with DNA pol [132], and our LCD, E-MAP and epistasis
experiments indicated that Ctf4p and Ctf18p likely function in the same pathway, it also
seemed likely that Ctf4p would be recruited to DNA damage sites. Moreover, Ctf4p and
Ctf18p were found to associate with chromatin during S phase and early M phase, consistent
with roles in cohesion and DNA replication [129].
59
Therefore, I decided to test whether Ctf18p and Ctf4p would become chromatin
associated after DNA damage. When Ogiwara et al. (2007) treated cells with MMS, a greater
fraction of the Ctf18p and Ctf4p became chromatin-associated. One possible role of
Asf1p/Rtt109p could be that it is required for the localization of the Ctf18-complex and/or
Ctf4p to the chromatin at sites of DNA damage. To rule out the possibility that the
redistribution of Ctf18p within the nucleus was an indirect effect of cells being arrested in the
G2/M phase of the cell cycle in response to DNA damage, cells were synchronized in the
original work. However, the Ctf18p preferential localization to chromatin was seen in both
synchronized and asynchronous cells, indicating the Ctf18p’s redistribution in the nucleus is
in response to DNA damage [149]. Hence, the cells were not synchronized in my work.
On performing a similar fractionation assay after using 0.1% MMS to treat strains
containing TAP-tagged Ctf18p or Ctf4p, I found that both Ctf18p and Ctf4p became
chromatin-associated, although the effect was stronger for Ctf18p than for Ctf4p (Figures 8
& 9). When the strains carried a deletion of RTT109, the absence of Rtt109p did not affect
the relocalization of either Ctf18p-TAP or Ctf4p-TAP after DNA damage (Figures 8 & 9).
This could be because Rtt109p/Asf1p works downstream of the Ctf18p complex and Ctf4p,
although this seems unlikely since deletion of Ctf4p or the Ctf18-complex does not affect
bulk histone H3K56 acetylation. Another possibility is that MMS causes many kinds of
damage, and these complexes may play a more specialized role in the repair of specific
DSB’s requiring the HR repair system [149, 150]. Finally, considering that deletion of
CTF18, CTF8, DCC1 or CTF4 does not affect global H3K56 acetylation, Rtt109p/Asf1p and
the cohesion promoting proteins may function to promote parallel aspects of DNA repair.
However, this assumption does not address the sensitivity of their double mutant to a DNA
60
Figure 8. Effect of MMS on the association of Ctf18p with the chromatin fraction of the cell.
Cells were grown in the absence (lanes 1-6) or presence (lanes 7-12) of 0.1% MMS and
fractionated for western blot assessment of the presence of Ctf18p in various cellular
fractions. The soluble fraction represents the nucleoplasmic part of the cell, the chromatin
fraction represents proteins bound to the chromatin and the WCE contains all the proteins in
the cell. WCE – Whole cell extract. Histone H3 levels are used as control. Antibodies used
here are: -TAP (made in-house), -H3 (Abcam #ab1791) and --H3K56 (Upstate #07-
677).
61
Figure 9. Effect of MMS on the association of Ctf4p with the chromatin fraction of the cell.
Cells were grown in the absence (lanes 1-6) or presence (lanes 7-12) of 0.1% MMS and
fractionated for western blot assessment of the presence of Ctf4p in various cellular fractions.
The soluble fraction represents the nucleoplasmic part of the cell, the chromatin fraction
represents proteins bound to the chromatin and the WCE contains all the proteins in the cell.
WCE – Whole cell extract. Histone H3 leves are used as control. Antibodies used here are:
-TAP (made in-house), -H3 (Abcam #ab1791) and --H3K56 (Upstate #07-677).
62
damaging agent (Figures 5 and 6) or the elevated levels of H2A S129 phosphorylation
(Figure 7). Hence, the roles of these genes were explored in another DNA damage repair
system.
3.3 Sensitivity of cells lacking cohesion promoting genes
to the presence of a single double-strand break
In response to a DSB, the MRX complex, which is comprised of Mre11p, Rad50p
and Xrs2p, arrives at the site of the break and carries out exonucleolytic cleavage of one of
the DNA strands to form single-stranded DNA. The single-stranded DNA is then recognized
by the single-stranded DNA-binding protein RPA, and checkpoint proteins are recruited in an
RPA-dependent manner. Finally, various proteins involved in HR are recruited depending on
the type of repair. Yeast YMV2 strains lacking the HML and HMR loci and having the HO
endonuclease under the control of a galactose-inducible promoter have proved useful for
identifying genes involved in the repair of DNA DSBs by SSA [58, 97]. When these strains
are grown in the presence of galactose, a single DSB is introduced at the leu2 locus where an
HO recognition site is inserted. The importance of Asf1p and Rtt109p in the process of DSB
repair was described previously using this system [97] and, in particular, Asf1p and Rtt109p
were found to be important for chromatin reassembly after the repair of the DNA DSB by
SSA.
Growth assays were performed by serially diluting cells and plating them on YP
medium containing either glucose or galactose as carbon source. Using this system the roles
of Ctf18p, Ctf8p, Dcc1p and Ctf4p were further explored. When deletions of CTF18, CTF8,
63
DCC1, CTF4, ASF1 or RTT109 were made in the YMV2 background and assayed for growth
on either sugar source, cells were sick when grown in the presence of galactose, indicating a
role for all these genes in the SSA repair pathway (Figure 10). Similarly, in the absence of
RAD52, which encodes a protein required for single-strand exchange during repair by SSA,
the cells exhibited very slow growth in the presence of galactose [58]. Using a similar
system, it has been shown that Ctf18p becomes associated with recombination intermediates
at the MAT and HML/HMR loci when mating type switching is initiated by HO cleavage,
and this association is partly dependent on Rad52p [149].
To confirm the sensitivity of asf1 , rtt109 , ctf18 , ctf8 , dcc1 and ctf4 strains to
a single DSB, deletions were made in a different strain, which uses the same SSA repair
mechanism but has the homologous regions located 25kb apart rather than 30kb apart [58].
As shown in Figure 11, the rad52Δ, asf1Δ, ctf8Δ, ctf18Δ and ctf4Δ strains again grew poorly
in the presence of galactose, confirming that, indeed, the Ctf18p complex and Ctf4p are
needed for DSB repair by SSA or possibly for checkpoint recovery.
3.4 The Ctf18p complex and Ctf4p are not required for
DSB repair by SSA
As discussed earlier, once a DSB is formed, the DNA around the site of the break is
resected by the MRX complex. For repair by SSA, the DNA is resected until the two
homologous regions become single-stranded. Because the two homologous regions are
separated by 30 kb, a lot of DNA is resected, which is thought to trigger a strong checkpoint
response. Using primers around the region of the break (P1, P2, P3, and P4, as shown in
64
Figure 10. Effects of deletions of cohesion promoting genes on the growth of a strain with a
single inducible DSB. Growth assay by 5-fold serial dilutions of deletion strains for ASF1,
RTT109, CTF18, CTF8, DCC1, CTF4 or RAD52 in the presence or absence of a single
double-strand break induced by galactose in the yeast YMV2 strain. Slow growth on
galactose indicates that the deleted gene is required for the DNA double-strand break repair
process or checkpoint recovery. Plates were incubated at 30oC for 3 days.
Ymv2
asf1∆
rtt109∆
ctf18∆
ctf8∆
dcc1∆
ctf4∆
rad52∆
Glucose ‘HO off’ Galactose ‘HO on’
65
Figure 11. Effects of deletions of cohesion promoting genes on the growth of a strain with a
single inducible DSB. Growth assay by 5-fold serial dilutions in the YMV80 strain. The
YMV80 strain has the two homologous sites separated by 25 kb (compared to 30 kb in YMV2
strain) but uses the same SSA process for repair of the DSB. Again, the growth of single
deletion mutants of RAD52, ASF1, CTF18, CTF8 or CTF4 were analyzed in the presence or
absence of a single DSB induced by galactose.
Ymv80
asf1∆
ctf18∆
ctf8∆
ctf4∆
rad52∆
Glucose ‘HO off’ Galactose ‘HO on’
66
Figure 4), the progress of repair can be monitored by PCR. Once the DSB is formed, the
amount of P1-P4 PCR product starts to decrease due to cutting by the HO endonuclease. The
region between P3-P4 represents the other homologous region, and it takes about 7-8 hours
for the resection machinery to reach this region and for the cells to repair the break [58]. The
P1-P4 PCR product only appears after repair is successfully completed.
Cells were grown in raffinose and galactose was added at time point 0, the first
sample being taken just before the addition of galactose. At subsequent intervals of about 2
hours, samples were taken to monitor DNA repair. Since galactose immediately induces the
Gal and HO genes, the break is induced within 30 mins of galactose induction. Genomic
DNA was amplified by PCR using P1-P2 and P1-P4 primers. In wild type cells, the P1-P2
(resection/cut product) PCR product decreased with time and the repair P1-P4 PCR product
started to appear by about 6-8 hours (Figure 12a).
Rad52p is a single-strand annealing protein involved in strand exchange during
homologous recombination. The absence of this protein eliminates the process of SSA [58],
although Rad52p becomes dispensable when the repair takes place on tandem repeats where
sufficiently large regions of homology compensate for the requirement of Rad52p [167]. As
expected, in rad52Δ cells the intensity of the P1-P2 (resection/cut product) decreases with
time, but no P1-P4 repair PCR product was formed (Figure 12f).
Both rtt109Δ and asf1Δ cells were shown previously to have normal repair of a DSB
by SSA [97]. Similar to rtt109Δ and asf1Δ cells (Figure 12b), I found that ctf18Δ, ctf8Δ and
ctf4Δ cells showed normal repair by SSA (Figure 12c, d, e). Taken together, the data
indicates that both the Ctf18-complex and Ctf4p are required downstream of the actual DNA
DSB repair process itself.
67
Figure 12. PCR analysis to assess the effects of mutations in cohesion promoting genes on
DSB formation and its repair. The P1-P2 primer pair amplifies the DNA spanning the
homologous sequence adjacent to the site where the DSB is induced. Hence the PCR product
decreases with time. The P1-P4 primer pair amplifies the DNA between the two distant
homologous ends, which come close only after the DNA is repaired. Hence this primer pair
only appears after the DNA is repaired (see Figure 4). a. The pattern of DSB induction and
its repair in wild type cells. b. Normal DSB induction and repair in the absence of ASF1 c.
Normal DSB induction and repair in the absence of CTF18 d. Normal DSB induction and its
repair in the absence of CTF8 e. The pattern of DSB induction and its repair in the absence of
CTF4 f. Normal DSB induction but no repair in the absence of RAD52
68
3.5 Rad53 hyperphosphorylation in the absence of
cohesion promoting genes
Emerging evidence indicates that checkpoint strength is variable depending on the
kind of DNA damage, or an inability to repair the damaged DNA [168]. In cells having intact
HML and HMR at mating type loci, where the damage caused by HO can be repaired in less
than 1 hour, the DNA damage checkpoint is not activated. In the repair system I am using,
where the break is not repaired for 7-8 hours, Rad53p is hyperphosphorylated as an index of
checkpoint activation, and this disappears once the DNA is repaired. Alternatively, cells can
also turn off the checkpoint by adaptation.
When the Rad53p becomes hyperphosphorylated, it appears as a slowly migrating
band upon SDS-PAGE when visualized using an antibody against Rad53p [58]. Following
galactose induction of a single DSB by HO, when samples of the culture were taken for
genomic DNA extraction to assess DSB formation and its repair by PCR, I simultaneously
took samples for western blotting to measure Rad53p phosphorylation (Figure 13).
As expected, wild type YMV2 cells exhibited hyperphosphorylation of Rad53p when
a DSB was induced, and this hyperphosphorylation disappeared once the DNA was repaired
by 8 hours (Figure 13a). Cells lacking ASF1 or RTT109 are known to have normal activation
of Rad53p phosphorylation, but fail to turn off the checkpoint and maintain high levels of
Rad53p hyperphosphorylation even after the repair has taken place (see Figure 13e for asf1 )
[97]. It is also known that cells lacking CTF18, CTF8 or DCC1 are capable of Rad53p
hyperphosphorylation in response to DNA damage by MMS [149, 150, 165]. When I
examined ctf8Δ, ctf18Δ and ctf4Δ strains in the YMV2 background, the cells were indeed
69
Figure 13. Western blotting to assess the effect of various mutations in cohesion promoting
genes on Rad53p hyperphosphorylation as an index of checkpoint activation and relief.
Rad53p becomes hyperphosphorylated upon checkpoint activation in response to an
unrepaired DNA DSB. Once the DNA is repaired, the Rad53p phosphorylation level drops,
leading to checkpoint relief and resumption of cell cycle progression. Using an antibody
against Rad53p (Santa Cruz #sc-6749), its hyperphosphorylated form can be observed as
slower migrating bands on SDS-PAGE. a. Rad53p phosphorylation levels for wild type cells
that repair the DNA DSB by 8 hours b. Rad53p phosphorylation levels remain high until 30
hours in the absence of CTF18 c. Rad53p phosphorylation levels remain high until 30 hours
in the absence of CTF4 d. Rad53p phosphorylation levels remain high until 24 hours in the
absence of CTF8 e. Rad53p phosphorylation levels remain high until 24 hours in the absence
of ASF1.
70
capable of normal activation of Rad53p hyperphosphorylation but, as was the case for asf1Δ
and rtt109Δ strains, hyperphosphorylation of Rad53p persisted after the DNA was repaired
(Figure 13b,c,d). This indicated that these genes are important for the checkpoint recovery
that normally occurs after the DSB is repaired.
3.6 Chromatin reassembly at the site of the DNA double-
strand break
After the repair of a DNA DSB, the repair proteins leave the site of the break and
chromatin is re-assembled on the repaired DNA. The ChIP of histone H3 can help to monitor
the progress of the resection and repair pathways by accounting for the presence or absence
of nucleosomes at the site of the break. Therefore ChIP was performed on the wild type
YMV2 strains and its derivative strains, YHS007 (asf1Δ), YHS009 (ctf18Δ), YHS010 (ctf8Δ)
and YHS012 (ctf4Δ) at various times up to 24 hours after the induction of a DSB by HO to
assess nucleosome reassembly after the repair of the DNA DSB (Figures 14 and 15). Primers
very close to the break site (less than 500 bp) were used to study nucleosome dynamics at the
site of the break.
PCR analysis of the input DNA used for ChIP monitors DNA resection and, as
expected, declined after the DSB was introduced at time 0 and recovered as the DNA was
repaired (Figures 14a and 15a). Reinforcing my previous observation that DNA cutting and
repair occur with similar efficiency in all the mutants, the input DNAs for all the mutants
(YHS007, YHS009, YHS010, YHS012) were similar to that of YMV2 (Figures 14 and 15).
Strikingly, however, all these mutants had very different H3 ChIP DNA patterns from that of
71
Figure 14. Effects of various mutations in cohesion promoting genes on nucleosome
occupancy at the DSB. Chromatin immunoprecipitation (ChIP) on histone H3 at the site of
the DNA DSB to analyze nucleosome occupancy during the process of DNA DSB formation
and its repair.The Input DNA represents samples that were not enriched with antibody. The
IP represents samples that were enriched with histone H3 antibody. The HO-cut where the
DSB is located is represented by a primer pair that was located very close to the DSB site,
while the control represents a primer pair located within the telomere, which is unaffected by
the DSB induction. a. Levels of histone H3 around the site of the DNA DSB after the
induction of a single DSB in wild type cells b. Levels of histone H3 around the site of the
DNA DSB after the induction of a single DSB in the ctf8 strain c. Levels of histone H3
around the site of the DNA DSB after the induction of a single DSB in the ctf18 strain d.
Levels of histone H3 around the site of the DNA DSB after the induction of a single DSB in
the ctf4 strain
72
Figure 15. Relative enrichment of histone H3 at the site of the DNA DSB. The levels of
histone (H3) at the site of the DSB were compared at various time points to those in another
region within the nucleus. The results from figure 14 are represented here as the ratio of
histone occupancy at the site of the DSB compared to another site which is unaffected by the
DSB. The values plotted on the graphs are the ratio normalized by the value at time 0. The
input reflects the effect of the resection and repair at the site of the DNA DSB, while the IP
represents the histone H3 occupancy at the same site. a. Normal distribution of histone H3
around the DSB site in wild type cells b. Histones are not reassembled onto the DNA after
the DNA is repaired in asf1Δ cells c. Histones are not reassembled onto the DNA after the
DNA is repaired in ctf18Δ cells d. Histones are not reassembled onto the DNA after the DNA
is repaired in ctf4Δ cells.
73
YMV2, although they were all similar to each other, such that the levels of histone H3 do not
increase after the DNA is repaired i.e. after 8-10 hours (Figures 14b,c,d and 15b,c,d).
As was previously observed for an asf1Δ strain, the nucleosomes were not re-
assembled in the absence of ASF1 (Figure 15b) [97]. Similarly, in the absence of CTF18,
CTF8 or CTF4, the level of H3 was relatively low at the site of the break even after 24 hours,
indicating that cells are incapable of efficient nucleosome reassembly in their absence. The
similarity in the patterns of genetic interactions of asf1Δ, rtt109Δ, ctf18Δ, ctf8Δ, dcc1Δ and
ctf4Δ with each other and with other genes involved in DNA repair suggests that these genes
might work together to reassemble nucleosomes onto the DNA at a DSB. There is, however,
an alternative to the possibility that Ctf4p and the Ctf18-complex directly help assemble
nucleosomes on repaired double-stranded DNA. The Ctf18-complex is an alternative RFC
complex which can load PCNA during DNA replication and possibly also during post-
replicative repair [149]. This issue is further addressed in the Discussion.
3.7 Kinetics of the appearance of Ctf18p and Ctf4p
around the site of the DSB
The results from the ChIP experiments on histone H3 indicated that both Ctf18p and
Ctf4p are required for proper nucleosome reassembly at a DSB. Therefore, it seemed likely
that they would be physically present at or near the site of the DSB. Indeed, in an earlier
study, using a strain carrying a repairable DSB which required 1 hour to repair by HR, had
shown that Ctf18p localizes to a region spanning about 10kb around the site of a DSB as
early as 2 hours after the induction of the DSB [149].
74
Ctf18p and Ctf4p were MYC-tagged to facilitate their detection and introduced into
the YMV2 strain, creating the strains YHS036 and YHS037, respectively. The association
with chromatin of Ctf18p and Ctf4p was then monitored for up to 6 hours, since the repair
itself occurs by 7 hours after the induction of the DSB. Primers were chosen to span about 20
kb on either side of the site of the DSB, with the primer pairs spaced about 2.5 kb apart.
Using anti-MYC antibody in a ChIP experiment on YHS036 and YHS037 strains, I observed
that, as was previously shown [149], Ctf18p was present around the site of the DSB by the
first time point (i.e. 2 hours)(Figure 16). The ChIP of Ctf18p-MYC also revealed two other
prominent features: firstly, in addition to Ctf18p being present around the site of the break by
2 hours, its level dropped by 6 hours (Figure 16); and secondly, the Ctf18p level was lowest
adjacent to the site of the break (Figure 17a), reminiscent of the patterns for cohesin and
γH2AX [67, 78, 127, 169].
A similar ChIP on Ctf4p in the strain YHS037 showed another very interesting
pattern around the site of the DSB (Figure 18). Unlike Ctf18p, Ctf4p was not enriched
around the site of the break by 2 hours. In fact, Ctf4p only began to appear after
about 4-6 hours, a time by which most of the Ctf18p had disappeared. However, the pattern
of occupancy of Ctf4p around the site of the break was similar to that of Ctf18p (Figure 17b).
It is possible that the decreased occupancy of cohesin, γH2A, Ctf18p and Ctf4p directly
adjacent to the site of a DSB could be simply due to resection and/or their inability to bind to
single-stranded DNA. Another possibility is that their low abundance at the site of DSB may
allow for the binding of various proteins involved in the repair process itself to facilitate
efficient repair. Indeed none of these proteins is required directly for the repair of the
damaged DNA.
75
Figure 16. Occupancy of Ctf18p at various positions around the site of the DSB at various
time points a & b. ChIP was done to assess levels of Ctf18p on either side of the break.
Control primers were designed to a region that is unaffected by DSB induction. Input
represents samples that were not enriched for Ctf18p, while IP represents enriched samples
c. Levels of Ctf18p at 0, 2, 4 and 6 hours after DSB induction in a region extending about 15
kb on either side of the DSB.
- indicates the region to the left of the DSB and + indicates region to the right of the DSB.
Primers used for this analysis are described in Table 5.
76
Figure 17. Occupancy of Ctf4p at various positions around the site of the DSB at various
time points a & b. ChIP was done to assess levels of Ctf4p on either side of the break.
Control primers were designed to a region that is unaffected by DSB induction. Input
represents samples that were not enriched for Ctf4p, while IP represents enriched samples c.
Levels of Ctf4p at 0, 2, 4 and 6 hours after DSB induction in a region extending about 15 kb
on either side of the DSB.
- indicates the region to the left of the DSB and + indicates region to the right of the DSB.
Primers used for this analysis are described in Table 5.
77
Figure 18. Relative enrichment of Ctf18p (at 2 hour) and Ctf4p (at 6 hour) for about 20kb on
either side of the DSB. The results of Figures 16 and 17 and other experiments are
represented here. The enrichment of Ctf18p or Ctf4p around the region of the break is the
ratio of DNA in the Ctf18p or Ctf4p IP to the DNA in the Input normalized to the ratio of the
control DNA. a. Ctf18p is present around the site of the DSB as early as 2 hours after DSB
induction b. Ctf4p is present around the site of the break at 6 hours after DSB induction.
78
3.8 Influence of histone H3K56 acetylation on the
recruitment of Ctf18p and Ctf4p around the site of a
DSB
Asf1p- and Rtt109p-dependent histone acetylation on H3K56 is required for
chromatin reassembly and checkpoint termination after the repair of a DNA DSB [97]. Since
cells lacking CTF18 or CTF4 have defects in both checkpoint deactivation and chromatin
reassembly, it seemed likely that these proteins could, in principle, have some role in the K56
acetylation pathway. To test this idea, ASF1 and RTT109 deletions were made in cells with
tagged Ctf18p or Ctf4p (strains YHS036 and YHS037). The absence of either ASF1 or
RTT109 did not affect the kinetics of appearance or abundance of Ctf18p around the site of
the break (Figure 19), indicating either that Asf1p- and Rtt109p-dependent histone
acetylation acts downstream of Ctf18p in chromatin reassembly or that Ctf18p and H3K56
acetylation have independent roles in chromatin assembly. However, the ChIP signal
indicating the abundance of Ctf4p was substantially decreased in the absence of ASF1 or
RTT109 (Figure 20), suggesting that H3K56 acetylation could be a mark that recruits Ctf4p
to the site of the DSB.
Since Ctf18p was present near the break site much earlier than was Ctf4p, it seemed
possible that Ctf18p might also act upstream of Ctf4p. To investigate this possibility, I
introduced a CTF18 deletion into cells with tagged Ctf4p. However, the CTF18 deletion did
not affect the occupancy of Ctf4p around the site of the break (Figure 21), consistent with
previous reports that Ctf18p is not required for the localization of Ctf4p. Therefore, it seems
79
Figure 19. The enrichment of Ctf18p around the site of the break in the absence of ASF1 or
RTT109 2 hours after DSB induction. A similar ChIP on Ctf18p, as shown in Figure 16, was
done in the absence of either ASF1 or RTT109 to see if that affects the localization of Ctf18p
around the site of the break 2 hours after DSB induction. Input is the DNA sample that was
not enriched with anti-myc antibody and represents the background levels of Ctf18p. IP is the
sample that was enriched with the antibody. The IP:Input ratio around the DSB was
normalized to the ratio of the control DNA The control bands normalize for non-specific
precipitation. a. ChIP gel on Ctf18p similar to that on Figure 16 b. ChIP gel on Ctf18p in the
absence of ASF1 c. ChIP gel on Ctf18p in the absence of RTT109 d. Levels of Ctf18p around
the site of the DSB 2 hours after DSB induction in the presence or absence of ASF1 e. Levels
of Ctf18p around the site of the DSB 2 hours after DSB induction in the presence or absence
of RTT109.
80
Figure 20. The enrichment of Ctf4p around the site of the break in the absence of ASF1 or
RTT109 6 hours after DSB induction. A similar ChIP on Ctf18p, as shown in Figure 18, was
done in the absence of either ASF1 or RTT109 to see if that affects the localization of Ctf4p
around the site of the break 6 hours after DSB induction. Input is the DNA sample that was
not enriched with anti-myc antibody and represents the background levels of Ctf4p. IP is the
sample that was enriched with the antibody. The IP:Input ratio around the DSB was
normalized to the ratio of the control DNA The control bands normalize for non-specific
precipitation. a. ChIP gel on Ctf4p similar to that on Figure 18 b. ChIP gel on Ctf4p in the
absence of ASF1 c. ChIP gel on Ctf4p in the absence of RTT109 d. Levels of Ctf4p around
the site of the DSB 6 hours after DSB induction in the presence or absence of ASF1 e. Levels
of Ctf18p around the site of the DSB 6 hours after DSB induction in the presence or absence
of RTT109.
81
Figure 21. Influence of CTF18 on the enrichment of Ctf4p around the site of the break 6
hours after DSB induction. A similar ChIP on Ctf4p as shown in Figure 18 was done in the
absence of CTF18 to see if that affects the localization of Ctf4p around the site of the break 6
hours after DSB induction. Input is the DNA sample that was not enriched with anti-myc
antibody and represents the background levels of Ctf18p. IP is the sample that was enriched
with the antibody. The IP:Input ratio around the DSB was normalized to the ratio of the
control DNA The control bands normalize for non-specific precipitation. a. ChIP gel on
Ctf4p at 6 hour similar to that in Figure 18 at 6 hour b. ChIP gel on Ctf4p at 6 hour in the
absence of CTF18 c. Levels of Ctf4p around the site of DSB at 6 hours after DSB induction
both in the presence and absence of CTF18.
82
likely that the Ctf18p-complex and the Asf1p/Rtt109p/Ctf4p influence chromatin reassembly
in distinct ways.
3.9 Physical interactions of Asf1p with Ctf18p and Ctf4p
Physical interactions are strong indicators of functional connections, and large scale
identification of yeast protein-protein interactions by affinity purification from TAP-tagged
strains followed by mass spectrometry has been carried out [31, 34]. Since our in-house
dataset has identified a physical association of Asf1p with Dcc1p, one of the subunits of the
Ctf18-complex (although only two Dcc1p peptides were identified), it seemed possible that
the Ctf18-complex and, perhaps, Ctf4p might associate with Asf1p. In view of this, I
introduced Asf1p tagged with the vsv epitope into strains containing TAP-tagged Ctf18p,
Dcc1p or Ctf4p. Using the standard two-step TAP-tag purification on IgG beads followed by
calmodulin beads, the three TAP-tagged proteins were purified but no Asf1p was detected.
The two-step purification is very stringent and discriminates against weak interactions. Since
proteins that assemble for DNA damage repair may have transient or weak interactions, I
also carried out one-step purifications, which are better suited for detecting weak
interactions. The cell lysates were incubated with beads coated with IgG to bind the protein
A component of the TAP-tag and run on a gel to test for the presence of Asf1p by western
blotting. On probing with anti-vsv antibody, a band corresponding to Asf1p was recovered
for both Ctf18p and Ctf4p purifications (Figure 22a), indicating that both these proteins
either directly or indirectly interact with Asf1p.
83
Figure 22. a. Physical interactions of Asf1p with Ctf18p and Ctf4p. Extracts derived from
strains with vsv-tagged Asf1p and TAP-tagged Ctf4p or TAP-tagged Ctf18 were precipitated
with IgG, and the input extracts and IgG precipitates were western blotted with anti-vsv
antibody. b. The interaction of Asf1p with Ctf18p and Ctf4p are not likely to be DNA-
mediated. The extracts were treated with DNAase I (25 units/ml) or EtBr (0.1 μg/ml) for 2
hours at room temperature prior to immunoprecipitation. EtBr- Ethidium bromide.
84
Since all these proteins accumulate around the site of a DSB, it seemed possible that
the interactions might be DNA-mediated. To investigate whether this might be true, ethidium
bromide (EtBr) and DNase were added to the cell extracts to reduce or eliminate DNA-
protein interactions [47]. As shown in Figure 22b, the interaction of Ctf18p or Ctf4p with
Asf1p was still observed, indicating that these interactions are not likely to be DNA-mediated
even though they are not very stable. Although I used the same amounts of EtBr and DNAse
I as were used in previous publications to efficiently destroy DNA-based interactions, I did
not have a control to check whether the DNA was properly degraded. Hence, I cannot
disregard the possibility of these interactions being DNA-mediated. Whether the interactions
of Asf1p with the Ctf18-complex and Ctf4p are direct or mediated by other proteins will also
require further research.
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Chapter 4
Discussion and Future Experiments
86
4 Summary
SGA and E-MAP datasets have facilitated the functional characterization of many
proteins. The E-MAP dataset on chromatin-related genes, which included both alleviating
and aggravating genetic interactions, assigned interaction scores based on the extent of
epistasis [21, 22]. Using the LCD algorithm [42], genes with partially overlapping patterns of
genetic interactions could be grouped, allowing for multiple predictions for a single gene.
Indeed, on applying the LCD algorithm to the E-MAP dataset on chromatin-related genes,
many new functional connections were predicted [42].
In my thesis work, I used genetic and biochemical approaches to further explore a
predicted functional relationship among Ctf18p, Ctf8p, Dcc1p, Ctf4p, Asf1p and Rtt109p
proteins. Upon galactose-induction of a single DSB in a strain that requires 7-8 hours to
repair by SSA [58], I found that neither the Ctf18-complex nor Ctf4p is required for efficient
repair of the DSB (Figure 12); however, elevated levels of Rad53p hyperphosphorylation
were maintained in the absence of these genes (Figure 13) long after the repair of the DSB
was completed. Furthermore, the Ctf18-complex and Ctf4p seemed to be needed for efficient
chromatin reassembly after the repair of the DNA DSB (Figures 14, 15 and 16). A previous
study produced similar findings in the absence of genes encoding Asf1p or Rtt109p [97],
using a similar system for DNA repair by SSA. While this indicated that all these genes are
needed for some aspect of the SSA repair process and/or checkpoint deactivation, their
functional interdependence remained unknown.
Since the proteins encoded by these genes seem to affect the DSB damage repair via
the DNA damage checkpoint, it seemed likely that these proteins would be present around
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the site of the DSB. Therefore, to further understand the role of the Ctf18-complex and Ctf4p
in the process of DSB repair along with Asf1p and Rtt109p, I examined the physical presence
of these proteins around the site of the DNA DSB. The patterns and kinetics of the
occupancy of Ctf18p and Ctf4p around the site of the DNA DSB were indeed very intriguing
(Figures 16, 17 and 18). Both proteins are found up to 15 kb away on either side of the DSB.
However, while the levels of Ctf18p peak as early as 2 hours after induction of the DSB and
decline afterwards, Ctf4p only appears about 6 hours after DSB induction, a time when the
Ctf18p levels are low. Moreover, in the absence of CTF18, Ctf4p was still present around the
site of the DSB.
Furthermore, I investigated whether the presence of Ctf18p and Ctf4p was dependent
on Asf1p or Rtt109p. I found that, while the presence of Ctf18p around the site of the DSB
was not dependent on Asf1p or Rtt109p, the presence of Ctf4p was dependent on the
presence of these proteins (Figures 19 and 20). This indicates that Ctf4p is downstream if
Asf1p/Rtt109p and, possibly, H3K56 acetylation.
Taken together these results indicate that, among these proteins, Ctf18p arrives first at
the DSB and possibly has a role independent of the roles of Ctf4p/Asf1p/Rtt109p, since its
presence is not dependent on any of these proteins. Subsequently, Asf1p and Rtt109p are
probably recruited to the site of the DSB, leading to the recruitment of Ctf4p, followed by
proper chromatin reassembly and checkpoint termination, including dephosphorylation of
Rad53p.
The poor growth caused by CTF18, CTF8, DCC1 and CTF4 deletions in YMV2
strains containing an inducible HO endonuclease suggested a possible immediate role of the
Ctf18-complex and Ctf4p in the DSB repair pathway. Ctf18p assembles into an alternative
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RFC complex, which loads PCNA during DNA replication, and Ctf4p interacts with DNA
polymerase. Assuming that there is no role for DNA replication, DNA polymerase, or PCNA
during resection or single-strand annealing (Figure 23), I found that cells were able to
efficiently repair the DNA DSB in the absence of CTF18 or CTF4, indicating that they play
different roles in this process. Similarly, Ctf4p was also not required for the repair process in
my experiments, indicating that its interaction with the replication machinery is also not
involved in resection or annealing.
During the SSA process, resection occurs on both ends of the DSB, as shown in
Figure 23, Step 3. Once the two homologous ends are annealed, there is relatively less DNA
to be made double-stranded at one end, compared to the other (Figure 23, Step 4). Since both
resection and DNA replication occur in the 5’ – 3’ direction, DNA repair analysis by using
primers around the site of the DSB might not be the best indication of the completion of
DNA repair, because the PCR product between P1 and P4 will still be formed even if the
DNA is still partially single-stranded. Hence, it is possible that the Ctf18-complex and/or
Ctf4p are required for filling in the single-stranded gaps that remains after annealing of the
homologous sequences, by interacting with the replication machinery.
Based on the results in this thesis, and the above described caveat in the experimental
design, there are three possible but closely related models for the roles of these genes during
the process of DNA DSB repair. Firstly, assuming that Ctf18p and Ctf4p are indeed required
for filling in the single-stranded gaps left after the HR process (Figure 23, Step 4), Ctf18p
could be required for converting the single-stranded DNA to double-stranded DNA by
loading the PCNA for DNA replication (Figure 23 Step 5); Ctf4p could be required to
interact with DNA polymerase for proper DNA replication to fill the ssDNA gaps and
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Figure 23. Repair of a DNA DSB by single strand annealing. P1 and P4 represent primers
that were used for DNA repair analysis. Both resection and DNA replication take place in a
5’ to 3’ direction. Hence, some single-stranded DNA may, still be present when PCR
indicates that the DNA has been repaired.
Single strand degradation and
beginning of repair DNA synthesis
Completion of repair
and DNA synthesis
Resected DNA
Replicated DNA To-be degraded DNA to be
degraded
Formation of DSB
Resection
Single-strand annealing
90
Asf1p/Rtt109p could modulate this activity by either acetylating Ctf4p or another target
around the DSB. Secondly, it is possible that the Ctf18-complex and Ctf4p are not required
for filling in the single-stranded gaps, but rather for the process of chromatin reassembly
accompanying Asf1p/Rtt109p-dependent H3K56 acetylation. Finally, it is possible that
Ctf18p, which arrives first at the site of the DSB, is required for filling in the ssDNA gaps,
whereas Ctf4p is required for chromatin reassembly, or chromatin reassembly coupled to
DNA replication, along with Asf1p/Rtt109p, as shown in Figure 24. Future experiments will
help to determine whether single-stranded region remains after the DSB repair in the absence
of Ctf18p or Ctf4p. If true, then those single-stranded regions could be responsible for the
failure to reassemble chromatin or deactivate the checkpoint in YMV2 strains.
4.1 Cellular response to DNA damage
Genomes are frequently damaged and must be efficiently and accurately repaired to
maintain their integrity. Many of these challenges come from environmental stresses, but
endogenous events also cause DNA damage during normal cell cycle progression. These
insults can lead to base damage, single-strand DNA breaks or double-strand DNA breaks
[170]. DSBs are particularly deleterious, as their inefficient or inaccurate repair can cause
mutations or chromosomal translocations, both of which are hallmarks of cancer.
A DSB can be repaired by either homologous recombination, which uses homologous
DNA sequences as templates for repair, or non-homologous end joining, which involves re-
ligation of the broken ends. The overall DSB repair process, however, involves much more
than simply the rejoining/repair of the broken regions of the chromosome. DSBs also result
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Figure 24. Model describing the possible role of Ctf18-complex and Ctf4 during the process
of the DNA DSB repair.
92
in the activation of a complex cascade of DNA-damage responses: sensing the DNA damage;
amplification and transmission of the damage signal; and activation of cell-cycle checkpoint
kinases, which results in cell cycle arrest until the DNA is restored to its original form.
After a DSB is formed and the damage site is detected, there is an amplification of the
damage signal through a cascade of protein kinases, leading to activation of a series of
downstream effectors that promote checkpoint activation and cell cycle arrest. The main
protein kinases in S.cerevisiae for proper checkpoint activation following a DSB are Mec1p,
Tel1p, Rad53p and Chk1p. Rad53p plays a pivotal role in the DNA damage checkpoint and
controls the majority of the DNA damage responses. Inactivation of the DNA damage
checkpoint so that the cell cycle can resume occurs in two situations: by recovery, after the
DNA lesions are repaired, or by adaptation, when some unrepaired DNA lesions persist.
Rad53p inactivation by its dephosphorylation is a key event for both recovery and adaptation,
because both are accompanied by the disappearance of Rad53p phosphorylation [58, 64, 76].
Hence, all adaptation- and recovery-defective mutants exhibit persistent phosphorylation of
Rad53p [58, 63, 64, 76]. These adaptation-defective mutants include the Ku family of genes,
Cdc4p and Srs2p [58, 97].
Previous studies have linked Asf1p, Rtt109p, Ctf18p, Ctf8p, Dcc1p and Ctf4p to
DNA repair via HR [97, 149, 150]. While histone H3K56 acetylation by Rtt109p and Asf1p
is required for chromatin reassembly after a DNA DSB is repaired [97], the roles of Ctf18p,
Ctf8p, Dcc1p and Ctf4p in this process have not been clear. Since all the genes encoding
these proteins share similar patterns of genetic interactions with other genes involved in
DNA repair (Figure 2), it is likely that the Ctf18-complex and Ctf4p are involved in the
Asf1p/Rtt109p pathway as well.
93
Asf1p and Rtt109p are necessary for adaptation in the presence of a single DSB that
can be repaired by SSA [97]. While the DSB is repaired efficiently by SSA in the absence of
ASF1 and RTT109, nucleosomes fail to reassemble onto the repaired DNA. Also, these cells
are also unable to turn off the checkpoint, indicating that chromatin reassembly is important
for Rad53p inactivation [97]. Similarly, cells lacking CTF18, CTF8 or CTF4 were
adaptation-defective and displayed activated checkpoints for prolonged periods of time
(Figure 13). While Asf1p becomes necessary for survival in the presence of a single DSB
that can only be repaired by SSA, it is not required when repair can occur by GC, where the
homologous region is present on another chromosome. While SSA and GC activate the
checkpoint in somewhat similar manners, it is possible that the type of repair by HR plays an
important role in some cellular responses. In particular, some of the repair proteins that are
recruited to the site of the DSB may be specific to both the type of break and the subsequent
repair process [51]. Repair by SSA is unique in that it leads to a complete loss of DNA
between the two homologous regions, and hence could be especially tightly controlled. I
observed that both Ctf18p and Ctf4p are recruited to the site of a DSB when repair occurs by
SSA, but this need not necessarily be true when repair occurs by GC.
Cells lacking either CTF18 or CTF4 are sensitive to growth in the presence of MMS
[149, 150, 165]. CTF4 and CTF18 deletions display genetic interactions with mutations in
DNA replication genes, spindle assembly checkpoint genes and genes involved in sister
chromatid cohesion, microtubule function and chromosome structure [24, 129, 146, 161].
The Ctf18-complex also appears to have a role in DNA damage checkpoint signaling.
CTF18, CTF8 and DCC1 deletion mutants have synthetic sick interactions with genes
encoding the DNA damage checkpoint proteins Rad9p, Ddc1p, Mec3p, Rad17p and Rad24p,
94
indicating that Ctf18p, Ctf8p and Dcc1p function in a pathway parallel to the Rad24-
dependent DNA damage checkpoint [25, 165]. My initial assessment of the effects of MMS
on the growth of ASF1, RTT109, CTF18, CTF8 and CTF4 deletion strains suggested a
functional interplay among these genes during DNA repair. Strains with a deletion of ASF1
or RTT109 combined with a deletion of CTF18, CTF8 or CTF4 were only slightly sicker than
the sickest single deletion strain, indicating that these genes could work in the same pathway
(Figures 5 and 6). Previous studies had shown that ctf18Δ cells are sensitive to MMS, but
only partially sensitive to HU, which interferes with the replication fork, because the
presence of a redundant protein complex containing Rad24p ensures faithful replication
[165]. Therefore, it is likely that the Ctf18-complex has more than one role in the repair of
DNA damage, and the sensitivity to MMS caused by deletion of CTF18 may reflect one of
its many functions.
As important as the acetylation of histone H3K56 is in the DNA damage response, its
deacetylation by Hst3p and Hst4p is equally important, as hst3Δ hst4Δ cells accumulate
spontaneous DNA damage [101, 171]. Cells lacking HST3 and HST4 show a plethora of
chromatin-associated phenotypes, which result from hyperacetylation of H3K56 since
mutation of K56 to an unacetylable arginine residue suppresses nearly all of the hst3Δ hst4Δ
phenotypes [99, 100].
Interestingly, deleting either CTF18 or CTF4 suppressed the hst3Δ hst4Δ MMS
sensitivity phenotype [159]. A CTF4 deletion has a positive genetic interaction with H3K56R
(where the lysine was mutated to arginine) in the presence of HU, suggesting that it works
directly in the H3K56 acetylation pathway during S phase [159]. Furthermore, deletion of
CTF4 in addition to HST3 and HST4 did not affect the levels of hyperacetylated H3K56 in
95
the cell, putting Ctf4p downstream of H3K56 acetylation, which is consistent with my
observation that Asf1p is needed for the assembly of Ctf4p at a DSB. Hence, it appears that
Ctf4p works downstream of DSB repair and is somehow required for chromatin reassembly
after the DSB repair by functioning in the H3K56 acetylation pathway.
Although ctf18Δ, like ctf4∆, suppressed hst3Δ hst4Δ MMS sensitivity, it had a
negative genetic interaction with H3K56R, indicating that it is likely working through a
separate pathway [159]. The different sensitivities for Ctf18p and Ctf4p could be caused by
Hst3p and Hst4p deacetylating other protein(s) involved in sister chromatid cohesion, since
hst3Δ hst4Δ cells have SCC defects as well [101].
4.2 Chromatin reassembly around the site of a DNA DSB
Recent advances in cell-biological approaches have led to a greater appreciation of
the spatial and temporal organization of the DNA-repair machinery [172-174]. In particular,
the organization of chromatin is important for assembling the repair machinery and making
the DNA lesion accessible to the repair complex for efficient repair [175]. Moreover,
chromatin must be properly reassembled after repair has taken place. Hence, studying the
DNA repair process in the context of chromatin will help achieve a better understanding of
the events that occur during the repair of DNA damage.
When a cell encounters a DSB, chromatin remodeling complexes like INO80, SWR1,
SWI/SNF and RSC are recruited to the DSB to reconfigure the nucleosomes around the DSB
to facilitate DNA repair and/or to modulate checkpoint activation [156, 176-179]. After a
DSB is formed, histone H2A gets phosphorylated in the vicinity of the break, leading to
96
recruitment of Ino80p, Arp5p and Arp8p to the DSB, which are necessary for subsequent
processing of both ends of the DSB into single-stranded DNA [156, 179]. Additionally, this
phosphorylation leads to the recruitment of three HATs: Gcn5p, Esa1p and Rtt109p (along
with Asf1p), to the region proximal to the DSB, resulting in localized histone acetylation
[93]. While the subsequent presence of cohesin around the site of the DSB is dependent on
γH2AX [127, 180, 181], deletion of ASF1 does not affect the loss of phosphorylated H2A
S129 from the vicinity of the break. The similarity of the genetic interaction patterns of the
genes encoding Asf1p, Rtt109p, Ctf18p, Ctf8p, Dcc1p and Ctf4p makes it unlikely that these
genes would differentially affect the SSA pathway. Consistent with this hypothesis, Ctf18p is
not required for the γH2AX-dependent de novo assembly of cohesin around a DSB [149,
169].
Some clarification of the roles of the Ctf18-complex and Ctf4p came from the ChIP
experiments on histone H3 as a proxy for nucleosome reassembly after the repair of the DSB.
Cells lacking ASF1 (or RTT109) have a defect in nucleosome reassembly [97], possibly
because histone H3K56 acetylation is a mark for nucleosome deposition and, in its absence,
the nucleosomes cannot be reassembled. Similarly, ChIP around the site of the break
indicated that, following religation of broken ends of the homologous DNA, the absence of
Ctf18p, Ctf4p, Rtt109p or Asf1p causes cells to continue to display Rad53p
hyperphosphorylation (Figure 13) and the nucleosomes were not re-assembled around the
repair site (Figures 14 and 15). This inefficient nucleosome reassembly in the absence of
Ctf18p or Ctf4p could be a consequence of either a direct defect in nucleosome assembly or a
failure to convert single-stranded DNA to double-stranded DNA after single strand
annealing.
97
Proteins that are required for the repair process are usually present around the site of
the break. Hence, using ChIP, I found that both Ctf18p (and hence the Ctf18-complex) and
Ctf4p were recruited to a region spanning about 15 kb on either side of the DSB (Figures 16,
17 and 18). Interestingly, while the patterns of their occupancy were similar, they were
physically present at the site of the break at different times. Ctf18p was present as early as 2
hours after the induction of a DSB (Figure 16), whereas Ctf4p was present only 6 hours after
the induction of a DSB (Figure 17), when the repair is nearly complete and nucleosomes are
being reassembled. Additionally, Ctf18p protein levels declined around the site of the break
prior to the assembly of Ctf4p. Since there was no decline in Ctf4p levels around the DSB in
the absence of CTF18 (Figure 21), it seems likely that Ctf18p and Ctf4p work on separate
processes (albeit in the same pathway) to facilitate chromatin reassembly. This result is
consistent with previous work showing that Ctf18p is not required for the recruitment of
Ctf4p [148]. Nevertheless, I found that Asf1p interacts physically with both Ctf18p and
Ctf4p (Figure 22), which suggests that Asf1p may independently chaperone Ctf4p and
Ctf18p for their separate functions. Indeed, I found that Asf1p is required for the recruitment
of Ctf4p around the site of the break but not for Ctf18p.
A previous study indicated the presence of electrophoretic mobility variants of Ctf18p
and Ctf4p [129]. It is, therefore, also possible that Ctf18p and Ctf4p are post-translationally
modified, possibly through their acetylation by Rtt109p in conjunction with Asf1p. Asf1p
also interacts with Rad53p, an interaction which is abolished in the presence of DNA damage
[45], making it possible that the Ctf18-complex and/or Ctf4p control not only nucleosome
reassembly but also an interaction of Asf1p with Rad53p.
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Hence, although this thesis sheds light on the roles of the Ctf18-complex and Ctf4p
after DNA repair in the SSA pathway, their precise roles and relationship to Asf1p/Rtt109p
and H3K56 acetylation remains to be determined.
4.3 Future experiments
I have shown that the Ctf18-complex and Ctf4p appear to play a role during the
process of DNA DSB repair by SSA. My results indicate that these proteins are required for
an event, which leads to proper chromatin reassembly and checkpoint exit. Further work will
be necessary to address the many unanswered questions as to where these proteins work in
the DSB repair pathway.
4.3.1 What are the defects in DNA DSB repair in the absence of
CTF18 and CTF4: DNA replication or chromatin reassembly?
I have provided evidence that the absence of CTF18 and CTF4 leads to efficient re-
ligation of the homologous regions but inefficient reassembly of the nucleosomes onto the
DNA and a defect in checkpoint exit. This indicates that the Ctf18-complex and Ctf4p may
either be required directly for chromatin reassembly via the histone chaperone, Asf1p, and
the HAT, Rtt109p, or have an indirect effect on checkpoint exit and chromatin reassembly
because of a defect in the completion of DNA replication that occurs after the re-ligation of
the homologous regions, which would be needed for chromatin reassembly.
After a DSB is formed, the DNA around the site of the break it resected to expose the
two homologous regions for HR (during SSA). After re-ligation, the resected single-stranded
99
DNA has to be replicated back to double-stranded form. The experimental system I
employed, using PCR at the site of the DSB to monitor DNA repair after DSB formation,
analyses only the re-ligation event. Hence, the DNA repair that I have observed in my
experiments is, in fact, only a part of the repair process.
If replication is indeed being affected in the absence of CTF18 and CTF4, then there
will be a persistent region of single-stranded DNA flanking the DSB. Dot- or Slot-Blotting, a
form of nucleic acid hybridization, can assess the single-stranded nature of the DNA around
the site of the break. By hybridizing complementary single-stranded probes separately, along
the arms of the break, which has an extended resected region of about 30 kb, the nature of
DNA in that region can be determined. If the region is single-stranded, it will bind to only
one of the complementary probes, giving support to the re-replication model. The advantage
of this experiment is that the DNA can be assessed in its native form. Furthermore, a wild-
type strain and aRAD52 deletion mutant can be used as controls. The DNA from the wild
type strain should bind to both the complementary probes. On the other hand, DNA from the
the RAD52 deletion strain should bind to only one of the complementary probes, because the
HR event is compromised while the resection occurs efficiently in the absence of RAD52,
and so the DNA remains single-stranded.
4.3.2 Where does Ctf18p function in this pathway?
Ctf18p is not needed for re-ligation of homologous regions and it does not depend on
Asf1p/Rtt109p for its presence around the site of the DSB. If Ctf18p is not required for
filling in the single-stranded gaps, then it is conceivable that it might be required for
100
recruiting the histone chaperones, since Ctf18p is present very early (2 hours after DSB
induction) around the site of the DSB. Further support for this notion is provided by the
physical interaction between Asf1p and Ctf18p (Figure 22). While, in theory, their
interdependence could be easily addressed by chromatin immunoprecipitation on Asf1p and
Rtt109p, many studies in the past have failed to ChIP either Asf1p or Rtt109p, possibly
because their high abundance in the cell leads to a very high background.
If Ctf18p is required for recruiting Asf1p and Rtt109p, then the levels of histone
acetylation (H3K56) at the site of the DSB should be low in the absence of Ctf18p. As well,
we may be able to eliminate the requirement for Ctf18p by changing the lysine residue on
histone H3K56 to glutamine (which will mimic an acetylated state). Hence, by ChIP on
histone H3K56, the role of Ctf18p can be further addressed.
4.3.3 Why does the presence of Ctf4p depend on Asf1p/Rtt109p?
As was the case for Ctf18p, Ctf4p was not required for re-ligation of the homologous
regions, but its presence around the site of the DSB is dependent on Asf1p and Rtt109p.
Studies in the past have suggested a downstream function for Ctf4p in chaperoning the
histones at the damage site, possibly via its interaction with H3K56 acetylated nucleosomes
[159]. In accordance with that possibility and results shown in my thesis, it appears likely
that Ctf4p functions as a part of the H3K56 acetylation pathway [22].
If Ctf4p must interact with Asf1p/Rtt109p to facilitate nucleosome deposition, then
we might be able to eliminate the requirement for Ctf4p by changing the lysine residue on
histone H3K56 to glutamine (which will mimic an acetylated state) or by deleting
101
HST3/HST4 (which will cause H3K56 hyper-acetylation). This will give a clearer picture of
the role of Ctf4p in the H3K56 acetylation pathway. However, as pointed our earlier, it
remains to be determined if there is efficient DNA replication after re-ligation in the absence
of Ctf4p.
4.3.4 Is there a role for other histone chaperones in DSB repair?
While Asf1p and Rtt109p are required for acetylating histone H3 before it gets
deposited at the DNA repair site, they are not necessary for the deposition of the acetylated
nucleosome: when the lysine residue on histone H3K56 is mutated to glutamate to mimic the
acetylated form, both Asf1p and Rtt109p are dispensable for the process of DNA DSB repair
[97]. Hence, it appears that another chaperone(s) is required for deposition of the histones
onto the DNA. Another bicluster consisting of the Hir-complex, the Ctf18-complex, Asf1p
and Rtt109p was also predicted by the LCD algorithm. The Hir-complex is a replication-
independent histone chaperone which functions together with Asf1p and Rtt109p for
deposition of acetylated histones H3/H4 onto DNA [47, 182]. Another histone chaperone,
Rtt106p, also functions together with the Hir-complex and Asf1p at histone promoters [26].
My preliminary work on Hir1p and Rtt106p suggests a role for these chaperones in BSB
repair, since their deletions, like that of Asf1p, were lethal to cells carrying a single DSB.
These proteins will presumably also be required for chromatin reassembly after the repair of
a single DSB. By carrying out a series of experiments similar to those described in this thesis,
the roles of these proteins can be addressed.
102
Chapter 5
References
103
1. Luger, K., Structure and dynamic behavior of nucleosomes. Curr Opin Genet Dev,
2003. 13(2): p. 127-35.
2. Akey, C.W. and K. Luger, Histone chaperones and nucleosome assembly. Curr Opin
Struct Biol, 2003. 13(1): p. 6-14.
3. Wu, J. and M. Grunstein, 25 years after the nucleosome model: chromatin
modifications. Trends Biochem Sci, 2000. 25(12): p. 619-23.
4. Donaldson, A.D., Shaping time: chromatin structure and the DNA replication
programme. Trends Genet, 2005. 21(8): p. 444-9.
5. Mellor, J., The dynamics of chromatin remodeling at promoters. Mol Cell, 2005.
19(2): p. 147-57.
6. Peterson, C.L. and J. Cote, Cellular machineries for chromosomal DNA repair.
Genes Dev, 2004. 18(6): p. 602-16.
7. Tsukuda, T., et al., Chromatin remodelling at a DNA double-strand break site in
Saccharomyces cerevisiae. Nature, 2005. 438(7066): p. 379-83.
8. Graff, J. and I.M. Mansuy, Epigenetic dysregulation in cognitive disorders. Eur J
Neurosci, 2009. 30(1): p. 1-8.
9. Sebova, K. and I. Fridrichova, Epigenetic tools in potential anticancer therapy.
Anticancer Drugs. 21(6): p. 565-77.
10. Botstein, D. and G.R. Fink, Yeast: an experimental organism for modern biology.
Science, 1988. 240(4858): p. 1439-43.
11. Kataoka, T., et al., Functional homology of mammalian and yeast RAS genes. Cell,
1985. 40(1): p. 19-26.
12. Sinclair, D., K. Mills, and L. Guarente, Aging in Saccharomyces cerevisiae. Annu
Rev Microbiol, 1998. 52: p. 533-60.
13. Sinclair, D.A., K. Mills, and L. Guarente, Accelerated aging and nucleolar
fragmentation in yeast sgs1 mutants. Science, 1997. 277(5330): p. 1313-6.
14. Sinclair, D.A., K. Mills, and L. Guarente, Molecular mechanisms of yeast aging.
Trends Biochem Sci, 1998. 23(4): p. 131-4.
15. Strand, M., et al., Destabilization of tracts of simple repetitive DNA in yeast by
mutations affecting DNA mismatch repair. Nature, 1993. 365(6443): p. 274-6.
16. Friedberg, E.C., Eukaryotic DNA repair: glimpses through the yeast Saccharomyces
cerevisiae. Bioessays, 1991. 13(6): p. 295-302.
104
17. Guarente, L., Synthetic enhancement in gene interaction: a genetic tool come of age.
Trends Genet, 1993. 9(10): p. 362-6.
18. Novick, P., B.C. Osmond, and D. Botstein, Suppressors of yeast actin mutations.
Genetics, 1989. 121(4): p. 659-74.
19. Schuldiner, M., et al., Exploration of the function and organization of the yeast early
secretory pathway through an epistatic miniarray profile. Cell, 2005. 123(3): p. 507-
19.
20. Tong, A.H., et al., Systematic genetic analysis with ordered arrays of yeast deletion
mutants. Science, 2001. 294(5550): p. 2364-8.
21. Collins, S.R., et al., A strategy for extracting and analyzing large-scale quantitative
epistatic interaction data. Genome Biol, 2006. 7(7): p. R63.
22. Collins, S.R., et al., Functional dissection of protein complexes involved in yeast
chromosome biology using a genetic interaction map. Nature, 2007. 446(7137): p.
806-10.
23. Krogan, N.J., et al., Regulation of chromosome stability by the histone H2A variant
Htz1, the Swr1 chromatin remodeling complex, and the histone acetyltransferase
NuA4. Proc Natl Acad Sci U S A, 2004. 101(37): p. 13513-8.
24. Tong, A.H., et al., Global mapping of the yeast genetic interaction network. Science,
2004. 303(5659): p. 808-13.
25. Pan, X., et al., A DNA integrity network in the yeast Saccharomyces cerevisiae. Cell,
2006. 124(5): p. 1069-81.
26. Kainth, P., et al., Comprehensive genetic analysis of transcription factor pathways
using a dual reporter gene system in budding yeast. Methods, 2009. 48(3): p. 258-64.
27. Bender, A. and J.R. Pringle, Use of a screen for synthetic lethal and multicopy
suppressee mutants to identify two new genes involved in morphogenesis in
Saccharomyces cerevisiae. Mol Cell Biol, 1991. 11(3): p. 1295-305.
28. Wang, T. and A. Bretscher, Mutations synthetically lethal with tpm1delta lie in genes
involved in morphogenesis. Genetics, 1997. 147(4): p. 1595-607.
29. Costanzo, M., et al., The genetic landscape of a cell. Science. 327(5964): p. 425-31.
30. Ooi, S.L., et al., Global synthetic-lethality analysis and yeast functional profiling.
Trends Genet, 2006. 22(1): p. 56-63.
31. Krogan, N.J., et al., Global landscape of protein complexes in the yeast
Saccharomyces cerevisiae. Nature, 2006. 440(7084): p. 637-43.
105
32. Gavin, A.C., et al., Functional organization of the yeast proteome by systematic
analysis of protein complexes. Nature, 2002. 415(6868): p. 141-7.
33. Ho, Y., et al., Systematic identification of protein complexes in Saccharomyces
cerevisiae by mass spectrometry. Nature, 2002. 415(6868): p. 180-3.
34. Gavin, A.C., et al., Proteome survey reveals modularity of the yeast cell machinery.
Nature, 2006. 440(7084): p. 631-6.
35. Bandyopadhyay, S., et al., Functional maps of protein complexes from quantitative
genetic interaction data. PLoS Comput Biol, 2008. 4(4): p. e1000065.
36. Ulitsky, I., et al., From E-MAPs to module maps: dissecting quantitative genetic
interactions using physical interactions. Mol Syst Biol, 2008. 4: p. 209.
37. Schuldiner, M., et al., Quantitative genetic analysis in Saccharomyces cerevisiae
using epistatic miniarray profiles (E-MAPs) and its application to chromatin
functions. Methods, 2006. 40(4): p. 344-52.
38. Typas, A., et al., High-throughput, quantitative analyses of genetic interactions in E.
coli. Nat Methods, 2008. 5(9): p. 781-7.
39. Roguev, A., et al., Conservation and rewiring of functional modules revealed by an
epistasis map in fission yeast. Science, 2008. 322(5900): p. 405-10.
40. Butland, G., et al., eSGA: E. coli synthetic genetic array analysis. Nat Methods, 2008.
5(9): p. 789-95.
41. Dixon, S.J., et al., Significant conservation of synthetic lethal genetic interaction
networks between distantly related eukaryotes. Proc Natl Acad Sci U S A, 2008.
105(43): p. 16653-8.
42. Pu, S., et al., Local coherence in genetic interaction patterns reveals prevalent
functional versatility. Bioinformatics, 2008. 24(20): p. 2376-83.
43. Fillingham, J., et al., Chaperone control of the activity and specificity of the histone
H3 acetyltransferase Rtt109. Mol Cell Biol, 2008. 28(13): p. 4342-53.
44. Driscoll, R., A. Hudson, and S.P. Jackson, Yeast Rtt109 promotes genome stability by
acetylating histone H3 on lysine 56. Science, 2007. 315(5812): p. 649-52.
45. Emili, A., et al., Dynamic interaction of DNA damage checkpoint protein Rad53 with
chromatin assembly factor Asf1. Mol Cell, 2001. 7(1): p. 13-20.
46. Franco, A.A., et al., Histone deposition protein Asf1 maintains DNA replisome
integrity and interacts with replication factor C. Genes Dev, 2005. 19(11): p. 1365-
75.
106
47. Green, E.M., et al., Replication-independent histone deposition by the HIR complex
and Asf1. Curr Biol, 2005. 15(22): p. 2044-9.
48. Tyler, J.K., et al., The RCAF complex mediates chromatin assembly during DNA
replication and repair. Nature, 1999. 402(6761): p. 555-60.
49. Longhese, M.P., et al., DNA damage checkpoint in budding yeast. Embo J, 1998.
17(19): p. 5525-8.
50. Paulovich, A.G., D.P. Toczyski, and L.H. Hartwell, When checkpoints fail. Cell,
1997. 88(3): p. 315-21.
51. Jain, S., et al., A recombination execution checkpoint regulates the choice of
homologous recombination pathway during DNA double-strand break repair. Genes
Dev, 2009. 23(3): p. 291-303.
52. Krogh, B.O. and L.S. Symington, Recombination proteins in yeast. Annu Rev Genet,
2004. 38: p. 233-71.
53. Paques, F. and J.E. Haber, Multiple pathways of recombination induced by double-
strand breaks in Saccharomyces cerevisiae. Microbiol Mol Biol Rev, 1999. 63(2): p.
349-404.
54. Kellis, M., B.W. Birren, and E.S. Lander, Proof and evolutionary analysis of ancient
genome duplication in the yeast Saccharomyces cerevisiae. Nature, 2004. 428(6983):
p. 617-24.
55. Fishman-Lobell, J., N. Rudin, and J.E. Haber, Two alternative pathways of double-
strand break repair that are kinetically separable and independently modulated. Mol
Cell Biol, 1992. 12(3): p. 1292-303.
56. Sugawara, N. and J.E. Haber, Characterization of double-strand break-induced
recombination: homology requirements and single-stranded DNA formation. Mol
Cell Biol, 1992. 12(2): p. 563-75.
57. Sugawara, N., et al., Role of Saccharomyces cerevisiae Msh2 and Msh3 repair
proteins in double-strand break-induced recombination. Proc Natl Acad Sci U S A,
1997. 94(17): p. 9214-9.
58. Vaze, M.B., et al., Recovery from checkpoint-mediated arrest after repair of a
double-strand break requires Srs2 helicase. Mol Cell, 2002. 10(2): p. 373-85.
59. Sugawara, N., G. Ira, and J.E. Haber, DNA length dependence of the single-strand
annealing pathway and the role of Saccharomyces cerevisiae RAD59 in double-
strand break repair. Mol Cell Biol, 2000. 20(14): p. 5300-9.
60. Lee, S.E., et al., Saccharomyces Ku70, mre11/rad50 and RPA proteins regulate
adaptation to G2/M arrest after DNA damage. Cell, 1998. 94(3): p. 399-409.
107
61. Lee, S.E., et al., The Saccharomyces recombination protein Tid1p is required for
adaptation from G2/M arrest induced by a double-strand break. Curr Biol, 2001.
11(13): p. 1053-7.
62. Toczyski, D.P., D.J. Galgoczy, and L.H. Hartwell, CDC5 and CKII control
adaptation to the yeast DNA damage checkpoint. Cell, 1997. 90(6): p. 1097-106.
63. Lee, S.E., et al., Yeast Rad52 and Rad51 recombination proteins define a second
pathway of DNA damage assessment in response to a single double-strand break.
Mol Cell Biol, 2003. 23(23): p. 8913-23.
64. Pellicioli, A., et al., Regulation of Saccharomyces Rad53 checkpoint kinase during
adaptation from DNA damage-induced G2/M arrest. Mol Cell, 2001. 7(2): p. 293-
300.
65. Sanchez, Y., et al., Control of the DNA damage checkpoint by chk1 and rad53 protein
kinases through distinct mechanisms. Science, 1999. 286(5442): p. 1166-71.
66. Sanchez, Y., et al., Regulation of RAD53 by the ATM-like kinases MEC1 and TEL1 in
yeast cell cycle checkpoint pathways. Science, 1996. 271(5247): p. 357-60.
67. Shroff, R., et al., Distribution and dynamics of chromatin modification induced by a
defined DNA double-strand break. Curr Biol, 2004. 14(19): p. 1703-11.
68. Gilbert, C.S., C.M. Green, and N.F. Lowndes, Budding yeast Rad9 is an ATP-
dependent Rad53 activating machine. Mol Cell, 2001. 8(1): p. 129-36.
69. de la Torre-Ruiz, M.A., C.M. Green, and N.F. Lowndes, RAD9 and RAD24 define
two additive, interacting branches of the DNA damage checkpoint pathway in
budding yeast normally required for Rad53 modification and activation. Embo J,
1998. 17(9): p. 2687-98.
70. Elledge, S.J., Cell cycle checkpoints: preventing an identity crisis. Science, 1996.
274(5293): p. 1664-72.
71. Weinert, T., DNA damage and checkpoint pathways: molecular anatomy and
interactions with repair. Cell, 1998. 94(5): p. 555-8.
72. Weinert, T.A. and L.H. Hartwell, The RAD9 gene controls the cell cycle response to
DNA damage in Saccharomyces cerevisiae. Science, 1988. 241(4863): p. 317-22.
73. Zhou, B.B. and S.J. Elledge, The DNA damage response: putting checkpoints in
perspective. Nature, 2000. 408(6811): p. 433-9.
74. Tercero, J.A., M.P. Longhese, and J.F. Diffley, A central role for DNA replication
forks in checkpoint activation and response. Mol Cell, 2003. 11(5): p. 1323-36.
108
75. O'Neill, B.M., et al., 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, 2007. 104(22): p. 9290-5.
76. Leroy, C., et al., PP2C phosphatases Ptc2 and Ptc3 are required for DNA checkpoint
inactivation after a double-strand break. Mol Cell, 2003. 11(3): p. 827-35.
77. Guillemain, G., et al., Mechanisms of checkpoint kinase Rad53 inactivation after a
double-strand break in Saccharomyces cerevisiae. Mol Cell Biol, 2007. 27(9): p.
3378-89.
78. Keogh, M.C., et al., A phosphatase complex that dephosphorylates gammaH2AX
regulates DNA damage checkpoint recovery. Nature, 2006. 439(7075): p. 497-501.
79. Ivanov, E.L., et al., Mutations in XRS2 and RAD50 delay but do not prevent mating-
type switching in Saccharomyces cerevisiae. Mol Cell Biol, 1994. 14(5): p. 3414-25.
80. Nakada, D., Y. Hirano, and K. Sugimoto, Requirement of the Mre11 complex and
exonuclease 1 for activation of the Mec1 signaling pathway. Mol Cell Biol, 2004.
24(22): p. 10016-25.
81. Tsubouchi, H. and H. Ogawa, A novel mre11 mutation impairs processing of double-
strand breaks of DNA during both mitosis and meiosis. Mol Cell Biol, 1998. 18(1): p.
260-8.
82. Iizuka, M. and M.M. Smith, Functional consequences of histone modifications. Curr
Opin Genet Dev, 2003. 13(2): p. 154-60.
83. Hyland, E.M., et al., Insights into the role of histone H3 and histone H4 core
modifiable residues in Saccharomyces cerevisiae. Mol Cell Biol, 2005. 25(22): p.
10060-70.
84. Masumoto, H., et al., A role for cell-cycle-regulated histone H3 lysine 56 acetylation
in the DNA damage response. Nature, 2005. 436(7048): p. 294-8.
85. Ozdemir, A., et al., Characterization of lysine 56 of histone H3 as an acetylation site
in Saccharomyces cerevisiae. J Biol Chem, 2005. 280(28): p. 25949-52.
86. Giannattasio, M., et al., The DNA damage checkpoint response requires histone H2B
ubiquitination by Rad6-Bre1 and H3 methylation by Dot1. J Biol Chem, 2005.
280(11): p. 9879-86.
87. Huyen, Y., et al., Methylated lysine 79 of histone H3 targets 53BP1 to DNA double-
strand breaks. Nature, 2004. 432(7015): p. 406-11.
88. McGill, C., B. Shafer, and J. Strathern, Coconversion of flanking sequences with
homothallic switching. Cell, 1989. 57(3): p. 459-67.
109
89. Paull, T.T., et al., A critical role for histone H2AX in recruitment of repair factors to
nuclear foci after DNA damage. Curr Biol, 2000. 10(15): p. 886-95.
90. Sanders, S.L., et al., Methylation of histone H4 lysine 20 controls recruitment of Crb2
to sites of DNA damage. Cell, 2004. 119(5): p. 603-14.
91. Clayton, A.L., C.A. Hazzalin, and L.C. Mahadevan, Enhanced histone acetylation
and transcription: a dynamic perspective. Mol Cell, 2006. 23(3): p. 289-96.
92. Doenecke, D. and D. Gallwitz, Acetylation of histones in nucleosomes. Mol Cell
Biochem, 1982. 44(2): p. 113-28.
93. Tamburini, B.A. and J.K. Tyler, Localized histone acetylation and deacetylation
triggered by the homologous recombination pathway of double-strand DNA repair.
Mol Cell Biol, 2005. 25(12): p. 4903-13.
94. Dorigo, B., et al., Chromatin fiber folding: requirement for the histone H4 N-terminal
tail. J Mol Biol, 2003. 327(1): p. 85-96.
95. Kayne, P.S., et al., Extremely conserved histone H4 N terminus is dispensable for
growth but essential for repressing the silent mating loci in yeast. Cell, 1988. 55(1):
p. 27-39.
96. Xu, F., K. Zhang, and M. Grunstein, Acetylation in histone H3 globular domain
regulates gene expression in yeast. Cell, 2005. 121(3): p. 375-85.
97. Chen, C.C., et al., Acetylated lysine 56 on histone H3 drives chromatin assembly after
repair and signals for the completion of repair. Cell, 2008. 134(2): p. 231-43.
98. Adkins, M.W., et al., The histone chaperone anti-silencing function 1 stimulates the
acetylation of newly synthesized histone H3 in S-phase. J Biol Chem, 2007. 282(2): p.
1334-40.
99. Celic, I., et al., The sirtuins hst3 and Hst4p preserve genome integrity by controlling
histone h3 lysine 56 deacetylation. Curr Biol, 2006. 16(13): p. 1280-9.
100. Maas, N.L., et al., Cell cycle and checkpoint regulation of histone H3 K56 acetylation
by Hst3 and Hst4. Mol Cell, 2006. 23(1): p. 109-19.
101. Thaminy, S., et al., Hst3 is regulated by Mec1-dependent proteolysis and controls the
S phase checkpoint and sister chromatid cohesion by deacetylating histone H3 at
lysine 56. J Biol Chem, 2007. 282(52): p. 37805-14.
102. Chen, C.C. and J. Tyler, Chromatin reassembly signals the end of DNA repair. Cell
Cycle, 2008. 7(24): p. 3792-7.
103. Anderson, D.E., et al., Condensin and cohesin display different arm conformations
with characteristic hinge angles. J Cell Biol, 2002. 156(3): p. 419-24.
110
104. Gruber, S., C.H. Haering, and K. Nasmyth, Chromosomal cohesin forms a ring. Cell,
2003. 112(6): p. 765-77.
105. Guacci, V., D. Koshland, and A. Strunnikov, A direct link between sister chromatid
cohesion and chromosome condensation revealed through the analysis of MCD1 in S.
cerevisiae. Cell, 1997. 91(1): p. 47-57.
106. Haering, C.H., et al., Molecular architecture of SMC proteins and the yeast cohesin
complex. Mol Cell, 2002. 9(4): p. 773-88.
107. Michaelis, C., R. Ciosk, and K. Nasmyth, Cohesins: chromosomal proteins that
prevent premature separation of sister chromatids. Cell, 1997. 91(1): p. 35-45.
108. Orr-Weaver, T.L., The ties that bind: localization of the sister-chromatid cohesin
complex on yeast chromosomes. Cell, 1999. 99(1): p. 1-4.
109. Blat, Y. and N. Kleckner, Cohesins bind to preferential sites along yeast chromosome
III, with differential regulation along arms versus the centric region. Cell, 1999.
98(2): p. 249-59.
110. Laloraya, S., V. Guacci, and D. Koshland, Chromosomal addresses of the cohesin
component Mcd1p. J Cell Biol, 2000. 151(5): p. 1047-56.
111. Megee, P.C., et al., The centromeric sister chromatid cohesion site directs Mcd1p
binding to adjacent sequences. Mol Cell, 1999. 4(3): p. 445-50.
112. Tanaka, T., et al., Identification of cohesin association sites at centromeres and along
chromosome arms. Cell, 1999. 98(6): p. 847-58.
113. Toth, A., et al., Yeast cohesin complex requires a conserved protein, Eco1p(Ctf7), to
establish cohesion between sister chromatids during DNA replication. Genes Dev,
1999. 13(3): p. 320-33.
114. Ciosk, R., et al., Cohesin's binding to chromosomes depends on a separate complex
consisting of Scc2 and Scc4 proteins. Mol Cell, 2000. 5(2): p. 243-54.
115. Skibbens, R.V., et al., Ctf7p is essential for sister chromatid cohesion and links
mitotic chromosome structure to the DNA replication machinery. Genes Dev, 1999.
13(3): p. 307-19.
116. Ivanov, D., et al., Eco1 is a novel acetyltransferase that can acetylate proteins
involved in cohesion. Curr Biol, 2002. 12(4): p. 323-8.
117. Ciosk, R., et al., An ESP1/PDS1 complex regulates loss of sister chromatid cohesion
at the metaphase to anaphase transition in yeast. Cell, 1998. 93(6): p. 1067-76.
111
118. Cohen-Fix, O., et al., Anaphase initiation in Saccharomyces cerevisiae is controlled
by the APC-dependent degradation of the anaphase inhibitor Pds1p. Genes Dev,
1996. 10(24): p. 3081-93.
119. Unal, E., et al., A molecular determinant for the establishment of sister chromatid
cohesion. Science, 2008. 321(5888): p. 566-9.
120. Onn, I., et al., Sister chromatid cohesion: a simple concept with a complex reality.
Annu Rev Cell Dev Biol, 2008. 24: p. 105-29.
121. Peters, J.M., A. Tedeschi, and J. Schmitz, The cohesin complex and its roles in
chromosome biology. Genes Dev, 2008. 22(22): p. 3089-114.
122. Strom, L. and C. Sjogren, Chromosome segregation and double-strand break repair -
a complex connection. Curr Opin Cell Biol, 2007. 19(3): p. 344-9.
123. Yanagida, M., Clearing the way for mitosis: is cohesin a target? Nat Rev Mol Cell
Biol, 2009. 10(7): p. 489-96.
124. Kim, S.T., B. Xu, and M.B. Kastan, Involvement of the cohesin protein, Smc1, in
Atm-dependent and independent responses to DNA damage. Genes Dev, 2002. 16(5):
p. 560-70.
125. Sjogren, C. and K. Nasmyth, Sister chromatid cohesion is required for postreplicative
double-strand break repair in Saccharomyces cerevisiae. Curr Biol, 2001. 11(12): p.
991-5.
126. Strom, L., et al., Postreplicative recruitment of cohesin to double-strand breaks is
required for DNA repair. Mol Cell, 2004. 16(6): p. 1003-15.
127. Unal, E., et al., DNA damage response pathway uses histone modification to assemble
a double-strand break-specific cohesin domain. Mol Cell, 2004. 16(6): p. 991-1002.
128. Spencer, F., et al., Mitotic chromosome transmission fidelity mutants in
Saccharomyces cerevisiae. Genetics, 1990. 124(2): p. 237-49.
129. Hanna, J.S., et al., Saccharomyces cerevisiae CTF18 and CTF4 are required for
sister chromatid cohesion. Mol Cell Biol, 2001. 21(9): p. 3144-58.
130. Kouprina, N., et al., CTF4 (CHL15) mutants exhibit defective DNA metabolism in the
yeast Saccharomyces cerevisiae. Mol Cell Biol, 1992. 12(12): p. 5736-47.
131. Kouprina, N., et al., CHL12, a gene essential for the fidelity of chromosome
transmission in the yeast Saccharomyces cerevisiae. Genetics, 1994. 138(4): p. 1067-
79.
112
132. Miles, J. and T. Formosa, Evidence that POB1, a Saccharomyces cerevisiae protein
that binds to DNA polymerase alpha, acts in DNA metabolism in vivo. Mol Cell Biol,
1992. 12(12): p. 5724-35.
133. Mossi, R. and U. Hubscher, Clamping down on clamps and clamp loaders--the
eukaryotic replication factor C. Eur J Biochem, 1998. 254(2): p. 209-16.
134. Mossi, R., et al., Replication factor C interacts with the C-terminal side of
proliferating cell nuclear antigen. J Biol Chem, 1997. 272(3): p. 1769-76.
135. Waga, S. and B. Stillman, The DNA replication fork in eukaryotic cells. Annu Rev
Biochem, 1998. 67: p. 721-51.
136. Mayer, M.L., et al., Identification of RFC(Ctf18p, Ctf8p, Dcc1p): an alternative RFC
complex required for sister chromatid cohesion in S. cerevisiae. Mol Cell, 2001. 7(5):
p. 959-70.
137. Bylund, G.O., J. Majka, and P.M. Burgers, Overproduction and purification of RFC-
related clamp loaders and PCNA-related clamps from Saccharomyces cerevisiae.
Methods Enzymol, 2006. 409: p. 1-11.
138. Ellison, V. and B. Stillman, Biochemical characterization of DNA damage checkpoint
complexes: clamp loader and clamp complexes with specificity for 5' recessed DNA.
PLoS Biol, 2003. 1(2): p. E33.
139. Shimomura, T., et al., Functional and physical interaction between Rad24 and Rfc5
in the yeast checkpoint pathways. Mol Cell Biol, 1998. 18(9): p. 5485-91.
140. Ben-Aroya, S., et al., ELG1, a yeast gene required for genome stability, forms a
complex related to replication factor C. Proc Natl Acad Sci U S A, 2003. 100(17): p.
9906-11.
141. Majka, J. and P.M. Burgers, The PCNA-RFC families of DNA clamps and clamp
loaders. Prog Nucleic Acid Res Mol Biol, 2004. 78: p. 227-60.
142. Bermudez, V.P., et al., The alternative Ctf18-Dcc1-Ctf8-replication factor C complex
required for sister chromatid cohesion loads proliferating cell nuclear antigen onto
DNA. Proc Natl Acad Sci U S A, 2003. 100(18): p. 10237-42.
143. Bylund, G.O. and P.M. Burgers, Replication protein A-directed unloading of PCNA
by the Ctf18 cohesion establishment complex. Mol Cell Biol, 2005. 25(13): p. 5445-
55.
144. Petronczki, M., et al., Sister-chromatid cohesion mediated by the alternative RF-
CCtf18/Dcc1/Ctf8, the helicase Chl1 and the polymerase-alpha-associated protein
Ctf4 is essential for chromatid disjunction during meiosis II. J Cell Sci, 2004. 117(Pt
16): p. 3547-59.
113
145. Formosa, T. and T. Nittis, Dna2 mutants reveal interactions with Dna polymerase
alpha and Ctf4, a Pol alpha accessory factor, and show that full Dna2 helicase
activity is not essential for growth. Genetics, 1999. 151(4): p. 1459-70.
146. Mayer, M.L., et al., Identification of protein complexes required for efficient sister
chromatid cohesion. Mol Biol Cell, 2004. 15(4): p. 1736-45.
147. Gambus, A., et al., GINS maintains association of Cdc45 with MCM in replisome
progression complexes at eukaryotic DNA replication forks. Nat Cell Biol, 2006.
8(4): p. 358-66.
148. Lengronne, A., et al., Establishment of sister chromatid cohesion at the S. cerevisiae
replication fork. Mol Cell, 2006. 23(6): p. 787-99.
149. Ogiwara, H., et al., Ctf18 is required for homologous recombination-mediated
double-strand break repair. Nucleic Acids Res, 2007. 35(15): p. 4989-5000.
150. Ogiwara, H., et al., Chl1 and Ctf4 are required for damage-induced recombinations.
Biochem Biophys Res Commun, 2007. 354(1): p. 222-6.
151. Haber, J.E., Mating-type gene switching in Saccharomyces cerevisiae. Annu Rev
Genet, 1998. 32: p. 561-99.
152. Haber, J.E., Mating-type gene switching in Saccharomyces cerevisiae. Trends Genet,
1992. 8(12): p. 446-52.
153. Jensen, R.E. and I. Herskowitz, Directionality and regulation of cassette substitution
in yeast. Cold Spring Harb Symp Quant Biol, 1984. 49: p. 97-104.
154. Kastan, M.B. and J. Bartek, Cell-cycle checkpoints and cancer. Nature, 2004.
432(7015): p. 316-23.
155. Jessulat, M., et al., Interacting proteins Rtt109 and Vps75 affect the efficiency of non-
homologous end-joining in Saccharomyces cerevisiae. Arch Biochem Biophys, 2008.
469(2): p. 157-64.
156. Morrison, A.J., et al., INO80 and gamma-H2AX interaction links ATP-dependent
chromatin remodeling to DNA damage repair. Cell, 2004. 119(6): p. 767-75.
157. Ramey, C.J., et al., Activation of the DNA damage checkpoint in yeast lacking the
histone chaperone anti-silencing function 1. Mol Cell Biol, 2004. 24(23): p. 10313-
27.
158. Han, J., et al., Rtt109 acetylates histone H3 lysine 56 and functions in DNA
replication. Science, 2007. 315(5812): p. 653-5.
159. Celic, I., A. Verreault, and J.D. Boeke, Histone H3 K56 hyperacetylation perturbs
replisomes and causes DNA damage. Genetics, 2008. 179(4): p. 1769-84.
114
160. Longtine, M.S., et al., Additional modules for versatile and economical PCR-based
gene deletion and modification in Saccharomyces cerevisiae. Yeast, 1998. 14(10): p.
953-61.
161. Xu, H., C. Boone, and G.W. Brown, Genetic dissection of parallel sister-chromatid
cohesion pathways. Genetics, 2007. 176(3): p. 1417-29.
162. Bellaoui, M., et al., Elg1 forms an alternative RFC complex important for DNA
replication and genome integrity. Embo J, 2003. 22(16): p. 4304-13.
163. Chang, M., et al., A genome-wide screen for methyl methanesulfonate-sensitive
mutants reveals genes required for S phase progression in the presence of DNA
damage. Proc Natl Acad Sci U S A, 2002. 99(26): p. 16934-9.
164. Kanellis, P., R. Agyei, and D. Durocher, Elg1 forms an alternative PCNA-interacting
RFC complex required to maintain genome stability. Curr Biol, 2003. 13(18): p.
1583-95.
165. Naiki, T., et al., Chl12 (Ctf18) forms a novel replication factor C-related complex and
functions redundantly with Rad24 in the DNA replication checkpoint pathway. Mol
Cell Biol, 2001. 21(17): p. 5838-45.
166. Recht, J., et al., Histone chaperone Asf1 is required for histone H3 lysine 56
acetylation, a modification associated with S phase in mitosis and meiosis. Proc Natl
Acad Sci U S A, 2006. 103(18): p. 6988-93.
167. Ozenberger, B.A. and G.S. Roeder, A unique pathway of double-strand break repair
operates in tandemly repeated genes. Mol Cell Biol, 1991. 11(3): p. 1222-31.
168. Kim, J.A. and J.E. Haber, Chromatin assembly factors Asf1 and CAF-1 have
overlapping roles in deactivating the DNA damage checkpoint when DNA repair is
complete. Proc Natl Acad Sci U S A, 2009. 106(4): p. 1151-6.
169. Fillingham, J., M.C. Keogh, and N.J. Krogan, GammaH2AX and its role in DNA
double-strand break repair. Biochem Cell Biol, 2006. 84(4): p. 568-77.
170. Wyman, C. and R. Kanaar, DNA double-strand break repair: all's well that ends well.
Annu Rev Genet, 2006. 40: p. 363-83.
171. Brachmann, C.B., et al., The SIR2 gene family, conserved from bacteria to humans,
functions in silencing, cell cycle progression, and chromosome stability. Genes Dev,
1995. 9(23): p. 2888-902.
172. Bekker-Jensen, S., et al., Spatial organization of the mammalian genome surveillance
machinery in response to DNA strand breaks. J Cell Biol, 2006. 173(2): p. 195-206.
173. Houtsmuller, A.B., et al., Action of DNA repair endonuclease ERCC1/XPF in living
cells. Science, 1999. 284(5416): p. 958-61.
115
174. Politi, A., et al., Mathematical modeling of nucleotide excision repair reveals
efficiency of sequential assembly strategies. Mol Cell, 2005. 19(5): p. 679-90.
175. Misteli, T., Beyond the sequence: cellular organization of genome function. Cell,
2007. 128(4): p. 787-800.
176. Bao, Y. and X. Shen, Chromatin remodeling in DNA double-strand break repair.
Curr Opin Genet Dev, 2007. 17(2): p. 126-31.
177. Chai, B., et al., Distinct roles for the RSC and Swi/Snf ATP-dependent chromatin
remodelers in DNA double-strand break repair. Genes Dev, 2005. 19(14): p. 1656-
61.
178. Downs, J.A., et al., Binding of chromatin-modifying activities to phosphorylated
histone H2A at DNA damage sites. Mol Cell, 2004. 16(6): p. 979-90.
179. van Attikum, H., et al., Recruitment of the INO80 complex by H2A phosphorylation
links ATP-dependent chromatin remodeling with DNA double-strand break repair.
Cell, 2004. 119(6): p. 777-88.
180. van Attikum, H. and S.M. Gasser, The histone code at DNA breaks: a guide to
repair? Nat Rev Mol Cell Biol, 2005. 6(10): p. 757-65.
181. Zou, L. and S.J. Elledge, Sensing DNA damage through ATRIP recognition of RPA-
ssDNA complexes. Science, 2003. 300(5625): p. 1542-8.
182. De Koning, L., et al., Histone chaperones: an escort network regulating histone
traffic. Nat Struct Mol Biol, 2007. 14(11): p. 997-1007.
116
Copyright Acknowledgements
Figure 1 has been reprinted from Pu, S., et al., Local coherence in genetic interaction
patterns reveals prevalent functional versatility. Bioinformatics, 2008. 24(20): p. 2376-83
with permission from Oxford University Press, License Number: 2474870650888