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Forkhead transcription factors regulate replication origin firing through dimerization and cell cycle-dependent chromatin binding in S. cerevisiae

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Forkhead transcription factors regulate replication origin firing through dimerization and cell cycle-dependent chromatin binding in S. cerevisiae

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Content Copyright 2014 Andrew Zachary Ostrow
FORKHEAD TRANSCRIPTION FACTORS REGULATE REPLICATION ORIGIN FIRING THROUGH DIMERIZATION
AND CELL CYCLE-DEPENDENT CHROMATIN BINDING IN S. CEREVISIAE

by

Andrew Zachary Ostrow

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

December 2014

ii

Dedication

This is dedicated to my family and friends who continue to make life great.

iii

Acknowledgements

I have enjoyed tremendous support from friends, family and colleagues during both my
time at USC pursuing a Ph.D. and leading up to it. Dr. Oscar Aparicio has been an
outstanding advisor and I am fortunate to have been a part of the innovative lab
environment he has fostered. His approach to science is nuanced and deliberate, and I take
pride in having been scientifically trained by him. He has always been available for
scientific, career, and general life advice. I also thank the other member of my committee,
Dr. Lin Chen, Dr. Matthew Michael, and Dr. Remo Rohs. Dr. Chen has been instrumental in
Chapter 4 of this dissertation, and it has been a pleasure working with him. Dr. Michael has
consistently provided valuable feedback , which has been most appreciated. Dr. Rohs has
been available for excellent discussion, and I look forward to progressing our collaborative
efforts. I would also like to acknowledge Dr. Susan Forsburg, Dr. Irene Chiolo, Dr. Michelle
Arbeitman, and Dr. Ian Ehrenreich for a great deal of constructive feedback.
I would also like to acknowledge Dr. Hinrich Boeger, my undergraduate advisor in UC Santa
Cruz. Dr. Boeger was extremely supportive of me during my time in his lab, and I largely
credit him with helping me progress to the next step. Similarly, Nancy Cox-Konopelski, the
director of an excellent academic program I was involved in during my time at UCSC, was
critical to my decision to pursue a Ph.D.
Members of the Aparicio Lab have greatly enriched my time here intellectually and socially.
Tittu Nellimoottil is an excellent friend and collaborator, I was fortunate to work with him
on the project described in Chapter 3. Jared Peace and I entered the lab together and
defended our dissertations within days of each other, and we’ve had years of constructive
discussion. Simon Knott made the lab a lively place to work and was always more than
happy to contribute his thoughts on a project. Yuan Zhong and I had many helpful and
enjoyable discussions over the years, sitting in the same bay. Yan Gan has also been an
excellent collaborator, and has made critical contributions to Chapter 4. Sandra Villwock is
a valuable member of the lab and I’ve enjoyed our many talks. Jeff Jancuska was an
excellent student and I had a great time working with him. John Zeytounian has been an
excellent member to the lab, and we’ve had a great deal of interesting discussion. From the
Lin Chen Lab, Reza Kalhor was a goodfriend and collaborator, and he made important
contributions to Chapter 4.
My friends, both in and out of science and the Aparicio Lab, have improved my time at USC
tremendously. It’s one thing to enjoy your time in lab, and another to enjoy your time out of
lab, and I’m grateful to my friends for helping me to have both. Dan McCoy is a real mensch,
and if not for him I probably would have forgotten to eat lunch every day like a normal
human. Ian Slaymaker has been a good influence when it comes to getting into trouble, and
we’ve had a lot of fun together. Aysen Erdem was a fun house mate and neighbor, and was a
great (i.e., ‘she let us run loose’) host to countless gatherings. Tara Mastro was a good
friend, key in bringing together the MCB incoming class of ’08. Richard Norte has been a
great friend since elementary school, and being neighbors with him while both getting our
PhDs will certainly be a happy source of nostalgia for the rest of my life. Thanks to Max
iv

Mayer and Clayton Kober for many good times, and Garrick Hogg and Josh Marx for
countless hours of biking. Thanks to Art Sanchez whose level-headedness is contagious,
and perhaps more importantly, for always being up for a BBQ.
My parents Ben and Coral have always been extremely supportive of my pursuits, and
always have been there to help guide me in my choices and foster my intellectual growth.
They are incredible role models, and have taught me the importance of doing what you
love, and I strive to live by their example. I won the parental lottery being raised by them
and my grandmother Oma, and I’m thankful for that every day.
My brother and sister, Aaron and Shana, are very important influences. My older brother’s
love of science and logic tremendously helped direct my own similar interests from an
early age. My sister’s amazing creativity and sense of humor have strongly influenced me
since we were kids. I’m also fortunate that Shana’s taste goes beyond the arts, and my
brother in law Jerrad Xavier has been a great addition to my life, and along with the rest of
my family, he reminds all of us to be good to others and to do good things in life.
Finally, my partner Erin Dollar has been wonderful in supporting me through all of this
time. Erin has an inexhaustible drive to achieve, and she is my inspiration. She has been
cheering me on for the entirety of my undergraduate and graduate studies in science, and
has been a constant source of love, stability, and happiness in my life.

v

Contents

Dedication ..................................................................................................................................................... ii
Acknowledgements ...................................................................................................................................... iii
List of Figures ............................................................................................................................................. viii
Abstract ......................................................................................................................................................... x
Chapter 1: Introduction to Eukaryotic Replication and Forkhead Transcription Factors ............................. 1
1.1 Summary of Replication ................................................................................................................ 1
1.2 Eukaryotic Replication .................................................................................................................. 2
1.3 Introduction to Forkhead Transcription Factors ............................................................................... 10
Chapter 2: Forkhead Transcription Factors Establish Origin Timing and Long-Range Clustering
in S. cerevisiae ............................................................................................................................................. 15
2.1 Background ....................................................................................................................................... 15
Summary ............................................................................................................................................. 15
Introduction ........................................................................................................................................ 17
2.2 Results ............................................................................................................................................... 20
Fkh1 and Fkh2 control genome-wide initiation dynamics of replication origins ................................ 20
Fkh-regulation involves establishment of replication timing domains............................................... 27
Fkh1/2 bind and function in cis to Fkh-activated origins .................................................................... 28
Fkh-dependent origin regulation is not correlated with transcription levels or changes .................. 31
Cdc45 preferentially associates with Fkh-activated origins in G1-phase ........................................... 33
Fkh1/2 are required for selective clustering of Fkh-activated origins in G1-phase ............................ 36
Fkh1 and Fkh2 interact with ORC ........................................................................................................ 40
2.3 Discussion .......................................................................................................................................... 42
Fkh1/2 establish replication-timing domains through origin clustering............................................. 42
Multiple, separable roles for Fkh1 and Fkh2 in regulation of the genome ........................................ 45
2.4 Experimental Procedures .................................................................................................................. 48
Yeast strains and methods .................................................................................................................. 49
Preprocessing of sequence data ......................................................................................................... 49
Analysis of linear clustering of Fkh-regulated origins ......................................................................... 50
Analysis of Fkh1 and Fkh2 binding sites .............................................................................................. 51
Analysis of global 4C ........................................................................................................................... 51
vi

Extended Experimental Procedures ........................................................................................................ 52
Yeast strain and plasmid constructions .............................................................................................. 52
Antibody methods. ............................................................................................................................. 53
BrdU-IP-Seq analysis ........................................................................................................................... 54
BrdU-IP-chip time-course data analysis .............................................................................................. 55
Analysis of linear clustering of Fkh-regulated origins ......................................................................... 55
Analysis of Fkh-regulated transcription versus Fkh-regulated origin function ................................... 56
Chromosome conformation capture on chip (4C) .............................................................................. 57
Chapter 3: Analysis of Fkh1 and Fkh2 Chromatin Binding .......................................................................... 61
3.1 Background ....................................................................................................................................... 61
3.2 Results ............................................................................................................................................... 63
An expanded map of Fkh1 and Fkh2 binding to the S. cerevisiae genome ........................................ 63
Fkh1 and Fkh2 are associated with distinct chromatin architectures ................................................ 68
Fkh1 and Fkh2 binding at regulated genes ......................................................................................... 69
Fkh1 and Fkh2 binding at replication origins ...................................................................................... 71
Other genetic elements associated with Fkh1 and Fkh2 binding ....................................................... 76
Cell cycle dynamics of Fkh1 and Fkh2 binding .................................................................................... 77
3.3 Discussion .......................................................................................................................................... 83
An expanded map of Fkh1 and Fkh2 binding to the S. cerevisiae genome ........................................ 83
Fkh1 and Fkh2 binding at replication origins ...................................................................................... 85
Novel genetic elements associated with Fkh1 and Fkh2 binding ....................................................... 86
3.3 Methods ............................................................................................................................................ 88
Yeast strains and methods .................................................................................................................. 88
Microarray data analysis and peak calling .......................................................................................... 89
Analysis of intersection between datasets ......................................................................................... 89
Calling Fkh1-only, Fkh2-only, and Fkh1and2 sets ............................................................................... 89
Analysis of Fkh1/2 enrichment at genetic .......................................................................................... 90
Cell cycle analysis of binding ............................................................................................................... 91
Data Accession .................................................................................................................................... 91
Chapter 4: Forkhead Transcription Factors Regulate Replication through Dimerization and Cell Cycle
Specific Binding at Origins ........................................................................................................................... 93
4.1 Background ....................................................................................................................................... 93
vii

4.2 Results ............................................................................................................................................... 94
Yeast Forkhead proteins share key homologies with mammalian FOXP proteins ............................. 94
Yeast Forkhead dimerization .............................................................................................................. 96
Forkhead domain swap mutants have knock-out replication profiles ............................................... 98
Forkhead dimerization is not required for CLB2-cluster gene regulation ........................................ 103
Chromatin binding of Forkhead domain swap mutants ................................................................... 106
Forkhead binding motifs differ at various Forkhead-regulated loci ................................................. 110
4.3 Discussion ........................................................................................................................................ 113
Fkh1/2 and epigenetic regulation ..................................................................................................... 113
Binding specificities of Forkhead proteins are altered spatially and temporally ............................. 113
Fkh-dsm as a tool for fundamental investigations............................................................................ 114
Residual activities of Fkh-dsm in replication timing ......................................................................... 115
Alternative mechanism of Forkhead regulation of replication timing or genome architecture ...... 115
Future directions ............................................................................................................................... 116
4.4 Methods .......................................................................................................................................... 118
Yeast strains and methods ................................................................................................................ 118
BrdU-IP-Seq ....................................................................................................................................... 118
Agar Scarring Assay ........................................................................................................................... 118
ChIP-chip ........................................................................................................................................... 119
Preprocessing of microarray data ..................................................................................................... 119
Analysis of global replication and Fkh1 occupation at features ....................................................... 119
Peak calling and differential origin firing .......................................................................................... 119
Analysis of Fkh1 dimerization using pull-down assay ....................................................................... 119
Supplemental Figures ............................................................................................................................... 121
References ................................................................................................................................................ 153
Chapters 1, 3,4 ...................................................................................................................................... 153
Chapter 2 ............................................................................................................................................... 164

viii

List of Figures

Figure 1.1. Anatomy of S. cerevisiae Replication Origins
Figure 2.1. Analysis of early S-phase BrdU incorporation
Figure 2.2. Temporal analysis of DNA replication by BrdU pulse-labeling
Figure 2.3. Analysis of Fkh1 and Fkh2 binding sites near origins
Figure 2.4. Transcription analysis surrounding Fkh-regulated origins in unsynchronized
and G1-synchronized cells
Figure 2.5. Genome-wide binding of replication initiation factors to Fkh-regulated origins
Figure 2.6. Chromosome-conformation capture analyses of origin interactions
Figure 2.7. Co-IP of Fkh1 with ORC
Figure S2.1. Related to Figure 1A
Figure S2.2. Related to Figure 2A
Figure S2.3. Related to Figure 6
Figure 3.1. Genome-wide analysis of Fkh1 and Fkh2 chromatin binding
Figure 3.2. Correlation of Fkh1 and Fkh2 binding sites identified in different experiments
Figure 3.3. Distinct nucleosome positioning at Fkh1-only loci versus loci that bind Fkh2
Figure 3.4. Fkh1 and Fkh2 binding with target genes
Figure 3.5. Fkh1 and Fkh2 binding with replication origins
Figure 3.6. Analysis of Fkh1 and Fkh2 binding proximal to various genetic elements
Figure 3.7. Cell cycle analysis of Fkh1 and Fkh2 binding
Figure 3.8. G1-specific binding of Fkh1/2 at Fkh-activated origins
Figure 4.1. Forkhead Sequence Analysis
Figure 4.2. Fkh1/2 Structure Model
ix

Table 4.1. Mutations introduced to Fkh1 and Fkh2 to create fkh1-dsm and fkh2-dsm
Figure 4.3. Fkh1 Pull-down Assay
Figure 4.4. Fkh-dsm BrdU Plots
Figure 4.5. Correlation coefficients between BrdU-IP-Seq of Forkhead mutants
Figure 4.6. Fkh-dsm and Fkh knockout strains have similar genome-wide replication
profiles
Figure 4.7. Fkh-dsm BrdU Heat Map Ratio
Figure 4.8. Forkhead mutant
Figure 4.9. Agar scarring assay of wild type and Forkhead mutants
Figure 4.10. Forkhead binding and BrdU incorporation at TEM1 and ARS1307
Figure 4.11. ChIP-chip Origin Classes and CLB2 Cluster Heat Map
Figure 4.12. EMSA of Fkh1-DBD and fkh1-dsm-DBD at Fkh-regulated loci
Figure 4.13. Strong and weak Forkhead motifs at origins by Fkh-regulation
Figures S3.1.1 – 2.1.16. Genome-wide analysis of Fkh1 and Fkh2 chromatin binding
Figures S3.2.1 – 2.2.16. Cell cycle binding of Fkh1/2 genome-wide
x

Abstract

Forkhead box (FOX) transcription factors regulate a wide variety of cellular functions in
higher eukaryotes, including cell cycle control and developmental regulation. In
Saccharomyces cerevisiae, Forkhead proteins Fkh1 and Fkh2 perform analogous functions,
regulating genes involved in cell cycle control, while also regulating mating-type silencing
and switching involved in gamete development. As described here, we revealed a novel
role for Fkh1 and Fkh2 in the regulation of replication origin initiation timing, which, like
donor preference in mating-type switching, appears to involve long-range chromosomal
interactions, suggesting roles for Fkh1 and Fkh2 in chromatin architecture and
organization. To elucidate how Fkh1 and Fkh2 regulate their target DNA elements and
potentially regulate the spatial organization of the genome, we undertook a genome-wide
analysis of Fkh1 and Fkh2 chromatin binding by ChIP-chip using tiling DNA microarrays.
Our results confirm and extend previous findings showing that Fkh1 and Fkh2 control the
expression of cell cycle-regulated genes. In addition, the data reveal hundreds of novel loci
that bind Fkh1 only and exhibit a distinct chromatin structure from loci that bind both
Fkh1 and Fkh2. The findings also show that Fkh1 plays the predominant role in the
regulation of a subset of replication origins that initiate replication early, and that Fkh1/2
binding to these loci is cell cycle-regulated. Finally, we demonstrate that Fkh1 and Fkh2
bind proximally to a variety of genetic elements, including centromeres and Pol III-
transcribed snoRNAs and tRNAs, greatly expanding their potential repertoire of functional
xi

targets, consistent with their suggested role in mediating the spatial organization of the
genome.
Sequence comparison and structural modeling of Fkh1 and Fkh2 with mammalian FOXP
family proteins reveals key homologies that suggest Fkh1 and Fkh2 are able to form
domain swapped dimers, providing a possible mechanism by which Fkh1 and Fkh2 bind
distal origins and sustain replication foci through dimerization in trans. Altering residues
thought to be involved in dimerization, we created domain swap mutants (dsm) Forkhead
proteins. We show that wild type Fkh1 dimerizes, and that this is diminished in fkh1-dsm.
BrdU-IP-Seq analysis of global replication in fkh-dsm reveals a null-like phenotype, yet
Forkhead regulation of transcription appears intact. Assessment of chromatin binding
using ChIP-chip shows that fkh1-dsm sufficiently binds at the promoters of CLB2-cluster
promoters, while binding at early origins is ablated, indicating a role of the mutated
residues in cell cycle-specific binding of Fkh1 at replication origins. Finally, we find that the
Forkhead binding motifs found at early origins differ from those found elsewhere,
supporting a role of alternative sequence specificities in Forkhead binding and regulation
of replication origins.
1

Chapter 1: Introduction to Eukaryotic Replication and Forkhead Transcription
Factors

1.1 Summary of Replication

Faithful duplication of genetic material in preparation for cell division and proliferation is
perhaps the most integral process a living cell must be able to perform. This process, DNA
replication, is governed by multiple layers of regulation in order to ensure that
chromosomes are properly duplicated, avoiding excessive mutagenesis and physical
collapse of DNA structures during replication. At a basal level, replication is regulated by
the binding of replication machinery to replication origins, specific chromosomal regions
where replication initiates. After an origin initiates replication, or “fires”, replication
machinery including polymerases, helicases, and other factors (together, “replisomes”)
extend bidirectionally along the chromosome at two replication forks until they encounter
neighboring replisomes, at which point replication of that region between two origins has
completed. However, DNA replication is not the only process occurring on DNA;
transcription, chromatin remodeling, and DNA repair, for example, are active throughout
the cell cycle, including in S phase during which DNA replication occurs. Interference of
these processes with replication can lead to replication fork collapse, a variety of
mutations, and cell death. These processes must therefore be coordinated spatially and
temporally to prevent disruption of replication.

2

1.2 Eukaryotic Replication

A replication origin must be prepared during G1 phase in order to fire during S phase.
Origin preparation, or licensing, is regulated in a cell cycle-dependent manner, with each
origin licensed for firing no more than once per cell cycle to prevent a particular region
from being replicated twice, a typically deleterious phenomenon known as re-replication.
The precise attributes of origins, such as conserved motifs and size ranges, differ among
eukaryotes. Replication in yeast is prototypical for metazoan replication origin selection,
and studies of yeast replication have provided great insight into conserved mechanisms
employed in eukaryotic DNA replication. In the budding yeast Saccharomyces cerevisiae,
replication origins were initially discovered as autonomously replicating sequences (ARSs),
100-200 base pair (bp) regions of chromosomes which when cloned into a vector allowed
the duplication and maintenance of the resulting plasmids through cell divisions. Many
hundred ARSs have been identified, and they are located primarily in intergenic regions
where they are less likely to encounter interference from transcription machinery.
Sequence analysis has revealed that ARSs contain a conserved A/T-rich element termed the
ARS Consensus Sequence (ACS) required for binding of the Origin Recognition Complex
(ORC), a six-subunit protein complex found at all origins that is required for origin
licensing and serves as a scaffold for other replication machinery. More precise analysis of
ARSs identified several additional features involved in replication origin activity. These
features, A and B1 through B4 by distance from the ACS, are found within a region
approximately 100 bps downstream of the ACS termed the B domain. Each of the four B
elements confers distinct regulatory properties to replication origins, and the specific B
element composition of a given origin varies due to the lack of complete B element
3

conservation in ARSs. An additional regulatory region termed the C domain was identified
upstream of the ACS.
The A element contains the ACS and is required for ORC binding. Of the five A and B
elements, the A element is the only one that is essential; deletion of the A element of an
origin ablates replication initiation. The B1 element is next proximal to the ACS, and, along
with the ACS in the A element, is involved in origin stimulation through binding of ORC and
additional replication factors
1
. Mutagenesis of certain B1 element bases decreases origin
function without reducing ORC binding
1
. The A/T-rich B2 element contains a DNA
Unwinding Element (DUE) involved in helicase function, and binds the single stranded DNA
(ssDNA) binding protein RPA, which is essential for replication initiation and elongation
2–
10
. RPA binds origins in a CDK- and DDK-dependent manner after loading of the MCM
complex
11
. The B3 element is a binding site for the transcription factor Abf1 at origins
including ARS1 and stimulates origin activity. The ability to stimulate replication is not
limited to Abf1; recruitment of Gal11, Rap1 or factors involved in the transcriptional
activation of RNA polymerase III-transcribed gene SNR6 relocated to ARS1, for example,
stimulated replication of the origin
12–14
. Additional transcription factors have been
demonstrated to have roles in origin stimulation at other origins, including Sum1 at HML-
E
15
. The B4 element also stimulates origin activity, and can substitute for other B elements
in their absence through unknown mechanisms
3
. Finally, the C domain upstream of the ACS
can substitute for various B elements, and at some origins including ARS121 contains a
binding site for Mcm1, another transcription factor involved in origin stimulation
16,17
.
4

Figure 1.1. Anatomy of S. cerevisiae Replication Origins. Budding yeast origins are well
defined and contain various domains including A, B elements, and C (not pictured).
Different origins contain different combinations of B and C elements, and each contribute
to origin activity through various functions. Adapted from Yoshida et al, 2013
18
.

Regulation of eukaryotic replication is dependent upon protein activity leading to the
licensing and initiation of replication origins. Origin assembly begins with binding of the
ORC to DNA, marking potential sites of replication initiation. ORC, along with Cdc6 and
Cdt1, loads inactive minichromosome maintenance (Mcm2-7, or MCM) helicase complex.
ORC, Cdc6 and Mcm2-7 are ATPases, and ATP binding by this complex facilitates proper
loading and stabilization of an active MCM double hexamer and release of Cdt1, forming
the pre-replication complex (pre-RC) and “licensing” the origin for later activation
19,20
. It is
critical to genome integrity that a given chromosomal position is replicated exactly one
time per cell cycle, and this is achieved through cell cycle regulated origin assembly and
firing. Under normal circumstances, origin licensing can only occur in G1 phase; after an
origin initiates, a pre-RC cannot form at that location until the next G1 phase, after
replication and subsequent cell division have completed. Cell cycle regulated origin
initiation is achieved by Dbf4-dependent-kinase (DDK) and cyclin-dependent-kinase (CDK),
which are activated and subsequently act on origins in G1 phase and beginning at the G1/S
phase transition, respectively. First, DDK phosphorylates the MCM complex and loads
initiation factors Cdc45 and Sld3. CDK then phosphorylates Sld3 and replication factor Sld2
5

and, along with Cdc45, leads to recruitment of a number of additional components of the
replisome including GINS, which plays a role in activation of the MCM helicase, and DNA
polymerases
21
.
After replication initiates, two replisomes extend from the origin in each direction until
terminating, usually by meeting an incoming replisome emanating from an adjacent origin.
Replisomes have more to contend with besides each other, and replication is therefore
regulated on another level to coordinate with transcription, chromatin remodeling, and
other genomic events. A given replication origin must go through the same licensing and
initiation steps outlined above, but other factors act to alter the timing and efficiency of
origins. The efficiency of an origin is a measure of the frequency with which it will initiate
during a given S phase across a population of cells, in a range of 0 to 1. Origin timing is a
measure of an origin’s temporal regulation, how much time passes between initiation of S
phase and initiation of the origin, between early to late S phase. Due to variability in origin
timings and efficiencies, individual cells will initiate various subsets of origins in a given S
phase. However, when is measured over a population, replication profiles are consistent
average representations of origin activity
22
. In a given cell, an efficient and early origin will
therefore usually initiate in S phase, as opposed to remaining dormant, and when it does it
is likely to do so early in S phase. Conversely, an inefficient and late origin will typically not
initiate, but when it does it is more likely to do so at a characteristic time near the end of S
phase. When an origin does not initiate, or if adjacent origins initiate sufficiently earlier, the
chromosomal region surrounding that origin will replicate passively; that is, replisomes
from adjacent origins will pass through the origin, replicating its chromosomal
neighborhood.
6

Measures of origin timing and efficiency are not, in principle, entirely independent of one
another. For example, a very early origin will intrinsically appear more efficient than a late
origin. A late origin has a high probability of being passively replicated by an earlier
neighboring origin even if it is highly efficient. As such, it will initiate less frequently, that is,
with an apparent lower efficiency, than the true efficiency of the origin. Genetically altering
the neighboring origins to remove their ability to initiate (or, “deleting” the origins)
removes their capability to passively replicate the late origin, allowing it to initiate with a
frequency closer to its potential. It has therefore been proposed that origin timing and
efficiency are both products of the same regulatory mechanisms. Precisely what these
mechanisms are and how they influence origin timing and efficiency are largely unclear,
and discovering and characterizing them in molecular detail is a major goal in the field.
The functionality of replication origins depends not only on their composition of origin
elements, but on larger scale chromosomal factors such as the broader neighborhood in
which an origin is located. Chromosomal regions like those near centromeres and
telomeres contain origins with characteristic replication times. Subtelomeric origins are
late-replicating and often inefficient to the point of dormancy, with their adjacent regions
duplicating entirely by passive replication
23
. In contrast, early origins are found near every
centromere. Experiments relocating early origin sequences to late replicating regions, and
vice versa, revealed that origin timing is dependent upon local chromosome context; the
timing of an origin can be altered by relocating it to a region with a different characteristic
time of replication. Relocating origins with substantial flanking sequence allows them to
maintain their timing, suggesting that local chromatin environment is involved in the
determination of origin timing
24,25
. Indeed, in addition to the differences in centromeric
7

and subtelomeric replication timing, differences in local chromatin environment affect
replication timing, with other heterochromatic regions replicating later than euchromatic
regions, often doing so passively
26–28
. This has been demonstrated in studies analyzing the
replication activity in strains lacking SIR3 and RPD3, factors that are involved in the
silencing of chromatin. SIR3 (Silent Information Regulator 3) is involved in subtelomeric
heterochromatin assembly, and is required for the integrity and subnuclear localization of
telomeres and, along with SIR4, represses gene expression in telomeres
29
. Deletion of SIR3
leads to initiation of the normally dormant subtelomeric replication origins, and relocation
of ARS1 into a subtelomeric region in the presence of Sir3 delays its initiation
26
. The
transcriptional regulator RPD3 is a histone deacetylase involved in chromatin silencing and
repression of gene expression. Studies of the effect of RPD3 on origin activity show that
Rpd3 delays initiation of most nontelomeric origins, correlating with Rpd3-regulated gene
expression and histone deacetylation. Additionally, direct recruitment of the histone
acetyltransferase Gcn5 to a late origin advances its initiation timing, implying an effect of
histone acetylation on replication timing
30–32
.
In addition to local chromatin environment, other factors like subnuclear localization,
chromosomal positioning, and global chromatin structure have been implicated in the
regulation of replication origin initiation. Both centromeres and telomeres have
characteristic times of replication initiation; early and late, respectively. Studies on
centromeric replication timing have been carried out to determine whether centromeres
impart early timing or if centromeres reside in otherwise early-replicating regions.
Deletion of centromeres in the budding yeast Candida albicans lead to the establishment of
new centromeres (neo-CENs) away from the original CEN site. An early-firing origin
8

formed proximal to the neo-CEN consistent with the idea that centromeric factors impart
early timing to the regions in which they reside
33
. In S. cerevisiae, experiments relocating
centromeres further supported this idea. Relocation of a CEN into a normally late-firing
region caused that region to replicate early. This effect was dependent upon the
functionality of the CEN, limiting the cause of the replication timing advancement to the
centromere itself
34
. Early CEN replication in the fission yeast Schizosaccharomyces pombe
has been demonstrated to be caused by DDK-recruitment to centromeric origins
35
.
Subtelomeric origins, as noted, replicate late in part due to their repressive,
heterochromatic local structure, and ablation of this structure leads to earlier firing.
Additionally, deletion of YKU70, a component of the Ku complex involved in telomere
localization to the nuclear periphery, leads to advancement of replication timing initiation
of origins near telomeres and within subtelomeric repeats, though tethering to the nuclear
periphery is not always sufficient, as demonstrated with the very early and efficient
ARS607
36
.
Initiation factors Cdc45 and Sld3 are loaded at pre-RCs by DDK in a replication timing-
dependent manner, with binding at origins occurring relative to the initiation timing of the
origin, indicating that initiation factor loading is the rate limiting step of origin
initiation
37,38
. Cdc45-Sld3 binds at early origins (including CEN-proximal origins) in G1-
phase, when the replication timing program is established, leading to the suggestion that
Cdc45 is a limiting factor that is preferentially accessible to early origins
37,39,40
. Several
studies in Xenopus, S. pombe, and S. cerevisiae have shown that overexpression of Cdc45
and other factors leads to an increase in the amount of both total replication and amount of
replication origins that initiate, supporting the idea that initiation factors are limiting
41–46
.
9

Beginning in late G1 phase, multiple origins converge to form replication foci, clusters of
origins and replication machinery that co-localize in the nucleus
47,48
. In agreement with
similar studies of mammalian cells, genome-wide conformation capture analysis of the
yeast genome detect contacts between early origins, indicating that replication foci contain
multiple origins with similar initiation times, including centromeric origins and early,
Cdc45-bound origins
49
. This suggests that the preferential access of early origins to
initiation factors is regulated by global genome structure. This thesis will describe factors
involved in such regulation and detail the mechanism by which they perform.

10

1.3 Introduction to Forkhead Transcription Factors

Forkhead Box (Fox) proteins comprise a large family of transcription factors involved in a
wide variety of cellular functions in eukaryotes from yeast to humans, ranging from
development and differentiation, cell cycle control, aging and stress response, including
apoptosis
50
. The uniting feature of Fox proteins is the Forkhead DNA Binding Domain (Fkh-
DBD), a ‘winged helix’ domain that recognizes a core conserved sequence (RYMAAYA),
where the ‘wings’ of the Fkh-DBD are responsible for recognition of flanking sequence,
providing a high degree of sequence specificity and variability of full binding motifs among
Fox factors. A subset of mammalian Fox proteins have been characterized as pioneer
transcription factors for their ability to recognize the Forkhead core sequence in
compacted chromatin, bind and remodel the chromatin to allow gene expression. Certain
Fox proteins act as canonical transcription factors functioning in activation or repression of
gene expression through the recruitment of chromatin-modifying co-factors. Fox proteins
have also been implicated in the formation of higher order chromatin structure and the
epigenetic regulation of gene expression and perhaps additional cellular functions
50–52
.
Four Forkhead proteins are present in the S. cerevisiae proteome, each containing a Fkh-
DBD: Fkh1, Fkh2, Hcm1 and Fhl1. Fhl1 is a regulator of ribosomal protein transcription, but
its Fkh-DBD has diverged substantially and it does not bind the Fkh motif
53
. Hcm1
promotes cell cycle progression through S phase through transcriptional regulation of
genes including FKH1 and FKH2
54
. Fkh1 and Fkh2 are the most closely related of the S.
cerevisiae Forkhead proteins, and share the greatest degree of similarity in both protein
sequence and recognized motif sequence
55–57
. Homology between Fkh1 and Fkh2 includes
11

significant similarity outside of the Forkhead domain, with both sharing a similar
Forkhead-associated (FHA) domain, an additional conserved region found in a subset of
Fox proteins that is involved in regulation of protein function. Fkh2 contains an additional
C-terminal regulatory domain
58,59
. Together, Fkh1 and Fkh2 regulate the CLB2-cluster, a set
of ~33 genes expressed during late S/G2 phase that promote cell cycle progression through
the G2/M phase transition
60
.
Combined deletion of FKH1 and FKH2 leads to deregulation of CLB2-cluster genes, severely
diminishing their expression levels and abolishing their pattern of cell cycle-regulated
expression
60
. Additionally, CLB2-cluster deregulation caused by deletion of these genes
induces pseudohyphal growth, a differentiated state in which cells become elongated,
remain attached after cell division, readily flocculate in liquid media and burrow into solid
growth substrate
60,61
. Single deletion of either FKH1 or FKH2 leads to less severe
transcriptional deregulation and does not induce pseudohyphal growth; thus, Fkh1 and
Fkh2 partially complement one another. The proteins do not have redundant roles,
however, as the deregulation of CLB2-cluster genes in the single deletions lead to different
transcriptional phenotypes: FKH1 deletion lacks transcriptional repression of CLB2-cluster
genes during G1 phase, and FKH2 deletion is defective in activation of CLB2-cluster gene
activation during S/G2 phase
59–62
. Although both Fkh1 and Fkh2 are thought to be involved
in repression of CLB2-cluster genes, Fkh2, but not Fkh1, also acts with transcription factor
Mcm1 to cooperatively bind DNA to activate cell cycle expression of CLB2-cluster
57,63
.
Transcription factor Ndd1 is a coactivator of Mcm1-Fkh2-mediated transcriptional
activation. Ndd1 and Fkh2 physically interact in a manner reinforced by Clb5-Cdk1-
mediated phosphorylation of the Fkh2 C-terminal domain, promoting interaction with
12

Ndd1, and Clb2-Cdk1-phosphorylated Ndd1 binds the FHA domain of Fkh2, leading to gene
activation
64–66
. NDD1 is essential both for CLB2-cluster gene activation and cell survival.
This lethality is overcome by deletion of FKH2 or the C-terminal regulatory domain of
FKH2, but not FKH1, supporting the idea that the function of Ndd1 in activating gene
expression is to overcome repression by Fkh2-Mcm1
66,67
.
In contrast to the major and well-studied role of Fkh2 in gene regulation, Fkh1 alone is
involved in the regulation of S. cerevisiae mating-type switching
68
. Mating-type switching is
the process of gene conversion at that MAT locus, and involves an HO-endonuclease-
induced dsDNA break at this locus and its repairing it by homologous recombination using
one of two mating type donor alleles (a or α) from genomic loci at opposite ends of
chromosome III. The result is a gene conversion of the mating type allele at the MAT locus
to that of the donor allele. Mating-type switching to each of the loci does not occur equally,
and MATa preferentially (approximately 90%) recombines with HML α over HMRa,
resulting in mating-type switch; this is referred to as “donor preference”. Donor preference
is the product of a cis-acting Recombination Enhancer (RE) proximal to HML α that binds
multiple copies of Fkh1 in MATa cells. Deletion of either RE or FKH1 eliminates donor
preference, and tethering of the Fkh1-FHA domain at a RE deletion is sufficient to restore
donor preference. FKH2 deletion does not affect donor preference
69–71
. Thus, Fkh1
facilitates long-range chromosomal interactions in this specific process.
More recently, we reported that Fkh1 and Fkh2 are involved in the establishment of the
replication timing program through a mechanism involving long-range chromosomal
interactions resulting in the clustering of early origins (discussed in Chapter 2)
72
. Deletion
13

of both FKH1 and FKH2 significantly alters the initiation timing of most early and late
replication origins genome-wide. Sequence analysis of early origins that are delayed in
fkh1Δfkh2Δ (referred to as Fkh-activated origins) revealed an enrichment of the Fkh-
binding motif, supporting a direct role for Forkhead proteins in determination of origin
timing. Single deletion of FKH1 had a more modest effect on global replication time,
altering the timing of ~50 early and late origins, and single deletion of FKH2 had no effect,
suggesting that Fkh1 plays the predominant role in determining replication timing, but that
Fkh2 can partially compensate in the absence of Fkh1. Investigation into the basis of this
difference is ongoing, but we show that differential binding of Fkh1 and Fkh2 at early
origins may partially explain this finding (discussed in Chapter 3).
As discussed in Chapter 2, we further examined the means by which Fkh1 and Fkh2
regulate replication initiation timing, first by chromatin immunoprecipitation analyzed by
microarray chip (chIP-chip) of pre-RC components and initiation factors. We found no
difference in MCM or ORC binding at origins with delayed or progressed initiation timing in
fkh1Δfkh2Δ, but did find that Cdc45 binding was diminished at Fkh-activated origins, and
increased at origins with advanced timing in fkh1Δfkh2Δ (Fkh-repressed origins),
indicating that in fkh1Δfkh2Δ origin licensing is not affected, but that loading of initiation
factors is altered. Although Fkh1 and Fkh2 are transcription factors involved in cell cycle
progression, the phenotype is not caused by transcriptional deregulation in fkh1Δfkh2Δ.
Gene expression as analyzed by high-throughput sequencing of RNA (RNA-Seq) revealed
that the replication defect is not due to local changes in transcription around origins.
Additionally, we introduced a plasmid-borne C-terminally-truncated FKH2 (pFKH2ΔC),
which rescues the pseudohyphal morphological defect caused by CLB2-cluster gene
14

deregulation of fkh1Δfkh2Δ cells. This strain exhibited normal morphology and growth,
indicating normal transcriptional regulation of the CLB2-cluster, but maintained the
replication initiation defect of fkh1Δfkh2Δ. Moreover, deletion of Forkhead binding motifs
at the early origin ARS305 in FKH1 FKH2 cells maintains pre-RC assembly while delaying
origin initiation; coupled with the enrichment of Forkhead binding motifs at Fkh-activated
origins, this suggests a direct role of Fkh1 and Fkh2 at origins in regulating the replication
timing program
72
.
Consistent with the observation that early origins cluster into replication foci, analysis of
global chromosome capture data revealed that Fkh-activated origins form contacts with
one another in late G1 phase
49,72
. This was confirmed with chromosome conformation
capture analysis of early origins ARS305 and ARS607 in wild type cells. However, in the
absence of FKH1 and FKH2, these early origins fail to make chromosomal contacts. Along
with the observation that Fkh1 and Fkh2 bind early origins, this indicates that the proteins
act to regulate global genome architecture by directly binding and recruiting early origins
into early-firing replication factories. This leads us to a model in which Forkhead factors
facilitate chromosomal interactions that provide origins with access to limiting initiation
factors. Interrupting these genomic interactions through deletion of FKH1 and FKH2 leads
to destabilization of the replication timing program by abolishing the preferential access of
certain origins to initiation factors, allowing Fkh-repressed origins to passively access
these factors and initiate with an earlier timing
72
. Elucidation of the mechanism of origin
recognition and clustering by Forkhead transcription factors is a major goal of this work.

15

Chapter 2: Forkhead Transcription Factors Establish Origin Timing and Long-Range
Clustering in S. cerevisiae

2.1 Background

The following chapter is taken from the 2012 Cell research article of the same name by
Simon R.V. Knott, Jared M. Peace, A. Zachary Ostrow, Yan Gan, Alexandra E. Rex,
Christopher J. Viggiani, Simon Tavaré, and Oscar M. Aparicio. Supplemental figures have
been inserted into the text; additional information such as supplemental tables are
available from Cell Press. My experimental contributions to this paper include: creation of
fkh1Δ (ZOy1) and subsequent first BrdU-IP-chip to identify its replication phenotype,
involvement in the time course experiments, involvement in ChIP-chip of replication
initiation factors, conception, along with Tittu Nellimmoottil, of the idea to use ‘4C’ to
detect chromatin interactions.

Summary: The replication of eukaryotic chromosomes is organized temporally and
spatially within the nucleus through epigenetic regulation of replication origin function.
The characteristic initiation timing of specific origins is thought to reflect their chromatin
environment or sub-nuclear positioning, however the mechanism remains obscure. Here
we show that the yeast Forkhead transcription factors, Fkh1 and Fkh2, are global
determinants of replication origin timing. Forkhead regulation of origin timing is
independent of local levels or changes of transcription. Instead, we show that Fkh1 and
16

Fkh2 are required for the clustering of early origins and their association with the key
initiation factor Cdc45 in G1-phase, suggesting that Fkh1 and Fkh2 selectively recruit
origins to emergent replication factories. Fkh1 and Fkh2 bind Fkh-activated origins, and
interact physically with ORC, providing a plausible mechanism to cluster origins. These
findings add a new dimension to our understanding of the epigenetic basis for differential
origin regulation and its connection to chromosomal domain organization.
17

Introduction

Chromatin structure and organization influence most every genomic process (reviewed in
(Jenuwein and Allis, 2001; Misteli, 2007)). Modification of chromatin structure to
accommodate one genomic task inevitably alters the landscape for other processes. To
function concurrently, fundamental processes such as DNA replication and transcription
must be coordinated to preserve the accuracy and integrity of both, failure of which may
lead to genome instability and developmental defects (reviewed in (Gondor and Ohlsson,
2009; Hiratani et al., 2009; Knott et al., 2009a; Mechali, 2010)). Epigenetic regulation of
replication origin activation is thought to play a role in coordinating DNA replication with
other genomic tasks, however our current understanding of how chromosomal replication
is regulated by chromatin, let alone organized in three dimensions, is mostly correlative
and sparse on mechanism.

Chromosomal DNA replication is governed primarily through regulation of replication
initiation at origins. Origin DNA binds ORC and these are joined, in G1-phase, by inactive
MCM helicase complexes resulting in assembly of pre-replicative complexes (pre-RCs),
which are competent to initiate replication. Upon S-phase entry, Dbf4-dependent kinase
(DDK) stimulates the loading of Cdc45 and Cyclin-dependent kinase stimulates the loading
of additional factors to convert pre-RCs into active replisomes (reviewed in (Bell and Dutta,
2002)). However, not all pre-RCs initiate replication synchronously at the onset of S-phase,
nor do all potential origins fire in every cell across a population. Instead, a subset of pre-
RCs initiates replication early while clustering into foci, each containing multiple
18

replisomes, that constitute replication factories (Kitamura et al., 2006; Meister et al., 2007).
The dynamic nature of the replication foci suggests that as early replicons terminate, these
factories are disassembled, allowing the next subset(s) of pre-RCs to initiate replication
and establish new factories (Sporbert et al., 2002). The process is not purely stochastic.
Whether in yeast or mammalian cells, certain origins reproducibly initiate more efficiently
(ie, the frequency of initiation per cell cycle, ≤1) and/or earlier than others (across a
population of cells), thereby giving rise to characteristic replication timing patterns of
chromosomes (reviewed in (Diller and Raghuraman, 1994; Weinreich et al., 2004)).

Replication timing generally correlates with gene activity and chromatin structure, with
earlier replicating regions being transcriptionally active and euchromatic, and later
replicating regions being transcriptionally silent and heterochromatic (reviewed in
(Gilbert, 2002; Gondor and Ohlsson, 2009; MacAlpine and Bell, 2005; Mechali, 2010;
Weinreich et al., 2004)). These correlations suggest that origins may be subject to similar
modes of regulation by local chromatin structure as promoters. Indeed, transcription
factors can stimulate origin activity (Chang et al., 2004; Danis et al., 2004; Marahrens and
Stillman, 1992), and active origins frequently co-localize with transcription start sites of
active genes in Drosophila and mammalian cells (Cadoret et al., 2008; Karnani et al., 2010;
MacAlpine et al., 2010; Sequeira-Mendes et al., 2009). The role of transcription factors here
is thought to involve the recruitment of chromatin remodelers or modifiers that position
nucleosomes or otherwise increase accessibility of origins to trans-acting factors (Flanagan
and Peterson, 1999; Hu et al., 1999; Li et al., 1998; Lipford and Bell, 2001). Similar to their
effects on transcription, local histone deacetylation typically delays or suppresses origin
19

firing, whereas histone acetylation advances or stimulates origin activity (Aggarwal and
Calvi, 2004; Aparicio et al., 2004; Goren et al., 2008; Knott et al., 2009c; Pappas et al., 2004;
Stevenson and Gottschling, 1999; Vogelauer et al., 2002; Weber et al., 2008). However,
distinct aspects of chromatin structure may affect origin timing versus efficiency. Recent
studies indicate that histone acetylation is required for pre-RC assembly (Miotto and
Struhl, 2007), and multiple, acetylated lysines in histone H3 and H4 N-termini are required
for efficient origin activity (Eaton et al., 2011; Unnikrishnan et al., 2010). The mechanism
of temporal control is less clear. Early firing is thought to represent a default state, with
deacetylated chromatin imposing a delay.

Recently, we reported that the Rpd3L histone deacetylase delays the activation of ~100
origins throughout the yeast genome (~1/3 of the active origins) (Knott et al., 2009c).
With this dataset we used classification-regression trees to identify annotated protein
binding-sites (from (Harbison et al., 2004) whose presence or absence near origins was
predictive of origin regulation by Rpd3L. This and further analysis identified binding sites
of Forkhead transcription factors, Fkh1 and Fkh2, as being depleted near Rpd3L-regulated
origins (data not shown). Fkh1 and Fkh2 have been well characterized for their role in
regulating G2/M-phase specific transcription of a group of genes known as the CLB2 cluster
(reviewed in (Murakami et al., 2010)), but have no known role in DNA replication. In this
study, we show that Fkh1 and Fkh2 regulate the initiation timing of most of the earliest
origins in the yeast genome through a novel mechanism involving origin clustering in G1-
phase.
20

2.2 Results

Fkh1 and Fkh2 control genome-wide initiation dynamics of replication origins: To
test whether Fkh1 and Fkh2 influence replication origin function, we examined genome-
wide origin-firing using BrdU immunoprecipitation analyzed by DNA sequencing (BrdU-IP-
Seq), in cells arrested in early S-phase with hydroxyurea (HU). In this analysis, BrdU peak
size is proportional to origin efficiency in HU: early-efficient origins produce large peaks
while late and/or dormant origins yield smaller or no peaks (Knott et al., 2009c). Because
Fkh1 and Fkh2 play partially complementary, yet opposing roles in regulation of G2/M-
phase regulated genes (Murakami et al., 2010), we analyzed single as well as double
deletion mutants of FKH1 and FKH2. Furthermore, because the double mutant cells exhibit
slow, pseudohyphal growth, which complicates their analysis, we also examined these cells
with over-expression of C-terminally truncated FKH2 (+pfkh2∆C), which largely restores
CLB2 cluster gene regulation (Reynolds et al., 2003). Consistent with this, we found that
expression of Fkh2∆C in fkh1∆ fkh2∆ cells suppressed their pseudohyphal growth and
restored nearly normal growth rate (Fig. S1.1A and data not shown).
21

Figure S1.1, related to Figure 1.1A. Suppression of pseudohyphal growth of fkh1∆ fkh2∆
cells by expression of Fkh2∆C. Phase-contrast images of the indicated strains grown in
liquid culture and sonicated mildly to disrupt cell aggregates. B. Origins deregulated in
fkh1∆, fkh1∆ fkh2∆, and fkh1∆ fkh2∆ + fkh2∆C cells. Venn diagrams showing overlap of
deregulated origins identified as Fkh-activated and Fkh-repressed.

In wild-type (WT) cells, 295 peaks of BrdU incorporation were detected genome-wide (Fig.
1.1A and Data S1). Combined deletion of FKH1 and FKH2 had an unprecedented effect on
origin activity throughout the genome, with the activities of the archetypal early origins
ARS305 and ARS607 being strongly reduced (Fig. 1.1A). Genome-wide, of the 352 origins
22

that were detected to fire in WT and/or fkh1∆ fkh2∆ cells, 106 (30%) origins were
significantly decreased in activity (Fkh-activated) and 82 (23%) were significantly
increased (Fkh-repressed) (Table S1 and Data S1). Deletion of FKH1 significantly
(FDR<0.005) altered the activity of specific origins, with 35 being Fkh-activated and 16
Fkh-repressed, whereas deletion of FKH2 had no significant effect on the replication
pattern (Fig. 1.1A, S1.1B, C, Table S1 and Data S1). Fortuitously, expression of fkh2∆C,
while complementing the pseudohyphal growth defects due to transcriptional
deregulation, did not complement the origin deregulation of fkh1∆ fkh2∆ cells, with
virtually all of the same origins being identified as Fkh-activated (95) or Fkh-repressed
(80) (Fig. 1.1A, S1.1B, C, Table S1 and Data S1). This result demonstrates that the C-
terminus of Fkh2 is required for origin regulation, and suggests that the effects on origins
are independent of transcriptional regulation by Fkh1 and Fkh2. We took advantage of the
ability of fkh2∆C expression to complement the transcriptional defects, but not the
replication defects, and to improve the growth of the double mutant cells to facilitate
further analyses of fkh1∆ fkh2∆ cells.

Two-dimensional clustering of the Fkh-regulated origins based on their peak sizes allows a
global comparison of origin activities in the WT, single and double mutant strains. This
analysis reveals the extensive deregulation of fkh1∆ fkh2∆ and fkh1∆ fkh2∆ +pfkh2∆C cells,
the strong similarity between replication patterns in the WT and fkh2∆ cells, and the
intermediate phenotype of fkh1∆ cells (Fig. 1.1B). These data indicate that Fkh1 and Fkh2
play a major and complementary role in selecting certain origins for early activation, while
23

repressing the activation of others. Fkh1 is sufficient to maintain normal (early) origin
regulation in the absence of Fkh2, whereas Fkh2 only partially compensates for the
absence of Fkh1.

Figure 1.1. Analysis of early S-phase BrdU incorporation. A. BrdU incorporation plots of
chromosomes III and VI are shown; plot colors and symbols correspond to the strain key
above. Origins discussed in the text are boxed. B. Two-dimensional clustering of peak
counts at Fkh-regulated origins is shown; columns (color-keyed above) correspond to
strains and rows to origins. C. All detected origins (in rows) are arranged from maximum
to minimum counts in WT, with the positions of Fkh-regulated origins indicated. See also
Figure S1.1, Table S1 and Data S1.To appraise the global relationship between origin
activities and regulation by Fkh1 and/or Fkh2 (Fkh1/2), we arranged origins according to
24

their WT activity levels (in HU) and plotted the positions of Fkh-activated and -repressed
origins (Fig. 1.1C). Fkh-activated origins were strongly enriched among earlier-firing
origins while Fkh-repressed origins were strongly enriched among later-firing (or
inefficient) origins (p<0.001, hypergeometric test). These results show that Fkh1 and Fkh2
are largely responsible for differential origin firing dynamics throughout the genome.

To examine in more detail the effect of Fkh1 and Fkh2 on temporal origin-firing dynamics,
we analyzed replication throughout an unperturbed, synchronous S-phase. Total DNA
content analysis showed similar overall replication kinetics in WT and fkh1∆ fkh2∆
+pfkh2∆C cells (hereon fkh1∆ fkh2∆C) (Fig. S1.2A). We next used BrdU pulse labeling
combined with BrdU-IP analyzed by microarray (BrdU-IP-chip) to analyze origin-firing
dynamics. At Fkh-activated ARS305 in WT cells, substantial BrdU incorporation occurred
during the 12-24min through 30-42min pulses, and ceased by the 36-48min pulse,
consistent with the early and synchronous replication of this origin (Fig. 1.2A). In fkh1∆
fkh2∆C cells, however, BrdU incorporation at ARS305 was delayed and reduced in
comparison, occurring mainly after replication had ceased in the WT (Fig. 1.2A). ARS607
and numerous other early origins showed similar delay of activity in fkh1∆ fkh2∆C cells
(Data S2). These data confirm the results of the analysis with HU and demonstrate that
Fkh1/2 are required for the early activation of many origins throughout the yeast genome.

25

Figure S1.2, related to Figure 1.2A. FACScan analysis of DNA content of WT and fkh1∆
fkh2∆C cells synchronized in G1-phase with –factor and released synchronously into S-
phase. B. Two-dimensional gel electrophoresis analysis of ARS305 (Fkh-activated) and
ARS1520 (Fkh-repressed) in unsynchronized WT and fkh1∆ fkh2∆C cells. Genomic DNA
was digested with NcoI and SalI. C. Non-random distribution of Fkh-regulated origins.
Chromosomal positions of Fhk-activated and –repressed origins are plotted. D. Histogram
displaying the frequency of “Cut” counts observed in the 10
5
simulations as well as the
experimentally observed “Cut” count. “Cuts” refers to the number of times a Fkh-activated
origin is followed by a Fkh-repressed origin, or vice-versa, given a random distribution (see
Methods).

The data also indicate that Fkh1/2 normally repress the earlier firing of many origins (Data
S2). For example, examination of the late-replicating region of chromosome XV
demonstrates that several later-firing origins, such as ARS1520, initiated replication earlier
26

in the mutant cells (Fig. 1.2A). To address the formal possibility that the observed
differences in origin activation timing derive from a change in origin activation efficiency,
we performed two-dimensional gel electrophoresis analysis of replication initiation
structures of Fkh-activated origin ARS305 and Fkh-repressed origin ARS1520. Both origins
exhibit high efficiency in both WT and fkh1∆ fkh2∆C cells (Fig. S1.2B). These data confirm
that that Fkh1/2 establish the temporal program of origin activation.
For a global view of the impact of Fkh1/2 regulation on the temporal program, we
clustered the Fkh-regulated origins according to their peak-count differences in the HU
analysis, and plotted the differences in their levels of BrdU-incorporation between WT and
mutant for each interval in the time-course (Fig. 1.2B). This analysis shows global
correspondence between the change in origin activity in HU and the change in origin
activity in the time course in the fkh1∆ fkh2∆C cells, with Fkh-activated origins firing earlier
and Fkh-repressed origins firing later in WT cells. Thus, Fkh1/2 play a major role in
determining the characteristic firing times of replication origins throughout much of the
yeast genome.

27

Figure 1.2. Temporal analysis of DNA replication by BrdU pulse-labeling. A. BrdU
incorporation plots of chromosome III and a region of XV are shown. Origins discussed in
the text are boxed. B. The matrix shows differences (WT-fkh1∆ fkh2∆C) in BrdU
incorporation (  M-value) at all Fkh-regulated origins (columns) across time (rows); the
origins are arranged from left to right by their differences (WT-fkh1∆ fkh2∆C) in BrdU
incorporation in HU (  HU Counts). Specific origins are indicated below. See also Figure
S1.2 and Data S2.

Fkh-regulation involves establishment of replication timing domains: Comparison of
the WT and mutant chromosomal replication profiles reveals additional features of
interest, including even earlier replication of centromere (CEN)-proximal sequences, such
that these became the earliest replicating region of each chromosome (Fig. 1.2A and Data
S1.2). Plotting CEN-proximal origins (ie, within 25kb) in the time-course clustergram
shows that many of these origins initiated earlier in the mutant cells and were among the
28

most strongly affected of the Fkh-repressed origins (Fig. 1.2B). Another striking feature of
the mutant replication profiles is the delayed replication of most telomere (TEL)-proximal
sequences (Data S2), particularly those with active origins, as evident on the right arm of
chromosome III (Fig. 1.2A). These results further demonstrate the global role of Fkh1/2 in
determining genome replication timing and suggest a function in chromosomal
organization.

We wondered whether the distribution of Fkh-regulated origins along chromosomes might
provide additional clues about their functional organization. Chromosomal plots of Fkh-
regulated origins (ignoring non-regulated origins) show frequent, linearly contiguous
groups of Fkh-activated and -repressed origins, suggesting a non-random distribution (Fig.
S1.2C). To test this notion rigorously, we applied a permutation test that determines the
likelihood that the contiguous groups are random. The result shows that the distribution of
Fkh-activated and -repressed origins is non-random and that origins of each class
frequently cluster linearly along the chromosome with other members of their class
(p<0.01, Fig. S1.2D). Together with the CEN- and TEL-specific effects, these results are
consistent with Fkh1/2 establishing domains of replication timing.

Fkh1/2 bind and function in cis to Fkh-activated origins: Fkh1 and Fkh2 exhibit similar
DNA sequence binding specificities in vitro and bind extensively throughout the genome,
with significant overlap of binding sites (data not shown and (Harbison et al., 2004;
Hollenhorst et al., 2001; MacIsaac et al., 2006). To examine the relationship of Fkh1 and
29

Fkh2 binding with origin regulation, we analyzed the distribution of putative Fkh1 and
Fkh2 binding sites within 500bp of Fkh-activated, -repressed, and -unregulated origins (see
Methods). This analysis shows that Fkh1 and Fkh2 binding sites are enriched near Fkh-
activated origins and depleted near Fkh-repressed origins (Fig. 1.3A, B, hypergeometric
test, p<0.01), as expected if Fkh1/2 act through direct binding near Fkh-activated origins.
Fkh1 was most enriched, being ~four-fold enriched at Fkh-activated versus -repressed
origins, consistent with a predominant role for Fkh1 rather than Fkh2 in origin regulation
as indicated by the single mutant analysis above.

The enrichment of Fkh1/2 binding sites near origins may explain the selection of these
origins for early activation, however, Fkh1/2 bind near some origins that are not Fkh-
activated suggesting that Fkh1/2 binding in the vicinity is not sufficient for origin
activation. To determine more precisely how Fkh1 and Fkh2 localize in relation to Fkh-
regulated origins, we calculated the distance from each origin’s ARS-consensus sequence
(ACS), which binds ORC, to the likeliest Fkh1 and Fkh2 binding site within 500bp and
plotted the results as a frequency distribution (see Methods). The distribution reveals
extraordinary proximity of Fkh1 and Fkh2 consensus sites to ACSs of Fkh-activated origins,
with frequent overlap of the Fkh1/2 binding sites and ACSs (Fig. 1.3C). In contrast, Fkh1
and especially Fkh2 showed poorer alignment and binding density with those few Fkh-
repressed origins proximal to Fkh1/2 binding sites. These results suggest that the
positioning and/or number of these sites may be important for origin regulation

30

To test directly whether Fkh1/2 regulate origin function through binding in cis to the
affected origin, we mutated two putative Fkh1/2 binding sites near ARS305 (ars305∆2BS).
Combined mutation of these sites significantly reduced BrdU incorporation at ARS305, but
not at more distal origins, indicating that Fkh1/2 regulate ARS305 directly through binding
in cis (Fig. 1.3D). Crucially, mutation of these binding sites eliminated Fkh1 binding to the
ARS305 region without eliminating ORC binding (Fig. 1.3D). These results also eliminate
concerns that origin deregulation results from mis-expression of a replication factor(s) in
fkh1∆ fkh2∆C cells. Overall, these results demonstrate that Fkh1/2 binding positively
influences origin activity.

31

Figure 1.3. Analysis of Fkh1 and Fkh2 binding sites near origins. A and B. Frequencies of
expected and actual Fkh1 (A) and Fkh2 (B) consensus binding sites near Fkh-activated,
Fkh-unregulated, and Fkh-repressed origins are shown. C. Frequency distribution plots of
Fkh1 and Fkh2 consensus binding sites relative to ACS position are shown. D. M-values for
BrdU-IP-chip and for ChIP-chip of Fkh1 and ORC binding along the ARS305 region in WT
cells harboring ARS305 or ars305∆2BS.

Fkh-dependent origin regulation is not correlated with transcription levels or
changes: The notion of a mechanistic link between replication origin timing and
transcriptional state, together with the well-characterized roles of Fkh1 and Fkh2 as
transcriptional regulators, suggested that altered transcription, particularly of genes
proximal to Fkh-regulated origins, might explain the altered origin firing. Although
32

expression of Fkh2∆C suppressed pseudohyphal growth, indicating that normal
transcriptional regulation had been at least partially restored, we nonetheless wished to
determine whether differences in transcription of genes proximal to the affected origins
could account for the differences in origin activity. Accordingly, we analyzed global RNA
transcript levels using strand-specific RNA quantification by sequencing (RNA-Seq) and
RNA Polymerase II (Pol II) occupancy using chromatin immunoprecipitation analyzed by
sequencing (ChIP-Seq) of the Pol II core subunit Rpb3 in WT and fkh1∆ fkh2∆C cells, in
unsynchronized cells and cells synchronized in G1-phase, when replication timing is
established (Dimitrova and Gilbert, 1999; Raghuraman et al., 1997). Up-regulation of CLB2
in G1-phase fkh1∆ fkh2∆C cells, which is consistent with the role of Fkh1 in CLB2
repression, and significant overlap between genes identified by the different methods
validated both analyses (Table S2). A permutation test indicates that genes deregulated in
fkh1∆ fkh2∆C cells are not significantly co-localized with or proximal to Fkh-regulated
origins (see Methods). We also plotted RNA transcript levels and Rpb3 occupancy, as well
as their differences in fkh1∆ fkh2∆C cells, within 10kb of Fkh-regulated origins (Fig. 1.4).
Visual inspection of these plots show no obvious correlation with the effects on origin
activities, regardless of the magnitude or directionality (positive or negative) of effect, the
orientation of the immediately flanking genes, or the cell cycle stage. Linear regression
analysis also shows no consistent correlation between the effects on origin activity and the
expression levels of the immediately flanking genes (see Methods). These findings
demonstrate that origin regulation by Fkh1/2 does not involve proximal changes in
transcription.
33

Figure 1.4. Transcription analysis surrounding Fkh-regulated origins in unsynchronized
and G1-synchronized cells. RNA-Seq (A) and Rpb3 ChIP-Seq (B) read counts of WT, fkh1∆
fkh2∆C, and WT-fkh1∆ fkh2∆C differences ( ), within 10kb of each Fkh-regulated origin, are
aligned by each origin’s predicted or verified ACS. Origins are grouped according to the
orientation of the flanking genes, and arranged by differences (WT-fkh1∆ fkh2∆C) in BrdU
incorporation in HU (  HU Counts). See also Table S2.

Cdc45 preferentially associates with Fkh-activated origins in G1-phase: We wondered
whether Fkh1/2 regulate replication timing by modulating the binding of replication
factors to origins. To determine whether Fkh1/2 influence ORC binding or MCM loading,
34

we used ChIP analyzed by microarray (ChIP-chip) to examine ORC binding in
unsynchronized cells and Mcm2+4 binding in G1-synchronized cells. The results show no
significant, global difference in ORC or Mcm2+4 origin-binding between WT and fkh1∆
fkh2∆C cells (Fig. 1.5A and Table S3), contrary to the idea that Fkh1/2 affect origin-firing by
modulating ORC or MCM binding.

Origin initiation requires the DDK-dependent recruitment of Cdc45 to pre-RCs. However,
Cdc45 associates specifically, albeit relatively weakly, with several early replication origins
in G1-phase (prior to DDK activation), presaging their characteristic early S-phase activity
(Aparicio et al., 1999). This suggests that these origins gain an early advantage (by G1-
phase) in their ability to recruit Cdc45 to enable early initiation. Examination of Cdc45
binding by ChIP-chip shows Cdc45 association with many early origins, including Fkh-
activated origins, such as ARS305 and ARS607, and a number of CEN-proximal origins (Fig.
5A, B, and Table S3). Of 28 origins that bind Cdc45 in WT G1-phase cells, 15 are Fkh-
activated and 14 are CEN-proximal (on 11 CENs), while only one is Fkh-repressed.
Strikingly, in the fkh1∆ fkh2∆C cells, Cdc45 binding is lost from the Fkh-activated origins,
which become significantly later firing, leaving only 13 origins binding Cdc45 (Fig. 5B and
Table S3). Of these 13, 12 are CEN-proximal, which as shown above, remain early firing.
Thus, Cdc45 origin-binding in G1-phase is robustly associated with early initiation. These
findings support the idea that Fkh1/2 influence origin function by regulating access to the
pool of replication factors such as Cdc45, whereas CEN-proximal origins have access to
Cdc45 independently of Fkh1/2.
35

Figure 1.5. Genome-wide binding of replication initiation factors to Fkh-regulated origins.
A. M-values from ChIP-chip analysis of ORC, Mcm2+4, and Cdc45 at Fkh-regulated origins
(in rows) are arranged by differences (WT-fkh1∆ fkh2∆C) in BrdU incorporation in HU ( 
HU Counts). B. Venn diagram of Cdc45 binding within different origin classes is shown.
See also Table S3.
36

Fkh1/2 are required for selective clustering of Fkh-activated origins in G1-phase:
The organization of selected origins into subnuclear domains or replication foci by Fkh1/2
may explain their preferential access to limiting or sequestered initiation factors like
Cdc45. In accord with this, a global analysis of intra- and inter-chromosomal interactions
of the yeast genome using a variation of 4C (Chromosome Conformation Capture-on-Chip)
suggests that early origins cluster in G1-phase (Duan et al., 2010). We analyzed this origin
interaction data to determine whether origin clustering was associated with Fkh-regulation
and/or Cdc45 binding in G1-phase. Two-dimensional clustering based on origin
interaction frequencies resulted in two main clusters of interacting origins, with 89 and 92
origins, respectively (Fig. 1.6A). One cluster contains most of the Cdc45-bound origins, the
most statistically significant Fkh-activated origins, and CEN-proximal origins. This cluster
also contains earlier-firing origins on average than the other main cluster and is depleted of
non-CEN proximal, Fkh-repressed origins (hypergeometric test, p<0.005). These findings
suggest that Fkh-regulation involves selective origin clustering.
37

Figure 1.6. Chromosome-conformation capture analyses of origin interactions. A. Two-
dimensional clustering of origin-origin interaction frequencies is shown, with origins in
columns and rows of the matrix. Columns to the right indicate Cdc45 ChIP-chip binding,
average BrdU ∆HU-counts, and ∆BrdU-pulse M-values. The top 5% (based on p values) of
Fkh-activated and Fkh-repressed origins are indicated. B. Venn diagram of overlap
between experimental replicates is shown. C. Plots of the ARS607 region including relevant
XbaI sites are shown. See also Figure S1.3 and Table S4.

To test whether Fkh1/2 have a role in origin clustering, we used 4C to analyze the trans
associations of Fkh-activated origin ARS305 with other genomic sequences (for scheme, see
Fig. S1.3A). We validated this analysis by comparing overlap between experimental
38

replicates of WT and mutant cells, with and without crosslinking, and by analyzing the
number of intra- versus inter-chromosomal interactions detected (Fig. S1.3B). As expected,
and consistent with the results of (Duan et al., 2010), intrachromosomal interactions were
enriched versus interchromosomal interactions (p<0.001). We detected 48 ARS305-
interacting loci in both WT replicates (of 71 and 72 in the replicates), and 41 ARS305-
interacting loci in both fkh1∆ fkh2∆C replicates (of 164 and 189 in the replicates) (Fig. 1.6B
and Table S4). The larger number of detected interactions with lower overlap between
them in the fkh1∆ fkh2∆C replicates is consistent with a decrease in specificity of ARS305
interactions in the mutant cells. Most of the 48 sites in WT cells were not detected in the
mutant cells, indicating that their interaction with ARS305 is Fkh1/2-dependent. For
example, ARS305 interacted with ARS607 (as shown previously (Duan et al., 2010)) in both
WT replicates and in neither fkh1∆ fkh2∆C replicate (Fig. 1.6C), indicating that Fkh1/2 are
required for interaction in G1-phase between these early-firing, Fkh-activated origins.
These results indicate that Fkh1/2 play a role in determining the long-range chromatin
contacts made by ARS305, and support the idea that Fkh1/2 function in origin regulation
through origin clustering.

39

Figure S1.3, related to Figure 1.6. 4C analysis of ARS305 interactions. A. Scheme of the
4C method showing relevant XbaI (X1-X4) and MseI (M1-M4) restriction sites surrounding
ARS305 (Bait) and a hypothetical interacting locus (Prey), and primers (P1-P4) used to
amplify captured loci for identification by microarray. The tethering agent represents
cross-linked protein(s) mediating interaction between the bait and prey. B. Statistical
analysis of ARS305 interacting sites by chromosome showing the expected preference for
intrachromosomal interactions (i.e., with chromosome III). The p value is based on the
number of observed versus expected interactions for each chromosome (the expected
number of interactions is directly proportional to the number of XbaI fragments per
individual chromosome).
40

Fkh1 and Fkh2 interact with ORC: The binding of Fkh1/2 adjacent to many Fkh-activated
origins, including ARS305 and ARS607 (data not shown and (Harbison et al., 2004; Keich et
al., 2008)), led us to hypothesize that Fkh1/2 bound near origins might stabilize origin
contacts in trans through interaction with ORC bound at other Fkh-activated origins.
Immunoprecipitation (IP) of Myc-tagged Fkh1 or Fkh2 from soluble cell extracts resulted in
co-precipitation of ORC (Fig. 1.7A, lanes 1 and 2, data not shown for Fkh2); Orc2 was
robustly detected, Orc1 and Orc3 were weakly detected, and Orc4-Orc6 were obscured by
co-migrating immunoglobulin heavy chain (data not shown). Reciprocal IP of ORC using a
polyclonal antibody co-precipitated Fkh1 (Fig. 1.7B, lanes 3 and 4). Taken together, these
results demonstrate a physical interaction (direct or indirect) between ORC and Fkh1.
These interactions persisted in the presence of the DNA-intercalating agent, ethidium
bromide, indicating that the interactions are likely not DNA-mediated (Fig. 1.7C, lanes 5-8).
Together with the close proximity of Fkh1/2 binding sites with origin ACSs, these results
support the idea that Fkh1/2 interact with ORC to bridge replication origins in trans.

41

Figure 1.7. Co-IP of Fkh1 with ORC. Soluble extracts from FKH1-MYC (lanes 1, 3, 5-8) and
untagged (lanes 2, 4) cells were subjected to IP with anti-Myc antibody (A) and anti-ORC
antibody (B and C). IPs were analyzed by immunoblotting with anti-Myc and anti-ORC
antibodies. C. Ethidium bromide (EtBr) was included in the IPs at 10, 40, and 100 µg/mL in
lanes 6, 7, and 8, respectively. ORC protein was included as standard.

42

2.3 Discussion

Fkh1/2 establish replication-timing domains through origin clustering: Our findings
reveal a novel, global mechanism for the regulation of origin initiation timing, involving the
spatial organization of replication origins by Fkh1/2. Previous studies have concluded that
yeast origins are early by default, and that late timing is imposed by flanking sequences of a
repressive nature (Ferguson and Fangman, 1992; Friedman et al., 1996). However, our
findings show that Fkh1/2 actively program the timing of most of the earliest origins
throughout the genome. Thus, we propose that Fkh1/2 establish early replication timing at
Forkhead-activated origins by recruiting these origins into clusters where Cdc45 is (and
likely other replication factors are) concentrated. The enrichment and distinct positioning
relative to the ACS of Fkh1/2 binding sites likely explains the selective preference for Fkh-
activated origins. Clustering may involve interaction of Fkh1/2 bound adjacent to an origin
with ORC bound to a distal, second origin. Likewise, Fkh1/2 bound near the second origin
might interact with a third origin, and so forth, providing a mechanism to cluster several
origins together. This congregation of origins and initiation factors provides a kinetic
advantage in assembling the factors needed for replication initiation upon S-phase entry,
which transforms these origin clusters into early replication factories. The ensuing
dynamics of the replication process, involving spooling of DNA through the replication
factories (Kitamura et al., 2006), eventually repositions more distal, unfired origins,
bringing them in proximity of the concentration of the replication factor(ie)s and thereby
allowing them to gain access as early replicons terminate and are released. This is
expected to result in an increasingly stochastic pattern of replication initiation as S-phase
43

proceeds and many unfired origins compete for limited access. However, later-replicating
regions also exhibit well-defined replication patterns indicating preferred origin timing
and usage. Indeed, chromosomes IV, XII, XIV, and XV each have distinctly late-replicating
regions >200kb in length, encompassing groups of contiguous Fkh-repressed origins,
which lose this unique character in the absence of Fkh1/2 (Fig. 1.2A and Data S2).

Origin clusters may define replication-timing domains. The organization of mammalian
chromosomes into spatial domains correlates strongly with replication timing (Ryba et al.,
2010). Analysis of global 4C in yeast shows clustering of early origins (in G1-phase), and
we have now shown that the early origin cluster contains Fkh-activated and Cdc45-bound
origins (in G1-phase). We have confirmed that Fkh-activated origins ARS305 and ARS607
interact in trans, and critically, show that this interaction depends on Fkh1/2. In addition,
Fkh-activated and Fkh-repressed origins often occur in separate, linearly contiguous
groups along chromosomes, suggesting the formation of distinct domains. This may
involve the anchoring of intrachromosomal chromatin loops by Fkh1/2 bound near origins,
perhaps through interaction with ORC, particularly in the case of Fkh-activated origins,
which are enriched for Fkh1/2 binding. In the case of Fkh-repressed origins, a dearth of
Fkh1/2 binding sites presumably reduces the likelihood that these origins join the Fkh-
activated clusters, which may permit other mechanisms, such as deacetylation or
localization to the nuclear periphery, to define replication timing of these regions.
Alternatively, the later timing may be a consequence of conformational or spatial
44

constraints imposed by the chromosomal architecture established by Fkh1/2 clustering of
Fkh-activated origins.

In the absence of Fkh1 and Fkh2, CEN-proximal origins dominate the early replication
landscape, suggesting that CENs confer early replication intrinsically. CENs normally
cluster and occupy a characteristic interior position in the nucleus (Jin et al., 1998) that we
suggest overlaps with the pool of replication factor(ie)s. Consequently, CEN-proximal
origins have favorable access to this pool and initiate early, independently of Fkh1/2. Thus,
CEN-proximal origins may act as organizing sites for early-replicating origin clusters that
include non-CEN-proximal origins. More distal Fkh-activated origins may utilize Fkh1/2 to
cluster with CEN-proximal origins, thereby drawing these more distal origins into the pool.
This is consistent with the finding that CEN-proximal origins localize to the large, early-
replicating cluster in the global 4C data together with the earliest Fkh-activated origins.
Thus, the advanced replication timing of CEN-proximal origins (and perhaps other Fkh-
repressed origins) in cells lacking Fkh1/2 may result from reduced competition from Fkh-
activated origins for limiting replication factor(ie)s, rather than a direct repressive function
of Fkh1/2. Incidentally, CEN-proximity may explain the finding in yeast that plasmid-borne
origins typically replicate early, as these studies were performed with CEN-harboring
plasmids (Ferguson and Fangman, 1992; Friedman et al., 1996).

In contrast to CENs, TELs form several clusters that occupy the nuclear periphery (Gotta et
al., 1996; Heun et al., 2001). The normally late replication of TEL-proximal regions is
45

consistent with the notion that the dynamic nature of the replication process eventually
relocates these distal regions to the interior of the nucleus, which ultimately enables their
access to replication factor(ie)s. In the absence of Fkh1 and Fkh2, most of the active
telomeric origins are further delayed. We imagine that the delayed activation of Fkh-
activated origins located along distal chromosomal arms results in a corresponding delay
in the relocation to TEL-proximal origins to the vicinity of replication factor(ie)s.
Alternatively, Fkh1/2 may act directly to regulate TEL-proximal origins. Further study will
be required to understand the regulation of CEN- and TEL-proximal origin timing.

Multiple, separable roles for Fkh1 and Fkh2 in regulation of the genome: A clear finding of
this study is the mechanistic independence of Fkh-origin regulation from transcription.
There is no correlation between the observed changes in replication timing and
transcriptional levels of proximal genes. Importantly, expression of Fkh2 lacking its C-
terminus in fkh1∆ fkh2∆ cells significantly restores transcriptional regulation of CLB2
cluster genes (only CLB2 remained deregulated and only in G1-phase cells) without
restoring origin regulation, directly demonstrating a separation of these Fkh1/2 functions.
Nevertheless, our results do not rule out the possibility that the function of Fkh1/2 in
origin clustering may also underlie transcriptional control not elicited under our growth
conditions.

As transcriptional regulators, Fkh1 and Fkh2 exhibit opposing, as well as partially
complementary functions (Murakami et al., 2010). Fkh1 and Fkh2 also demonstrate
46

distinct abilities to regulate origins, suggesting that the features that distinguish Fkh1 and
Fkh2 functions in transcription also impinge on their functions as origin regulators.
Whereas Fkh2 plays the lead role in transcriptional regulation, Fkh1 plays the lead role in
origin regulation. Fkh1 differs from Fkh2 most significantly in the presence of a C-terminal
extension in Fkh2, which regulates its interaction(s) with transcriptional co-activator(s)
(Darieva et al., 2010; Darieva et al., 2003; Koranda et al., 2000; Pic-Taylor et al., 2004;
Reynolds et al., 2003). This domain is also required for Fkh2’s function in origin regulation,
suggesting that proper regulation of co-activator interactions is critical, and that factors
interacting with Fkh2 but not with Fkh1 may disrupt origin regulation. Mcm1, which binds
cooperatively with Fkh2, but not Fkh1 (Boros et al., 2003; Koranda et al., 2000; Kumar et
al., 2000; Pic et al., 2000), is an intriguing candidate, as it has been reported to modulate
origin function (Chang et al., 2004). We note that Mcm1 binding sites are not enriched near
Fkh-activated origins (data not shown). Thus, consistent with the lack of effect on origin
firing of FKH2 deletion, it is possible that Fkh2 normally plays no role in origin regulation,
and only substitutes (partially) in Fkh1’s absence.

Fkh1, but not Fkh2, also regulates donor preference in yeast mating-type switching (Sun et
al., 2002). Mating-type switching involves homologous recombination between the MAT
locus (recipient) and one of two silent mating-type loci (donor) distally located on opposite
arms of the same chromosome, HML  and HMRa. This mechanism presumably
necessitates chromosomal looping of either arm to juxtapose the donor and recipient loci.
Remarkably, in MATa cells, HML  is preferentially selected as the donor in over 90% of
47

cells, which ensures efficient mating-type switching. This preference depends on Fkh1
binding to the recombination enhancer (RE), which is proximal to HML . Our finding that
Fkh1/2 mediate long-range origin interactions suggest that Fkh1 mediates a stable, long-
range interaction between MATa and the RE to specify the recombination between MATa
and HML , which conspicuously, like early origin clustering, occurs during G1-phase. The
role of Fkh1 in regulating recombination over long distances together with Fkh1/2’s role in
regulating replication initiation timing through long-range origin clustering suggests that
establishing long-range chromatin contacts may be a common mechanism of Fkh1/2
function, likely extending to transcriptional control.

Our proposed mechanism of origin clustering may also explain how the long-range
interaction necessary for recombinational donor preference is established. Dormant
origins are closely associated with the RE (ARS304) and with MAT (ARS313 and ARS314).
Thus, interactions between Fkh1 bound to the RE and ORC bound to the distal ARS313 or
ARS314 may stabilize long-range contacts between these loci; similar interactions between
ORC bound to ARS304 and Fkh1 bound near MAT may also participate (though an RE-like
element has not been identified near MAT). The dormancy of these origins is consistent
with the idea that these loci form a separate chromosomal domain dedicated for
recombination, which delays replication (by inhibiting initiation and allowing passive
replication from distal, flanking origins). Exactly how such domains are dedicated to one
function over another will require more investigation, but may reflect combinatorial
48

regulation by Fkh1/2 together with other factors, along with defined sub-nuclear
localization of these activities.

The findings presented here provide a clearer understanding of the epigenetic basis for
differential origin regulation and its connection to the spatial organization of
chromosomes. Rather than a direct connection with transcription, the results indicate that
the organization of origins into functional clusters determines their activation kinetics. Our
study identifies Fkh1 and Fkh2 as factors that participate in the establishment of the three-
dimensional structure of the yeast genome and the epigenetic regulation of genome
replication. This regulation through structure may be analogous to epigenetic mechanisms
of transcriptional memory wherein gene looping or sub-nuclear localization is correlated
with the maintenance of a transcriptional state or a potentiated state primed for rapid
response (Misteli, 2007). Furthermore, this organization may contribute to a coordination
of replication and transcription, perhaps with consequence for genome stability (Knott et
al., 2009a). Indeed, this study’s findings provide a new handle to investigate the
consequences of deregulating replication timing on gene regulation or genome stability.
The identification of yeast members of the conserved Fox transcription factor family as
physical mediators of chromosomal architecture and epigenetic regulation suggest
conservation of this function, which may link replication timing control and the role of Fox
proteins in metazoan development.

2.4 Experimental Procedures

49

Additional details are provided below in the Extended Experimental Procedures
Yeast strains and methods
W303-derived, BrdU-incorporating strains were used for all strain constructions (Viggiani
and Aparicio, 2006). Cell cycle block-and-release, DNA content analysis, and two-
dimensional gel analysis have been described (Aparicio et al., 2004). Co-IP was performed
as described (Hu et al., 2008), except Dynabeads Protein G (Invitrogen) was used. BrdU-
labeled DNA was isolated as described (Viggiani et al., 2010); salmon sperm DNA was
omitted for sequencing. 80ng of BrdU-IPed DNA was prepared for single-end sequencing
by Illumina ChIP-Seq protocol or 10ng of BrdU-IPed DNA was prepared for hybridization to
microarrays as described (Viggiani et al., 2010). ChIP-chip was performed and analyzed as
described (Knott et al., 2009b; Viggiani et al., 2009). ChIP-Seq was performed identically
except that culture was scaled-up four-fold to generate 5-10ng of IP material for single-end
sequencing by Illumina ChIP-Seq protocol. RNA was isolated from 20mL cultures using
RiboPure Yeast Kit (Ambion). rRNA was depleted with Ribominus Beads (Invitrogen), and
purified RNA was prepared for strand-specific RNA-Seq as described in (Parkhomchuk et
al., 2009). We used a custom microarray design (Nimblegen) that tiles one ~60bp
oligonucleotide for every ~80bp of unique genomic sequence. For hybridization and
washing we followed Nimblegen protocols, and for image capture used an Axon 4100A
Scanner.

Preprocessing of sequence data
Sequencing was carried out with an Illumina GAII. BrdU-IP-Seq and ChIP-Seq were
analyzed with 36bp single-end reads, while RNA-Seq was analyzed with 36bp paired-end
50

reads. Reads were aligned to S. cerevisiae genome release r.64 with PerM (Chen et al.,
2009), allowing only unique matches with a maximum of two mismatches per end. BrdU-
IP-Seq and Rpb3 ChIP-Seq reads were binned into non-overlapping 50bp bins; bin-counts
were median-smoothed (1000bp and 500bp windows, respectively) and quantile-
normalized across all experiments. This smoothing step was repeated. For all other gene
expression analysis, each RNA-Seq read was assigned to a gene only when at least one of its
paired-ends was fully contained within the gene’s ORF and when the read’s orientation
corresponded to the gene’s orientation. Reads whose paired-ends mapped to two or more
genes were discarded. Gene read-counts were quantile-normalized prior to differential
expression analysis.

Analysis of linear clustering of Fkh-regulated origins
We performed Monte Carlo simulations to determine the likelihood of the observed level of
clustering between like-regulated origins (e.g. both Fkh-activated) along the chromosome
occurring by chance. In each simulation we randomly assigning (from 352 total origins) 95
origins as Fkh-activated and 80 as Fkh-repressed (on each simulation) and determined the
number of occurrences where two Fkh-repressed or –activated origins neighbored each
other. We then compared the observed level of such instances to the empirical distribution
obtained through simulations to calculate a p-value.

51

Analysis of Fkh1 and Fkh2 binding sites
To determine whether Fkh1 and Fkh2 are differentially bound at Fkh-regulated versus
Fkh-unregulated origins we used the Position Weight Matrices (PWMs) defined in
(Morozov and Siggia, 2007)) to identify all putative Fkh1/2 binding sites near origins
(PWM-score cutoff =5.5). We defined Fkh1/2-bound origins as those with a putative site
within 500bp of its BrdU-peak apex. To determine the distribution of Fkh1/2 binding sites
relative to ACSs, for each Fkh1/2-bound origin with a defined ACS, we calculated the
distance from the ACS to the highest scoring binding site (ACS locations from (Eaton et al.,
2010)); we applied a kernel density function to these distances to define the probability
curves.

Analysis of global 4C
226 origins whose defined regions (as listed in OriDB) were fully contained within an EcoRI
and a HindIII restriction fragment were analyzed. The restriction fragment interaction map
from (Duan et al., 2010) was used to build two-dimensional interaction matrices for each
restriction fragment set containing the 226 origins. The matrix value (0 to 4) represents
the interaction distance between two origin-containing restriction fragments defined in
(Duan et al., 2010). The two matrices were summed and the two-dimensional clustering
algorithm defined in (Duan et al., 2010) was applied. 17 clusters containing fewer than ten
origins each (45 total) were not analyzed further.

52

Extended Experimental Procedures

Yeast strain and plasmid constructions
W303-derived, BrdU-incorporating strains CVy43 (Mata ade2-1, bar1::hisG, can1-100, his3-
11,15, leu2-3,112, trp1-1, ura3-1::BrdU-Inc::URA3) or CVy63 (Mata ade2-1, bar1::hisG, can1-
100, his3-11,15, leu2-3,112, trp1-1, leu2::BrdU-Inc::LEU2) were the WT parents for all strain
constructions (Viggiani and Aparicio, 2006). FKH1 and FKH2 were deleted in CVy43 as
described (Longtine et al., 1998), yielding strains: ZOy1 (fkh1∆::kanMX6), CVy138
(fkh2∆::His3MX6), and CVy139 (fkh1∆::kanMX6 fkh2∆::His3MX6); only differences in
genotype from CVy43 are indicated. Plasmid pfkh2∆C contains a C-terminally truncated
NotI-KpnI fragment of FKH2 (truncated at the native KpnI site in FKH2, deleting amino acids
624-862; this maintains the entire DNA binding domain and all homology with Fkh1) into
pRS424 digested with the same enzymes; pfkh2∆C was transformed into CVy139 yielding
strain SKy1. CDC45-HA3 (LEU2) was introduced into strains CVy43 and CVy139 + pfkh2∆C
using p405-CDC45-HA/C as described (Aparicio et al., 1997), yielding strains CVy46 and
T2y3, respectively. FKH1-MYC9 replaced FKH1 in CVy138 using plasmid pTOPO-Fkh1-
Myc9, yielding strain ZOy22. pTOPO-Fkh1-Myc9 was constructed using Phusion High-
Fidelity PCR kit (New England Biolabs, M0530) to amplify FKH1-MYC9-TRP1 from genomic
DNA of strain Z1448 (Harbison et al., 2004), and inserting it into pCR2.1-TOPO vector
(Invitrogen).

Strain ARy23 containing mutations of two Fkh1/2 binding sites at ARS305 was constructed
by pop-in/pop-out of plasmid p306-ARS305-∆2BS into strain CVy63 and confirmed by
53

sequencing of PCR-amplified genomic DNA. Plasmid p306-ARS305∆2BS was constructed
as follows: Two ~1kb fragments covering ARS305 with overlapping ends were amplified
from genomic DNA (using primers: 5´-gtcaagcttggcaatgtcaagagcagagc with 5´-
gtcctcgaggaatacataacaaaaatataaaaacc for one fragment and 5´-
tgagaattcaggcatcagtttgatgttgg with 5´-gtcctcgaggtccctttaattttaggatatgaaaac for the second
fragment), digested with EcoRI +XhoI and with XhoI + HindIII, respectively, and three-way
ligated into pRS306 digested with EcoRI + HindIII. The XhoI site changes the first predicted
Fkh1/2 binding site (chr III coordinates 39,563-39,570) without deleting or inserting
additional sequence. The resulting plasmid, p306-ARS305∆1BS was sequenced to confirm
that only the desired sequence changes were introduced. This plasmid was mutagenized
using QuikChange Lightning Multi Site mutagenesis kit (Agilent# 210515-5) using primer
(5´-caaagaaaaaaatcttagctttaagaactacaaagtcctcgaggaataataaatcacaccggacagtacatg) to change
the second predicted Fkh1/2 binding site (chr III coordinates 39,483-39,490) to an XhoI
site without deleting or inserting additional sequence. The resulting plasmid p306-
ARS305∆2BS was sequenced to confirm that only the desired sequence changes were
introduced.

Antibody methods.
For BrdU and chromatin IPs we used: anti-BrdU at 1:1000 (GE Healthcare, RPN202), anti-
Fkh1 at 1:200 (Casey et al., 2008), anti-ORC at 1:500 (Wyrick et al., 2001), anti-Mcm2 at
1:50 (Santa Cruz Biotech., SC-6680), anti-Mcm4 at 1:50 (Santa Cruz Biotech., SC-33622),
anti-Ha 16B12 at 1:200 (Covance, MMS101R), and anti-Rpb3 at 1:500 (Neoclone, W0012).
54

We used anti-Myc 9E10 at 1:100 and 1:2000 (Covance, MMS150P), and anti-ORC at 1:100
and 1:1000, for co-IP and immunoblotting, respectively.

BrdU-IP-Seq analysis
To identify an initial set of peaks in each experiment, a set of apices (bins whose count was
higher than any neighboring bin within 500bp) were detected. We assigned a magnitude to
these peaks equal to the number of reads mapping to within 500bp of the apex; only peaks
with a magnitude >10 were considered further. For each strain we aligned replicate apex
chromosomal locations using the dynamic programming algorithm as described (Knott et
al., 2009), with a gap penalty of 1000bp. Apices that did not align across all replicates were
removed from consideration. Next, for each strain we aligned peaks (387) with the set of
previously annotated origins listed in OriDB (Nieduszynski et al., 2007); peaks (35) that did
not align to an annotated origin were not considered further.

Origins that were not detected to incorporate BrdU within a given strain were assigned a
count equal to the number of reads that mapped to within 500bp of the average of its
corresponding detected apices. To test for differential BrdU-incorporation across strains,
we employed DESeq (Anders and Huber, 2010). Origin counts were normalized using
DESeq’s internal size and variance normalization strategies and were called as different
between two strains with a significance cutoff of FDR<0.005.

55

BrdU-IP-chip time-course data analysis
Due to the high proportion of enriched probes in BrdU-IP-chip experiments, within-array
normalization methods designed for ChIP-chip are not suitable (Knott et al., 2009). To
compensate for this, we developed a procedure and tested it on BrdU-IP-chip experiments
performed in the presence of HU. This method requires that un-enriched probes form a
dense cluster in the M=log(IP/Total) vs. A=(log(IP)+log(Total))/2 plane (Knott et al., 2009).
However, in BrdU incorporation experiments without HU (where the percentage of
enriched probes can reach 80%), this requirement is sometimes not met. To account for
this, we developed a technique specifically for such experiments. This method requires a
mock control, for which we hybridized BrdU-IP material obtained from a 12min BrdU pulse
using G1-arrested (non-replicating) cells against genomic DNA. First, we identified the best
axes on which to transform the experimental data by applying our previous method on the
control data (Knott et al., 2009). After transforming the control and experimental data onto
these axes, the median absolute deviations of both datasets were normalized to 1. Then,
the M values of the experimental data were location-normalized such that mean of the
lowest 20% of probes were equal to mean of the lowest 20% of control probes.
Subsequently, we followed our previous method (Knott et al., 2009).

Analysis of linear clustering of Fkh-regulated origins
To test whether Fkh-activated and –repressed origins cluster in separate groups linearly
along chromosomes, we defined a clustering metric equal to the number of “cuts” required
to separate Fkh-repressed and Fkh-activated origins (this is equivalent to the number of
instances where a Fkh-activated origin neighbors a Fkh-repressed origin, ignoring non-
56

Fkh-regulated origins). A low “cut” count indicates higher clustering of like-regulated
origins. We obtained a “cut” count of 65 in the experimental data. To test if this was
significantly low, we performed 10
6
simulations on the 352 origins that were detected in
WT or fkh1∆ fkh2∆C cells. In each simulation we randomly assigned 95 origins as Fkh-
activated, 80 as Fkh-repressed, and the remaining as Fkh-unregulated. Fewer than 1% of
the simulations resulted in a “cut” count <65 (Fig. S2D).

Analysis of Fkh-regulated transcription versus Fkh-regulated origin function
To determine whether proximal genes show co-regulation with Fkh-regulated origins, we
performed a permutation test on the distances between Fkh-regulated origins and the
nearest Fkh-regulated genes. Fkh-regulated genes were identified as those that showed
differential expression (DESeq FDR<0.01) between WT and fkh1∆ fkh2∆C cells in the same
condition (unsynchronized or G1-synchronized). This analysis was performed using both
the RNA-Seq and Rpb3-ChIP-Seq datasets, (genes detected as differentially expressed in
each of the experiments are listed in Table S1). For each experiment we calculated the
distance from each Fkh-regulated origin to the nearest Fkh-regulated gene’s promoter.
Next, 10
5
simulated origins sets were identified by randomly selecting 172 origins, and
randomly assigning 95 as Fkh-activated and 82 as Fkh-repressed. For each of these sets,
the minimum distances to the nearest Fkh-regulated genes were calculated. With this
analysis we determined for all possible pair-wise combinations (e.g. up-regulated gene and
Fkh-activated origin, down-regulated gene and Fkh-activated origin, etc.) that Fkh-
regulated origins are not significantly clustered with Fkh-regulated genes along the
chromosome.
57

To test for correlation of Fkh-regulated origins with flanking gene expression, we
performed regression analysis separately on Fkh-regulated origins lying within intergenic
regions flanked by diverging, converging, and tandemly oriented genes. For converging
and diverging intergenic regions, we used two covariates representing the unsynchronized
and G1-phase fkh1∆ fkh2∆C-WT RNA-Seq read count differences of the closest transcript
(as measured in bp between the origin’s ARS-consensus sequence (ACS) and the gene’s
nearest end) and two covariates representing the same difference measure in the farther of
the two transcripts. For tandem intergenic regions, two covariates represented
unsynchronized and G1-phase fkh1∆ fkh2∆C-WT RNA-Seq read count differences for the
converging gene and another two covariates represented the differences for the diverging
gene. In this analysis the only covariate that showed significant correlation with origin
regulation was the gene farthest away from origins within converging intergenic regions in
unsynchronized cells (p<0.05). A closer inspection revealed that this correlation was due
to four outlying data points, and when these were removed, the same analysis found no
covariate to be significantly correlated with origin regulation. Furthermore, the application
of this same analysis to read count differences in the Rpb3 ChIP-Seq data showed no
covariate to be significantly predictive of origin regulation.

Chromosome conformation capture on chip (4C)
Chromatin isolation: 50mL of G1-sychronized cells were crosslinked and harvested as
described for ChIP-chip (Viggiani et al., 2009). Cells were suspended in 9.5mL Buffer Z
58

(0.7M Sorbitol, 50mM Tris (pH 7.4), heat sterilized) plus freshly added 2-mercaptoethanol
(20mM final) and protease inhibitor cocktail (Roche, Mini Complete). 0.5mL Zymolyase
100T (ICN, 10 mg/mL freshly made in Buffer Z) was added and the suspension was
incubated at 30°C with gentle agitation, 35 min. The suspension was split into six 2mL
microcentrifuge tubes and centrifuged at 16,000g, 20 min at 4°C. The supernatants were
discarded, each pellet was suspended in 300µL NP buffer (1M Sorbitol, 100mM Tris (pH
7.4), 50mM NaCl, 5mM MgCl 2, 1mM CaCl 2, heat sterilized) containing 0.5mM Spermidine
(freshly added from 250mM stock) by gently pipetting with a wide-bore pipet tip, and the
samples were pooled in a 2mL microcentrifuge tube.

Digestion and ligation I: The suspension was centrifuged as above and the pellet was
suspended in 500µL ice-cold 1X NEB (New England Biolabs) digestion buffer II, and
centrifuged again. This wash step with digestion buffer was repeated and the pellet was
suspended in 50µL 1X NEB digestion buffer II. 42µL 1% SDS was added, mixed gently,
incubated at 60°C, 15 min. 328µL of ice-cold 1X NEB digestion buffer II was added and the
resulting suspension was centrifuged at 600g, 1 min at 4°C. 400µL of the supernatant was
transferred to a fresh microcentrifuge tube (the remainder was discarded), and 44µL 10%
Triton-X100 was added and mixed gently by pipetting with a wide-bore pipet tip. This
suspension was placed on ice for 15 min, after which 58.4µL of H 2O, 16µL 10X NEB
digestion buffer II, and 1.6µL BSA (NEB, 10mg/mL) were added.

59

4µL XbaI (NEB, 100 U/µL) was added, mixed gently, and incubated at 37°C for a minimum
of 8hr while shaking at 275 rpm. 10µL H 2O, 50µL 10% SDS, and 9µL 0.5 M EDTA was
added, mixed, and incubated at 65°C for 10 min, followed by 60°C for 10 min, and on ice for
5 min. The sample was transferred to a 15mL conical screw-cap tube on ice and 3554µL
H 2O, 250µL 10X T4 DNA ligase buffer (NEB), 50µL BSA (10mg/mL), 500µL 10% Triton-
X100, and 125µL 1M Tris (pH 7.5) were added, mixed gently, and incubated on ice, 15 min.
While on ice, 2µL T4 DNA ligase (NEB, 400U/µL) was added, mixed gently, and incubated at
16°C for 4 hr, after which 60µL 0.5 M EDTA was added.

To the ligated sample, 50µL 5M NaCl and 5µL RNAase A (20mg/mL) were added, mixed,
and incubated at 37°C, 1 hr. 25µL Proteinase K (20mg/mL) was added, mixed, and
incubated overnight at 65°C. The sample was transferred to a 15mL phase-lock tube (5-
Prime, 2302850) and the DNA was purified by extraction with 6mL
phenol:chloroform:isoamyl alcohol (25:24:1) and centrifugation according to the
manufacturer’s instructions. To the 4.2mL of aqueous solution recovered, 225µL 5M NaCl
and 6µL glycoblue were added and mixed, and 11mL of ice-cold ethanol was added, mixed,
and incubated at -20°C, 8 hr. The sample was aliquoted into eight 2mL microcentrifuge
tubes and centrifuged at ~16,000g, 30 min at 4°C. After discarding the supernatant, each
pellet was dissolved in 50µL 1X TE and the samples were pooled. 30µL 3M NaOAc (pH 5.2)
and 825µL of ice-cold ethanol were added, mixed, and incubated at -80°C, 2 hr. The
precipitate was recovered by centrifugation at 16,000g, 30 min at 4°C, and after discarding
the supernatant, the pellet was dissolved in 50µL TE.
60

Digestion and ligation II: To 25µL (~100ng) of the ligated sample, 64µL H 2O, 10µL 10X NEB
digestion buffer II, 1µL BSA (10mg/mL, NEB) were added and mixed, and 2µL of MseI (10
U/µL, NEB) was added, mixed, and incubated at 37°C, 2 hr. 1µL 20% SDS was added and
incubated at 65°C, 20 min; 30µL 10% Triton-X100 was added and incubated on ice for 15
min. 757µL H 2O, 100µL T4 DNA ligase buffer, and 10µL BSA (10mg/mL) were added and
incubated on ice for 15 min. While still on ice, 2µL T4 DNA Ligase (400U/µL) was added,
mixed by pipetting gently, and incubated overnight at 16°C. The sample was split into two
500µL aliquots (in 2mL microcentrifuge tubes) and 25µL 5M NaCl, 2µL glycoblue, and
1.2mL ice-cold ethanol was added to each, mixed and incubated at -20°C, 2 hr. The
precipitate was recovered by centrifugation at 16,000g, 30 min at 4°C; the supernatant was
discarded and each pellet was dissolved in 25µL TE and pooled.

Amplification and microarray analysis: 5µL was amplified by standard PCR (25 cycles) with
the following primers: 5'CTAAGTGTCCTGTTTCGGAAC, and 5'CAGGCCGCTCTTATAAAATGA.
1µg amplified DNA was labeled with Cy5 and hybridized against Cy3-labeled reference
DNA (G1-synchronized total genomic DNA) as described for BrdU-IP-chip (Viggiani et al.,
2010). Analysis was performed as described in (Knott et al., 2009) to identify enriched
probes, and Xba1 fragments containing >3 enriched probes immediately adjacent to either
cut site were deemed to be interacting.
All original and processed data files can be found at http://www.ncbi.nlm.nih.gov/geo/
under accession number GSE33704.
61

Chapter 3: Analysis of Fkh1 and Fkh2 Chromatin Binding

3.1 Background

Previous studies on transcription factor binding have used chromatin immunoprecipitation
analyzed by DNA microarrays to investigate chromatin binding of Fkh1 and Fkh2, and
comparison of enriched regions with locations of Forkhead binding motifs revealed a few
hundred genomic binding loci for each protein
73–75
. However, these studies sought to
broadly investigate the binding of an array of transcription factors, and the binding data of
individual transcription factors was lacking for a variety of reasons including the use of
early microarray technology with only intergenic regions represented, and with only one
probe per region. As such, existing binding data failed to detect Fkh1 and Fkh2 binding at
sites where these proteins would be expected to bind, including Fkh-activated origins with
Forkhead binding motifs present. We sought to generate a more complete map of Fkh1 and
Fkh2 chromatin binding, and undertook an in-depth, Fkh1- and Fkh2-specific study
utilizing an improved microarray platform.
Our results indicate more abundant binding of both Fkh1 and Fkh2 broadly throughout the
genome with both shared and unique binding sites, despite the similarity of Fkh1 and Fkh2
binding motifs. Chromatin architecture differs at sites that bind either or both Fkh1 and
Fkh2 where we find nucleosome free regions (NFRs) of differing widths. We also uncover
cell cycle-regulation of Fkh1 and Fkh2 binding at certain genomic elements such as Fkh-
activated replication origins, and uncover a variety of additional genomic and genetic
elements which bind Fkh1 and Fkh2, including RNA Pol III-transcribed genes and
62

centromeres. These findings provide an expanded map of Fkh1 and Fk2 chromatin binding,
provide novel insight into origin regulation, and suggest novel roles for Fkh1 and Fkh2 in
genome regulation.
The following chapter is taken from the 2014 PLoS One research article “Fkh1 and Fkh2
Bind Multiple Chromosomal Elements in the S. cerevisiae Genome with Distinct
Specificities and Cell Cycle Dynamics” by A. Zachary Ostrow
‡
, Tittu Nellimoottil
‡
, Simon R.
V. Knott, Catherine A. Fox, Simon Tavaré, and Oscar M. Aparicio (
‡
equal contribution).
Supplemental figures are listed below in Supplemental Figures, and other supplemental
materials, including data sets, are available from PLoS.

63

3.2 Results

An expanded map of Fkh1 and Fkh2 binding to the S. cerevisiae genome: To assess
the genome-wide distribution of Fkh1 and Fkh2, we performed ChIP-chip using several
immunologic approaches. First, we used a polyclonal antibody that immunoprecipitates
Fkh1 and Fkh2 (herein referred to as “anti-Fkh1/2 poly”) and carried out experiments in
wild type (WT) and fkh1∆ fkh2∆ (control) strains. To validate and supplement these
results, we also performed the analysis with anti-MYC monoclonal antibody in WT strains
expressing C-terminally epitope-tagged Fkh1 (Fkh1-Myc9), Fkh2 (Fkh2-Myc13), and an
untagged (control) strain. Experiments were performed in triplicate and analyzed with
tiling microarrays covering unique sequences of the S. cerevisiae genome (one ~60bp
oligonucleotide probe every ~80bp of unique sequence). Data from individual replicates
were analyzed to identify significantly enriched regions (p ≤ 0.05) having a minimum
length of 500bp (see Methods). Segments of these enriched regions that overlapped by at
least 500bp in at least two replicates were deemed “bound loci”, while any such regions
overlapping substantially (  50% of length) with regions deemed bound in the control
strains (fkh1∆ fkh2∆ for anti-Fkh1/2 poly and untagged for anti-Myc) were excluded from
the set. Plots of the data across chromosome VI show the average from the three replicates
of each experiment with bound loci colored (Fig. 3.1; plots of all chromosomes are
presented in Fig. S3.1).
64

Figure 3.1. Genome-wide analysis of Fkh1 and Fkh2 chromatin binding. Plots show
averaged ChIP-chip signal (M) from three experimental replicates along chromosome VI,
with enriched regions plotted in purple. The antibody and strain genotype used for each
experiment are indicated to the left of each panel; the corresponding strains from top to
bottom are: CVy43, ZOy1, CVy138, CVy139, ZOy3, ZOy4, and CVy43. Triangles on the
bottom panel indicate the position of determined binding sites as described in the text,
color-coded by classification.

Analysis with anti-Fkh1/2 poly identified 1503 Fkh1 and/or Fkh2 (Fkh1/2)-bound loci
that were not detected in the control fkh1∆ fkh2∆ cells (Table S1). To investigate the
dependence of these bound loci on Fkh1 and Fkh2, we performed ChIP-chip on fkh1∆ and
fkh2∆ strains with anti-Fkh1/2 poly (Fig. 3.1, Table S1). We analyzed the resulting binding
65

maps to identify overlapping regions (see Methods), which are indicated in the
corresponding intersection of the Venn diagram (Fig. 3.2A). Focusing on the intersection of
the WT with the fkh1∆ and fkh2∆ sets, 702 bound loci in WT and fkh2∆ cells were not bound
in fkh1∆ cells, defining these as Fkh1-dependent loci and suggesting these loci specifically
bind Fkh1 (Fig. 3.2A). 63 sites bound in WT and fkh1∆ cells were not bound in fkh2∆ cells,
defining these as Fkh2-dependent loci and suggesting that these sites specifically bind
Fkh2. The remaining 605 loci are defined as Fkh1/2-dependent loci, suggesting that these
sites can bind both Fkh1 and Fkh2, either simultaneously or in the absence of the other.
Analysis with anti-Myc identified 1013 Fkh1-Myc- and 700 Fkh2-Myc-bound loci, which
were not detected in the untagged strain (Table S3.1). These sets showed substantial
overlap with the Fkh1/2-poly set, with 81% of the Fkh1-Myc and 70% of the Fkh2-Myc
bound loci intersecting with the Fkh1/2 poly set, while the union of Fkh1-Myc and Fkh2-
Myc sets intersected with 61% of the larger Fkh1/2 poly set (Fig. 3.2B). The Fkh1-Myc and
Fkh2-Myc sets also showed substantial overlap with each other, with 452 loci exhibiting
binding to both proteins. An additional 530 loci bound Fkh1-Myc specifically, and 221 loci
bound Fkh2-Myc specifically.
To test these inferred specificities, we examined Fkh1-Myc and Fkh2-Myc binding at Fkh1-
and Fkh2-dependent loci determined in the experiments with anti-Fkh1/2 poly. Fkh1-
dependent loci showed greater overlap with Fkh1-Myc (58%) than Fkh2-Myc (19%) loci
(Fig. 3.2C), whereas a more balanced proportion of all Fkh1/2 poly loci overlapped with
Fkh1-Myc (55%) and Fkh2-Myc (34%) loci (Fig. 3.2B), consistent with specific or
preferential binding of Fkh1 to the set of Fkh1-dependent loci. In contrast, the
comparatively small number of Fkh2-dependent loci showed similar overlap with Fkh2-
66

Myc (22%) and Fkh1-Myc (21%) loci (Fig. 3.2D). Overall, the multiple approaches, use of
controls, and good overlap between datasets suggests we have generated robust Fkh1 and
Fkh2 binding data. We consolidated the data into three, non-overlapping sets for further
analysis, yielding: 828 Fkh1-only loci, which were only detected to bind Fkh1, 285 Fkh2-
only loci, which were only detected to bind Fkh2, and 541 Fkh1and2 loci, which were
detected to bind Fkh1 and Fkh2 (see Methods, Table S2).
To examine these sets of Fkh1 and Fkh2 binding loci further, we searched for Fkh1 and
Fkh2 consensus binding sequences within the called regions. Using previously reported
position-weight matrices of Fkh1 and Fkh2 consensus sequences
76
, we determined
coordinates for Fkh1 and Fkh2 consensus sequences in the yeast genome (Table S3). The
Fkh1 and Fkh2 consensus sequences are very similar to each other, so we searched for the
presence of either one, within each set of bound loci. 72%, 45%, and 81% of the Fkh1-only,
Fkh2-only, and Fkh1and2 bound loci, respectively, contained at least one Fkh1/2
consensus sequence match (Fig. 3.2E).
67

Figure 3.2. Correlation of Fkh1 and Fkh2 binding sites identified in different
experiments. A-D) Venn diagrams show overlap between binding regions identified
and/or categorized in different experiments. The area of the circle representing each
group and the degree of intersection between groups are proportional to the number of
binding loci in each group and degree of intersection, respectively. Discrepancies in
number of total binding loci corresponding to datasets between the different Venn
diagrams result from the method for calculating intersection between the sets (see
Methods S1). E) Pie charts show the percentage of binding loci in each group for which the
indicated number of matches to Fkh1 and/or Fkh2 consensus binding site(s) were
identified. Because the values were rounded to the nearest whole number, the sum of
percentages in two of the pie charts exceeds 100%.
68

Fkh1 and Fkh2 are associated with distinct chromatin architectures: Fkh1 and Fkh2
have been implicated in the regulation of chromatin structure through the recruitment of
chromatin modifiers and remodelers
77–79
, so we examined the chromatin structure
associated with Fkh1 and Fkh2 binding. To achieve base-pair resolution necessary to
compare binding with nucleosome positioning, we examined Fkh1- and Fkh2-bound loci
containing a single Fkh1/2 consensus sequence(s) and aligned these sequences with a
published map of nucleosome positions
80
. We plotted the nucleosome density in a 2kb
region surrounding each consensus sequence bound by Fkh1-only, Fkh2-only, and
Fkh1and2 loci, as separate sets (Fig. 3.3A). The data show differences in the nucleosome
densities associated with these bound loci, with Fkh1-only loci localizing to narrower
nucleosome-depleted regions than Fkh2-only and Fkh1and2 loci. We consolidated the data
into an average nucleosome density profile for each set and plotted the profiles together for
comparison (Fig. 3.3B). Estimation of the size of the nucleosome-depleted regions
indicates a length of ~400bp at Fkh1and2 loci versus ~275bp at Fkh1-only loci, a
difference of approximately one nucleosome.
69

Figure 3.3. Distinct nucleosome positioning at Fkh1-only loci versus loci that bind
Fkh2. A) The heat maps show density of MNase-protected sequences (Eaton et al 2010)
for 2kb regions centered on Fkh1/2 consensus sequences within enriched regions that
have only a single Fkh1/2 consensus sequence. B) Averaged signal intensities from (A) are
plotted. Arrows indicate the positions used to estimate length of nucleosome-depleted
regions reported in the Results.

Fkh1 and Fkh2 binding at regulated genes: Next, we examined the Fkh1 and Fkh2
binding data at genes previously reported to be under Fkh1/2 regulation. We generated
70

heat maps of Fkh1-only, Fkh2-only, and Fkh1and2 binding frequency for 10kb regions
centered and oriented on the start codons of 32 CLB2-cluster genes and, for comparison,
two additional groups of co-regulated genes: 13 “CLN2-cluster” genes expressed in late G1-
phase and 18 “SIC1-cluster” genes expressed in late M-early G1-phase (Fig. 3.4A)
81
. The
heat maps show enrichment of Fkh1 and Fkh2 over the promoter regions of CLB2-cluster
genes, with 38% of these regions binding both proteins, an additional 21% binding only
Fkh2, and an additional 8% binding only Fkh1. In comparison, Fkh1 and Fkh2 were not
enriched over the promoters of the CLN2-cluster genes, as expected. Interestingly, some
enrichment of Fkh1 and Fkh2 was apparent over SIC1-cluster genes, which is consistent
with Fkh1 and Fkh2 acting as anti-activators of a subset of SIC1-cluster genes resulting in
their activation by Ace2 but not by Swi5
79
. To examine this more closely, we divided the
SIC1 gene cluster into subsets activated by transcription factor Ace2 only, Swi5 only, or
either factor, and generated heat maps of Fkh1 and Fkh2 binding frequencies (Fig. 3.4B).
The results show occupancy of Fkh1 and Fkh2 at 38% of Ace2-only genes, but little to no
occupancy at other SIC1-cluster genes, confirming that Fkh1 and Fkh2 specifically bind
Ace2-only genes
79
. These findings demonstrate that our data recapitulate known features
of Fkh1 and Fkh2 binding.
71

Figure 3.4. Fkh1 and Fkh2 binding with target genes. Heat maps show 10kb regions of
summed binding data for the indicated types of binding loci (Fkh1-only, Fkh2-only,
Fkh1and2) surrounding the groups of features indicated above the heat map. The color
represents the frequency of enriched binding sequences called for each group of features,
amongst total number of features (n) included in each group. A) Fkh1 and Fkh2
enrichment frequencies surrounding CLB2-, CLN2-, and SIC1-cluster genes are plotted as
separate groups, with the respective ORFs aligned by their start codons at coordinate 0,
with transcription toward positive coordinates to the right. B) Fkh1 and Fkh2 enrichment
frequencies surrounding Ace2-only-regulated genes, Ace2- or Swi5-regulated genes, and
Swi5-only-regulated genes are plotted with the ORFs aligned and oriented as in (A).

Fkh1 and Fkh2 binding at replication origins: Fkh1/2 were recently identified as
regulators of the initiation timing of replication origins throughout the budding yeast
genome
72,82
. In fkh1∆ fkh2∆ cells, the initiation of many early origins is delayed, and these
origins are locally enriched for Fkh1/2 consensus binding sequences. For a few tested
origins, Fkh1/2 binding sequences in cis were shown to be essential for regulation of the
proximal origin. However, previous ChIP-chip analysis did not report Fkh1/2 binding at
72

many Fkh-regulated origins
73–75
, suggesting that Fkh1/2 might act over longer distances to
regulate some origins. To examine the Fkh1- and Fkh2-bound loci we have identified in
relation to replication origins, we divided origins (termed ARS in yeast) into three groups
defined by their change in origin activity in fkh1∆ fkh2∆ cells in our previous study: Fkh-
activated origins, which showed reduced early firing, Fkh-repressed origins, which showed
increased early firing, and Fkh-unregulated origins, which showed no significant change in
early firing
72
. For each set of origins, we generated heat maps representing the frequency
of Fkh1-only, Fkh2-only and Fkh1and2 bound loci for a 10kb region centered and oriented
on the ARS Consensus Sequence (ACS), which is the essential origin-defining sequence (Fig.
3.5A). Fkh-activated origins are enriched for proximal Fkh1 binding, with 42% of these
origins associated with Fkh1-only loci and an additional 27% associated with Fkh1and2
loci, while only 2% are associated with Fkh2-only loci. Fkh-unregulated origins are also
enriched for Fkh1, with 31% of these origins associated with Fkh1-only loci and 21%
associated with Fkh1and2 loci. Only 11% of Fkh1-only and no Fkh2-only loci are
associated with Fkh-repressed origins, however, 20% of Fkh-repressed origins are
associated with Fkh1and2 binding loci. These results are consistent with our previous
demonstration that Fkh1/2 consensus binding sequences are enriched near Fkh-activated
origins and required for their regulation, whereas Fkh1/2 consensus sequences are
depleted near Fkh-repressed origins
72
. However, these results also suggest that Fkh1/2
binding is not sufficient to establish Fkh-activation or that Fkh-unregulated origins are
associated with factors that oppose Fkh-origin regulation (see Discussion). The results
further suggest that Fkh-repression of origins may in some cases derive from direct
binding by Fkh1/2.
73

The predominance of Fkh1 over Fkh2 binding near origins was consistent with our
previous finding that fkh1∆ cells deregulate origin timing whereas fkh2∆ cells do not (see
Introduction). However, our previous study also showed that fkh1∆ fkh2∆ cells deregulate
many additional origins than fkh1∆ cells, suggesting a primary role for Fkh1 in origin
timing regulation and a secondary role for Fkh2
72
. Given our previous findings that both
Fkh1 and Fkh2 consensus binding sequences are enriched near Fkh-activated origins, the
preference for Fkh1 binding indicates the existence of additional determinants of Fkh1
versus Fkh2 binding specificity. Possible candidates for determining Fkh1 versus Fkh2
binding specificity are Mcm1 and Ndd1. In vitro, Mcm1 binds cooperatively with Fkh2 , but
not Fkh1, to DNA sequences containing closely juxtaposed Fkh1/2 and Mcm1 consensus
binding sequences
57,63
. In vivo, Fkh2 recruits Ndd1 to CLB2-cluster gene promoters
through interactions involving the unique C-terminus of Fkh2
65,66
.
To examine the relationship of Mcm1 and Ndd1 with Fkh1 and Fkh2 binding, we plotted
Fkh1-only, Fkh2-only, and Fkh1and2 binding loci for 10kb regions centered on 79 Mcm1
and 315 Ndd1 binding sites, which were previously reported to bind the respective protein
in ChIP experiments and contain a recognizable consensus sequence for the respective
protein
73,74
. The heat maps show strong enrichment of Fkh1and2-bound loci proximal to
Mcm1 binding sites, with 41% of Mcm1 binding sites overlapping with a Fkh1and2 locus. A
few Fkh1-only and almost no Fkh2-only loci were associated with Mcm1 binding sites (Fig.
3.5B). Ndd1 exhibited a similar pattern of association, with 52% of Ndd1 binding sites
proximal to Fkh1and2 loci, 13% of Fkh1-only loci and almost no Fkh2-only loci are
proximal to Ndd1 binding (Fig. 3.5B). Because Fkh-activated replication origins are
associated predominantly with Fkh1-only binding loci, this result implies that neither
74

Mcm1 nor Ndd1 associates with most Fkh-activated origins. We tested this directly by
searching for Mcm1 and Ndd1 binding sites proximal to replication origins, and for
comparison, to CLB2-cluster genes. The results show no instances of Mcm1 binding sites
within 500bp of any of the replication origin classes, whereas 19% of CLB2-cluster genes
are within 500bp of an Mcm1 binding site (Fig. 3.5C). Like Mcm1, Ndd1 binding sites are
also enriched at CLB2-cluster genes, with 22% of CLB2-cluster genes proximal to an Ndd1
site. In contrast to Mcm1, however, Ndd1 binding sites are associated with 10% of Fkh-
unregulated origins, representing significant enrichment with this origin class, and with
3% and 4% of Fkh-activated and Fkh-repressed origins, respectively (Fig. 3.5C). These
results suggest that recruitment of Ndd1 to replication origins might counteract Fkh1/2-
regulation of origin function (see Discussion).
75

Figure 3.5. Fkh1 and Fkh2 binding with replication origins. (A-B) Heat maps show
10kb regions of summed binding data for the indicated types of binding loci (e.g.: Fkh1-
only, Fkh2-only, Fkh1and2) surrounding the groups of features indicated above the heat
map. The color represents the frequency of enriched binding sequences called for each
group of features, amongst total number of features (n) included in each group. A) Fkh1
and Fkh2 enrichment frequencies surrounding Fkh-activated, Fkh-unregulated, and Fkh-
repressed origins are plotted with each group aligned and oriented at coordinate 0 by each
origin’s ARS consensus sequence (ACS). B) Fkh1 and Fkh2 enrichment frequencies are
plotted around Mcm1 and Ndd1 binding sites, which are aligned and oriented by Mcm1 and
Ndd1 consensus sequences, respectively. C) The graph shows the percentage of each
element class having an Mcm1 or Ndd1 binding site within 500bp. Asterisks indicate
values significantly greater than expected on a random basis at p<0.01 (see Methods S1).

76

Other genetic elements associated with Fkh1 and Fkh2 binding: To determine
whether Fkh1 and Fkh2 bind and potentially regulate other genomic elements, we plotted
Fkh1 and Fkh2 binding loci near different sets of genomic elements (as defined in
Saccharomyces Genome Database) (Fig. 3.6). Fkh1 and Fkh2 showed remarkable
occupancy near several of these elements, with occupancy rates comparable to those at
CLB2-cluster genes and Fkh-activated origins. As a group, ORFs show minor enrichment of
Fkh1 or Fkh2 relative to flanking sequences. ARSs, telomeres, and subtelomeric X and Y´
elements, are associated predominantly with Fkh1-only, with 15-20% of these elements
proximal to a Fkh1-only locus. In contrast, centromeres, 5  UTR introns, snoRNAs, and
tRNAs are more frequently associated with Fkh1and2 binding loci, which are proximal to
40-60% of these elements; these elements show more modest levels of enrichment for
Fkh2-only loci (see Table S4 for list of genes with Fkh1/2 enrichment upstream).
Fkh1and2 binding loci are also proximal to 20-30% of introns, ncRNAs, retrotransposons,
and dispersed long terminal repeats (LTRs). Interestingly, ncRNAs were associated with
Fkh1-only binding loci at a similar frequency as with Fkh1and2 loci. These findings suggest
that Fkh1 and Fkh2 have unrecognized roles in the regulation of Pol III-transcribed genes,
intron processing, and centromere function.
77

Figure 3.6. Analysis of Fkh1 and Fkh2 binding proximal to various genetic elements.
Fkh1 and Fkh2 enrichment frequencies surrounding different classes of genetic elements
are oriented and aligned at coordinate 0 according to the first base position of each
element. The maximum frequency reached within 100bp (500bp for Y') of coordinate 0 is
indicated above each heat map. The asterisk indicates significant enrichment (p<0.001)
near coordinate 0 (see Methods).

Cell cycle dynamics of Fkh1 and Fkh2 binding: To gain further insight into the
mechanisms that Fkh1/2 use to regulate genes and origins in the cell cycle context, we
78

performed ChIP-chip of Fkh1 and Fkh2 with anti-Fkh1/2 poly in cells synchronized in
G2/M with nocodazole, in late G1 with -factor, and in early S with hydroxyurea. Data from
these experiments corresponding to the Fkh1/2 binding loci identified above were
subjected to k-means clustering analysis according to the binding patterns of individual loci
across the three cell cycle stages (Fig. 3.7A, Table S2, see Methods). This analysis revealed
four distinct clusters that can account for most of the data, with each cluster representing a
distinct binding pattern across the cell cycle (Fig. 3.7A). The largest cluster of ~865
binding loci, named “High-S”, shows higher binding in early S-phase and lower binding in
G2/M and late G1. The High-G1 cluster shows higher binding in late G1 and lower binding
in G2/M and early S. The High-G2/M cluster shows higher binding in G2/M and lower
binding in late G1 and early S, while the Low G1 cluster shows lower binding in late G1 and
higher binding in early S and G2/M.
79

Figure 3.7. Cell cycle analysis of Fkh1 and Fkh2 binding. A) The k-means cluster-gram
on the left shows the average signal intensity of individual binding loci across the three
experiments, divided into four groups, which was found to account well for the data. The
graphs to the right of each cluster show the averaged signal of all sites in the cluster. B) For
each class of genetic element indicated, the number of proximal (+/- 500bp) Fkh1 and Fkh2
binding loci in each of the four clusters in (A) was counted to determine the distribution of
these binding sites amongst the four clusters. The colors in the graph correspond to the
colors of the four clusters in (A). C) The average signals of Fkh1 and Fkh2 binding loci
proximal (+/- 500bp) to the indicated genetic element class was determined and plotted
for the three cell cycle points.

80

To ascertain whether these cell cycle binding patterns are associated with specific
functional classes of Fkh1/2 binding loci such as those associated with CLB2-cluster genes
or replication origins, we determined the binding patterns of Fkh1/2 binding loci within
500bp of specific classes of genomic features analyzed above (Fig. 3.7B). This analysis
indicates that Fkh1/2 binding loci proximal to distinct genomic elements exhibit
significantly distinct cell cycle patterns of Fkh1/2 binding (see Methods). For example, the
High G1 binding pattern, which is the least frequent overall when all binding loci are
considered, is the most frequent pattern associated with ARS and X elements, and is also
significantly enriched at LTRs, ncRNAs, retrotransposons, tRNAs, and telomeres. The High
G1 pattern is also depleted at snoRNAs. The Low G1 pattern, which is infrequent in the
overall distribution, is significantly enriched at Introns, 5´ UTR Introns, snoRNAs, and
tRNAs; this pattern is also depleted at ARSs, X elements and telomeres. The High G2/M
pattern is modestly enriched at Introns, LTRs, and tRNAs, and is most notably depleted
near ARSs. The High-S pattern, which is most frequent overall, is correspondingly depleted
at most of the aforementioned elements that are enriched for another pattern. However,
the High-S pattern is not depleted at binding loci proximal to ORFs, telomeres,
centromeres, and 5´ UTR Introns.
81

Figure 3.8. G1-specific binding of Fkh1/2 at Fkh-activated origins. Plots show
averaged ChIP-chip signal from three experimental replicates along a 100kb region of the
left arm of chromosome III, with enriched regions plotted in purple. The cell cycle arrest
for each experiment is indicated to the left of each panel. Boxed loci are discussed in the
Results.

To scrutinize the binding dynamics more directly at these genomic elements, we plotted
Fkh1/2 binding profiles at loci specifically proximal to each set of elements (Fig. 3.7C). The
plots show distinct binding patterns associated with different element types. For example,
ARSs and telomeres show lower signals in G2/M and sharply higher signals in late G1 and
in early S. In contrast, centromeres and 5´ UTR Introns showed intermediate signals in
G2/M decreasing in early G1 followed by strikingly higher signals in early S. The remaining
82

elements also generally showed higher signals in early S compared with G2/M and early
G1, however, the overall degree of fluctuation was somewhat lower. With the exception of
the very low binding at telomeres in G2/M, binding levels show the greatest differences
amongst elements in late G1.
To examine Fkh1/2 binding at specific loci, particularly Fkh-activated origins, we plotted
the cell cycle ChIP data for a 100kb region of chromosome III (Fig. 3.8, see Fig. S3.2 for plots
of all chromosomes). This region includes early-efficient origins ARS305 and ARS306, the
silent mating-type locus HML, the Recombination Enhancer (RE) for mating-type donor
preference, and BUD3, a Fkh1/2-regulated CLB2-cluster gene, all of which are associated
with Fkh1/2 binding. A previous study reported binding of Fkh1 and Fkh2 to CLB2-cluster
target genes in late G1- and G2/M-synchronized cells, suggesting that Fkh1/2 bind
constitutively to CLB2-cluster target genes
67
. In agreement with these previous reports,
Fkh1/2 binding was strongly enriched at BUD3 at all cell cycle times tested. In contrast,
previous analysis of Fkh1 binding at the RE showed binding in G2/M but not in late G1
69
.
However, our data show binding of Fkh1/2 at all three cell cycle times, though we note a
decreased signal in late G1. At HML-I, Fkh1/2 binding was detected at all cell cycle times,
though the signal was decreased in G2/M. Unlike Fkh1/2 binding at all of these loci,
however, Fkh1/2 binding at Fkh-activated origins ARS305 and ARS306 showed strong
enrichment in G1-phase, but little or no enrichment in S- or G2/M-phases. These findings
reveal a new dimension of Fkh1/2 regulation and support the notion that Fkh1/2 function
through distinct mechanisms to regulate distinct classes of genetic elements.

83

3.3 Discussion

An expanded map of Fkh1 and Fkh2 binding to the S. cerevisiae genome: The recent
discovery that Fkh1 and Fkh2 regulate replication initiation timing
72,82
, along with exciting
new mechanistic insight into how Fkh1 regulates donor preference in mating-type
switching
70
, in addition to their well-established roles as transcription factors, have stoked
new interest into these versatile regulators of the genome. A primary goal of this study was
to gain a greater understanding of the relationship between Fkh1/2 binding and regulation
of replication origins. Elucidating a more complete and dynamic map of Fkh1 and Fkh2
binding loci throughout the genome enabled robust, genome-scale analyses of these
binding loci in relation to replication origins, as well as other functional genetic elements.
We identified hundreds of novel binding loci for both proteins, including shared and
specific loci. Analyses of these data showed binding to known binding loci and targets of
regulation such as CLB2-cluster genes, serving to validate these results. These new
genomic maps of Fkh1 and Fkh2 binding also provide a valuable resource for future
genome-wide and locus-specific studies.
Our analysis of Fkh1/2 binding throughout the genome paints a somewhat different
picture than previous studies
73,75
, with several-fold more binding loci, especially loci
binding only Fkh1, identified here. To provide further confidence for our sets of identified
binding loci, we searched for matches to Fkh1/2 consensus binding sequences. We found
that a large majority of Fkh1-only and Fkh1and2 loci contained at least one consensus
match within the enriched region, however, only slightly fewer than half of the Fkh2-only
84

loci contained a match. We chose not to use the presence of a consensus sequence as a
filter to reduce the number of called loci to avoid imposing this possible bias, as it remains
possible that close matches to the consensus sequence were missed, or that Fkh1/2 binds
some sequences independently of a consensus sequence. A related possibility is that
binding loci lacking a consensus sequence represent sites of “indirect” binding (as coined
by Bulyk and colleagues in
83
) where Fkh1/2 do not bind DNA directly but bind chromatin
through interaction with other DNA-binding proteins.
The much larger number of Fkh1-only versus Fkh2-only loci suggests that Fkh2 binding is
more specific or otherwise restricted. This might be explained by additional specificity
provided by its interacting partners Mcm1 and/or Ndd1. Hence, it is surprising that Mcm1
and Ndd1 binding sites are located proximal to Fkh1and2 loci but not to Fkh2-only loci.
This finding suggests that a different factor is responsible for the exclusive binding of Fkh2
at Fkh2-only loci. Whereas the more extensive nucleosome-depleted regions associated
with Fkh1and2 binding loci may be related to Mcm1 and/or Ndd1 binding, this does not
explain the similar nucleosomal structure observed at Fkh2-only loci, which are not
associated with Mcm1 or Ndd1. Instead, the narrower nucleosome-depleted regions
associated with Fkh1-only loci and the larger numbers of loci that bind Fkh1 (i.e., Fkh1-
only and Fkh1and2 loci) suggest that Fkh1 is better able to access potential binding
sequences in chromatin than Fkh2. A related possibility is that greater abundance of Fkh1
(1720 molecules/cell) versus Fkh2 (656 molecules/cell) results in a more restricted set of
binding loci for Fkh2
84
. Alternatively, Fkh1 and Fkh2 binding may regulate the remodeling
of chromatin in distinct ways resulting in the observed differences. This is currently under
investigation.
85

Fkh1 and Fkh2 binding at replication origins: In contrast to the high occupancy of both
Fkh1 and Fkh2 at CLB2-cluster genes, Fkh-activated replication origins are most frequently
bound by Fkh1 only, and whereas a minority of origins is also bound by Fkh2, almost none
binds only Fkh2. These findings are consistent with the differential effects on individual
origin function when either FKH1 or FKH2 is deleted
72
. These results also reinforce
previous findings that Fkh1/2 act directly in cis to regulate origin function
72,82
.
Nevertheless, we did not detect Fkh1/2 binding near one-third of Fkh-activated origins,
leaving open the possibility that the regulation of some origins occurs over a longer
distance or indirectly. We also detected Fkh1, and to a lesser degree Fkh1 and Fkh2,
binding at a fraction of origins in the Fkh-unregulated group. Some of these may represent
bona-fide Fkh-activated origins within this set that did not reach the significance threshold
to be classified as Fkh-activated in the previous study. However, another possibility is that
additional chromatin regulators binding in the vicinity of these origins oppose Fkh1/2
function, resulting in their Fkh-unregulated phenotype. Indeed, the presence of Ndd1
binding sites near Fkh-unregulated origins may explain why some of these origins are Fkh-
unregulated despite many of these origins being bound by Fkh1 and Fkh2.
The cell cycle-regulated association of Fkh1/2 with replication origins reported here is an
important advance toward a complete understanding of the mechanism of Fkh1/2-
regulation of origin timing. Previous studies have indicated that the establishment of the
replication-timing program occurs in the M to early G1 period
39,85
. More recent studies
indicate that the selective recruitment of replication initiation factors to licensed origins
during G1-phase determines early origin firing, and Fkh1/2 are required for this
recruitment (reviewed in Aparicio, 2013
86
). This strongly suggests that the G1-phase
86

recruitment of Fkh1/2 is essential for initiation factor recruitment and is linked to the
origin licensing cycle. This might involve interactions with protein(s) that license origins in
early G1-phase such as Mini-Chromosome Maintenance proteins, and/or might involve
regulation by CDK or DDK activities. Experiments are in progress to determine the
mechanism of cell cycle-regulated binding of Fkh1/2 to replication origins.
Novel genetic elements associated with Fkh1 and Fkh2 binding: A novel finding of this
study is the association of Fkh1/2 with a large number of functional genetic elements,
including centromeres, telomeres, transposable elements, introns and RNA Pol III-
transcribed genes, suggesting a possible role for Fkh1/2 in regulating the function of these
elements. Enrichment of Fkh1 upstream of tRNA genes has been previously reported
87
.
The high Fkh1/2 occupancy at tRNAs and snoRNAs, which are transcribed by RNA Pol III is
particularly intriguing given the known role of Fkh1/2 as a regulator of some Pol II-
transcribed genes. Furthermore, Fkh1/2 are thought to regulate origin timing and mating-
type donor preference by mediating long-range intra- and/or inter-chromosomal
interactions (reviewed in Haber, 2012 and Aparicio, 2013
68,86
), while highly expressed
tRNAs aggregate into clusters surrounding the nucleolus (reviewed in Hopper et al,
2010
88
). It will be interesting to determine whether Fkh1/2 regulate tRNA clustering or
expression. Similarly, the association of Fkh1/2 with transposable elements, centromeres
and telomeres, all suggest a function in chromosomal organization.
The Fkh1/2 association with one or more of these element classes may reflect co-
localization of two or more element classes where a single class is the functional target of
Fkh1/2. A possible case is the enrichment of Fkh1 with telomeres and subtelomeric X and
Y elements, which are associated with a high density of ARS elements
89,90
. Thus, the
87

binding of Fkh1 near subtelomeric origins likely explains their observed proximity to
subtelomeric elements and telomeres. Although telomeres and subtelomeres are late-
replicating, many of these regions become even later replicating in fkh1∆ fkh2∆ cells,
consistent with Fkh1/2 regulating subtelomeric origins
72
. tRNAs and retrotransposons
also co-localize with replication origins more frequently than expected at random
90
;
however, this relationship probably does not explain the Fkh1/2 association with these
elements because tRNAs and retrotransposons are primarily associated with Fkh1and2
binding loci whereas origins are primarily associated with Fkh1-only loci. Nevertheless,
yeast transposable elements frequently co-localize with tRNAs and Pol III-transcribed
genes so the association seen with these various elements may result from this co-
localization. Given the much higher occupancy of Fkh1/2 at tRNAs and snoRNAs and the
larger number of these elements compared with retrotransposons, we think it is more
likely that the association with retrotransposons reflects functional Fkh1/2 binding near
tRNAs and snoRNAs, rather than the converse. Whereas further studies will be required to
elucidate fully the role(s) of Fkh1 and Fkh2 at these various elements, these remarkably
robust associations strongly suggest that Fkh1 and Fkh2 have more global functions than
previously appreciated. It remains to be seen whether the association of Fkh1 and Fkh2
with a broad array of genetic elements can be explained by a common mechanism involving
higher-order chromatin organization.

88

3.3 Methods

Yeast strains and methods: All strains (see Table S5) are congenic with the W303
background, including FKH1 and FKH2 MYC-tagged strains, Z1448 and Z1370 respectively,
from the Young lab
73
. ZOy3 and ZOy4 were constructed by deletion of BAR1 in strains
Z1448 and Z1370, respectively, using BamHI-BglII-digested plasmid p∆bar1::URA3 with
lithium acetate transformation
91
. Cells were grown at 23°C for all experiments and
synchronized in late G1, early S, and G2/M by incubation for 3 h in 7.5 nM α-factor (Sigma,
T6901), 200mM hydroxyurea (Sigma, H8627), or 10 µg/mL nocodazole (Sigma, M1404),
respectively. ChIP-chip experiments were performed as described previously
92
, with the
following modifications and reagents: chromatin was sheared to an average size of 300bp
using a Covaris S2 instrument; immunoprecipitations were performed with 9E10 (Covance,
MMS150) at 1:100 followed by pull-down with Protein G Dynabeads (Invitrogen, 10004D),
or with anti-Fkh1/2 polyclonal antibody
93
, which was pre-crosslinked to protein A-
Sepharose 4B beads (Invitrogen, 10-1041), at 1:40 (packed bed volume). Up to 10ng
immunoprecipitated (IP) and total DNA samples were subjected to whole genome
amplification (Sigma, WGA2), followed by primer extension labeling with Cy5 and Cy3 end-
labeled random nonamers, as described previously
92
. Cy5-labeled IP and Cy3-labeled total
DNA samples were combined and hybridized to custom oligonucleotide tiling microarrays
(Roche-Nimblegen, 124k HX12) that tile one ~60bp oligonucleotide probe per ~80bp of
unique genomic sequence; the Maui hybridization system and reagents (Roche) were used
according to the manufacturer’s instructions, and image capture was performed using an
Axon 4100A scanner.
89

Microarray data analysis and peak calling: We used RINGO package
(http://www.biomedcentral.com/1471-2105/8/221) in BIOCONDUCTOR suite to perform
the microarray normalization. The ChIP peaks were calculated with a distCutOff value of
5000. The upperBoundNull method with a p-value of 0.05 was used to calculate the
threshold for calculating the enriched regions. M is the log 2 ratio of bound to total signal.
From each microarray experiment, we obtained a set of enriched regions defined by
chromosome number, start, stop, maxLevel, and score of each peak. For experimental
triplicates, all nucleotides were examined to identify those enriched in at least two of the
replicates. Nucleotides pertaining to contiguous stretches of enriched nucleotides ≥ 500bp
were identified. Finally, these enriched regions were eliminated if 50% or more of their
nucleotides overlaps with enriched nucleotides in the control datasets. The remaining
enriched regions are deemed “bound”.
Analysis of intersection between datasets: Bound regions from different datasets that
overlap by ≥ 100bp were deemed to intersect and were enumerated within the intersecting
region of the Venn diagrams. Details on set functions and construction of the Venn
diagrams are described in Methods S1
Calling Fkh1-only, Fkh2-only, and Fkh1and2 sets: Fkh1-only loci were defined as the
union of Fkh1-dependent and Fkh1-Myc loci followed by subtraction of Fkh2-Myc loci.
Fkh2-only loci were defined as the union of Fkh2-dependent and Fkh2-Myc loci followed by
subtraction of Fkh1-Myc loci. Fkh1and2 loci were defined as all loci with subtraction of
Fkh1-only and Fkh2-only loci. For union of sets, all nucleotides in the sets being combined
were included in the union. For subtraction of a set B from a set A, enriched regions in set
A were entirely eliminated from set A if they overlapped by ≥ 100bp with enriched
90

region(s) from set B. For smaller overlaps, only the overlapping nucleotides were
eliminated from set A. The remaining enriched sequences of set A comprise the subtracted
set.
Analysis of Fkh1/2 enrichment at genetic elements: Heat maps of Fkh1/2 binding
proximal to features of interest were constructed from two-dimensional binary matrices.
Each row of the matrix represents nucleotides on either side of one instance of the
chromosomal element of interest; there are as many rows as there are instances of the
element class under analysis. The central column (plotted as coordinate 0) represents a
central reference nucleotide for each instance of that chromosomal element, and on either
side are the surrounding nucleotides, with one nucleotide per column. A matrix value of 1
indicates that the nucleotide position was called as enriched in the ChIP analysis, whereas a
value of 0 indicates that the nucleotide was not enriched. The average value for each
column was plotted as the binding frequency. Values given in the text and figure are the
maximum binding frequency within 100bp (500bp for Y') of coordinate 0. Coordinates for
all genetic elements were acquired from SGD, with the exception that coordinates for ACSs
in Figure 5A were taken from
80
.
To test the significance of enrichment of Fkh1/2 binding in the vicinity of genetic elements,
we performed simulations to model the null distribution and then tested whether the
actual distribution was significantly higher than the null distribution. This method was not
applicable to X, Y', telomeres, or retrotransposons because of the lack of unique sequences
downstream of these elements. Details of the simulation and statistical tests are described
in Methods S1.
91

Cell cycle analysis of binding: Each enriched region identified by RINGO is associated
with a total score, which is a measure of enrichment across the entire region. We
normalized the total score to a score per nucleotide by dividing the total score by the length
of the enriched region. Next, we calculated a union set of all the enriched regions across
the three cell cycle experiments (G2/M; late G1; early S), which included all nucleotides
within enriched regions in any of the sets. The score associated with each enriched region
in the union set was calculated as the total of the per nucleotide score of each nucleotide
that belongs to that enriched region. Hence we ended up with three tracks of enriched
regions with the same chromosomal coordinates, but different total scores. These three
sets of total scores were subjected to k-means clustering with k=4, and distance measure
being Pearson's correlation coefficient. Fkh1/2 binding loci that were not enriched in any
of the three cell cycle experiments were excluded from this analysis.
We also assigned subsets of these union sets to genetic elements from SGD annotation file,
if the enriched region overlapped with or was < 100bp from a boundary of the feature.
Then we determined the class to which each feature-associated enriched region belonged
and constructed the stacked bar graphs of their distribution for each genetic element. A
chi-squared test was applied to the corresponding ratios of each set of Fkh1/2 binding loci
associated with a particular genetic element to test whether it was significantly different
from the null distribution after Bonferroni correction. The null distribution was chosen as
the membership ratios of all Fkh1/2 binding loci in the four cell cycle clusters. The
distributions at all individual classes of genetic elements were found to be significantly
different from the null.
Data Accession: Microarray pair files are available at GEO, accession number: GSE42567.
92

93

Chapter 4: Forkhead Transcription Factors Regulate Replication through
Dimerization and Cell Cycle Specific Binding at Origins

4.1 Background

The Forkhead Box (FOX) family of transcription factors is comprised of members sharing a
conserved Forkhead domain, a winged-helix DNA binding domain, and is conserved among
eukaryotes from yeast to humans
50,55
. Members of a mammalian FOX protein subfamily,
FOXP, have been shown to form domain swapped dimers in which a helix in the Forkhead
domain of each member of a pair extend, interlocking the proteins in a highly stable dimer.
The region of the Forkhead domain in which this extension occurs is highly conserved
among FOX proteins with an important substitution in the FOXP subfamily where a
conserved proline is replaced with an alanine. The flexible alanine at this position allows
helical extension and domain swapping, which is otherwise prohibited by the rigid
backbone of proline. The conservation of this alanine among the FOXP family members
suggests that a shared feature among this subfamily is the ability to form domain swapped
dimers with functional importance
51,52
.
The following chapter describes a project performed in collaboration with Yan Gan, Reza
Kalhor, Lin Chen and Oscar Aparicio.

94

4.2 Results

Yeast Forkhead proteins share key homologies with mammalian FOXP proteins: S.
cerevisiae Fkh1 and Fkh2 are homologous to mammalian FOX family proteins, performing
analogous functions of multiple FOX proteins in higher eukaryotes such as transcriptional
regulation, cell differentiation and cell cycle control
50
. Amino acid sequence analysis
between yeast Fkh1 and Fkh2 and mammalian FOX proteins reveals a particular homology
in the Forkhead DNA Binding Domain (Fkh-DBD) largely specific to the FOXP subfamily,
FOXP1 – FOXP4: an alanine in substituting for a highly conserved proline in conventional
FOX proteins (Fig. 4.1)
51
. In previous crystallographic and Nuclear Magnetic Resonance
(NMR) studies, this alanine has been demonstrated to be a structural necessity for the
domain swapped dimerization of FOXP1, FOXP2 and FOXP3, and functional dependency
has been demonstrated in FOXP3
51,52,94,95
. This alanine resides at the base of a helix in the
Forkhead domain and imparts flexibility to the helix, allowing extension of helices within
the Fkh-DBD between paired Forkhead proteins, creating the domain swapped dimer. In
other FOX family proteins, the rigid proline at this position prevents such flexibility,
disallowing domain swapped dimerization. In addition to this alanine, sequence analysis
identified in Fkh1 and Fkh2 two other residues, glutamine and asparagine, at key positions
found to be important in stabilization of domain swapped FOXP dimers
51
. The high
sequence homology between FOX family proteins and Fkh1 and Fkh2 allowed us to model
the structures of S. cerevisiae Forkhead proteins using the previously determined
crystallographic structures of domain swapped FOXP family proteins (Fig. 4.2). The
resulting model revealed that in Fkh1 and Fkh2, the alanine and two stabilizing residues
95

are positioned to function as in FOXP family proteins, supporting the idea of a role of
dimerization in Fkh1 and Fkh2 function in yeast.

Figure 4.1. Forkhead Sequence Analysis. Sequence alignment of mammalian (Homo
sapiens) FOXA, FOXO, and FOXP proteins and yeast (S. cerevisiae) Forkhead proteins are
shown. Residues involved in domain swapping in FOXP family proteins are highlighted in a
range from green to magenta according to helical propensity
96–98
.

Our previous finding that Fkh1 and Fkh2 interact with ORC and establish clustering of
replication origins was incorporated into a model in which a Forkhead protein binds an
origin and interacts with ORC at a second origin, thereby facilitating origin clustering
72
. The
finding that Fkh1 and Fkh2 share homology with FOXP family proteins in key residues
involved in domain swapped dimerization of Forkhead factors expands greatly on the
mechanism by which Fkh1 and Fkh2 bridge distal loci, both at replication origins at which
these factors have been demonstrated to act, and at other potentially architecturally-
regulated loci that bind Fkh1 and/or Fkh2
99
. Importantly, this homology does not exist
between FOXP factors and the remaining two S. cerevisiae Forkhead transcription factors
Hcm1 and Fhl1, which have not been implicated in higher order genomic regulation.
96

Figure 4.2. Fkh1/2 Structure Model. A model structure of domain swapped yeast
Forkhead DNA binding domain is shown bound to two separate dsDNA strands. The
structure is modeled and optimized from the crystal structure of a DNA-bound FOXP3
domain swapped dimer. Image created by Reza Kalhor and Lin Chen.

Yeast Forkhead dimerization: To investigate whether S. cerevisiae Forkhead proteins are
able to dimerize and whether Forkhead dimerization plays a role in replication origin
initiation, we constructed alleles in which specific residues involved in formation of domain
swapped dimers were mutated in such a way as to reduce or abolish dimerization. Using
site-directed mutagenesis, we mutated the FOXP- and S. cerevisiae Fkh1- and Fkh2-specific
alanine to a helix-breaking proline as is conventionally found in FOX proteins. Additionally,
we mutated the glutamine and asparagine found at the predicted core of the domain
swapped dimer which, based on our model, stabilize the complex, to their acid
counterparts glutamate and aspartate in order to repel paired Fkh-DBDs to destabilize and
disallow dimerization. We term these proteins Forkhead Domain Swap Mutants (Fkh1-dsm
and Fkh2-dsm), and have integrated these alleles in strains in which FKH1 and FKH2 have
been deleted (Table 4.1).
97

fkh1-dsm fkh2-dsm
A338P A375P
N335D N373D
Q314E Q351E

Table 4.1. Mutations introduced to Fkh1 and Fkh2 to create fkh1-dsm and fkh2-dsm:
Site directed mutagenesis was used to alter highlighted residues of Fkh1 and Fkh2 to create
the domain swap mutant (dsm) versions of the proteins.

We focus on Fkh1 for many analyses as we have found that this protein plays the
predominant role in regulation of replication initiation compared with its counterpart,
Fkh2. Additionally, Fkh1 binds replication origins that do not bind Fkh2 whereas Fkh2 only
binds origins in which we have detected Fkh1 binding, in accordance with the greater role
in replication timing played by Fkh1
72,99
. To determine whether Fkh1 is able to dimerize,
we performed a pull-down assay in which purified His-tagged Fkh1-DBD (6xHis-Fkh1-wt-
DBD) was incubated with yeast cell extract containing Myc-tagged full-length Fkh1 (Fkh1-
Myc). Samples were incubated with antibody against Myc (α-Myc), and
immunoprecipitated material was resolved by SDS-PAGE. Subsequent blotting with a
Forkhead-specific antibody (α-Fkh) uncovered the presence of both Fkh1-Myc and 6xHis-
Fkh1-wt-DBD, revealing that wild type Fkh1 is able to homodimerize, as expected from
crystal structure-based modelling. To determine whether the Fkh1-dsm mutations
diminish this interaction, the same experiment was performed with whole cell extract
containing Myc-tagged Fkh1-dsm (Fkh1-dsm-Myc). Western blotting with α-Fkh revealed a
greatly diminished interaction between wild type Fkh1 and Fkh1-dsm. Finally, both of
these experiments were performed using His-tagged Fkh-DBD of Fkh1-dsm (6xHis-Fkh1-
98

dsm-DBD). In the case of two Fkh1-dsm (Fkh1-dsm-Myc and 6xHis-Fkh1-dsm-DBD),
interaction between proteins was further diminished as compared with the Fkh1 (wild
type) and Fkh1-dsm hybrid interaction, revealing that dimerization of Fkh1-dsm is nearly
abolished and supporting a role in domain swapping for the three critical homologous or
analogous residues conserved between Fkh1, Fkh2, and mammalian FOXP-family proteins
(Fig. 4.3).

Figure 4.3. Fkh1 Pull-down Assay. Pull-down assay of whole cell extract (WCE)
containing Myc-tagged Fkh1 or fkh1-dsm which were isolated with antibody against the
Myc epitope. Purified Fkh1-Myc or fkh1-dsm-Myc were incubated with His-tagged Fkh1
DNA binding domain and analyzed by Western blot using antibody against yeast Forkhead
proteins.

Forkhead domain swap mutants have knock-out replication profiles: In order to test
the biological significance of Forkhead protein dimerization and ascertain whether or not
this phenomenon is involved in regulation of DNA replication initiation, we assayed
replication activity genome-wide (Fig. 4.4).
99

Figure 4.4. Fkh-dsm BrdU Plots. BrdU incorporation plots are shown; origins identified
as Fkh-activated (red), Fkh-unaffected (grey), and Fkh-repressed (green) are highlighted
beneath each plot.

To do this, cells were arrested in late G1 phase and synchronously released into S phase
with bromodeoxyuradine (BrdU) and hydroxyurea (HU). BrdU allows labeling of active
replication forks through its incorporation in place of thymidine in nascent DNA, while HU
activates the inter-S checkpoint through nucleotide depletion, allowing analysis of early S
phase by suppression of late origin initiation and continued replication of early origins.
Immunoprecipitation of BrdU-labeled DNA and subsequent high-throughput DNA
sequencing analysis (BrdU-IP-Seq) yields a genome-wide snapshot of replication activity,
with early-efficient origins yielding peak sizes proportional to their activity, and late or
dormant origins yielding small or no peaks
32,72
. To ensure that the replication phenotype
observed does not reflect potential transcriptional alterations, Fkh1-dsm and Fkh2-dsm
100

were separately integrated in the previously characterized fkh1Δfkh2Δ + pFKH2ΔC
background, which does not exhibit pseudohyphal growth caused by deregulation of CLB2
cluster genes, and exhibits transcription-independent deregulation of the replication
timing program
72
. Control strains were created by integrating wild type FKH1 or FKH2 in
the same parent strain as fkh1-dsm and fkh2-dsm. We found high similarity in the
replication profiles between Fkh1-dsm, Fkh2-dsm and fkh1Δfkh2Δ + pFKH2ΔC, with both
domain swap mutant strains exhibiting the global replication timing defect seen in
fkh1Δfkh2Δ + pFKH2ΔC, implicating dimerization as a functional necessity of Fkh1 and
Fkh2 in replication timing control. Both single mutant control strains exhibited
characteristic phenotypes of near-wild type replication profile of FKH2 deletion, and an
intermediate replication profile of FKH1 deletion, showing that the method of integrating
Fkh-dsm alleles did not lead to an effect on replication timing. Additionally, fkh1-dsm and
fkh2-dsm correlate well with fkh1Δ, potentially suggesting a somewhat intermediate
replication phenotype, but more likely reflecting unrelated differences between the dsm
strains and both the double mutant and the Fkh1 single mutant (Fig. 4.5).
101

Figure 4.5. Correlation coefficients between BrdU-IP-Seq of Forkhead mutants.
Correlation coefficients were obtained for strains used in BrdU-IP-Seq by removing inter-
origin regions and analyzing 5 kb regions centered around Fkh-activated and Fkh-
repressed origins. All correlations shown are significant.

Observing replication activity globally, the defect caused by deleting FKH1 and FKH2 or
mutation to their domain swap mutant counterparts has a similar effect on firing (Fig. 4.6).
Comparison of mutants to their respective controls at different classes of Fkh-regulated
origins shows that this trend is conserved among pairs (Fig. 4.7), and the trends in the
difference between mutant and control follows the same trend, lessening in intensity as the
control strain’s phenotype deviates further from wild type as seen in replication phenotype
of fkh1Δ.

102

Figure 4.6. Fkh-dsm and Fkh knockout strains have similar genome-wide replication
profiles. BrdU incorporation signal was summed over 5 kb centered at origins identified as
Fkh-activated, Fkh-unaffected, and Fkh-repressed. Heat maps are grouped both by strain
and its respective control, and origin classification.
103

Figure 4.7. Fkh-dsm BrdU Heat Map Ratio. BrdU incorporation signal was summed over
5 kb centered at origins. Ratio of signal in control strains over that of respective mutant
Forkhead strains are shown at each Forkhead-regulated origin class.

Forkhead dimerization is not required for CLB2-cluster gene regulation: Previous
work from our lab demonstrated that the replication timing defect of FKH1 and FKH2
double deletion is independent of the transcriptional defect caused by Forkhead deletion
72
.
To determine whether Forkhead dimerization is required for the proteins to perform their
known regulatory transcriptional functions, namely in regulation of CLB2-cluster genes to
promote cell cycle progression, and to ensure that Fkh1-dsm and Fkh2-dsm are functional
in that regard, we observed the morphological characteristics of Fkh1-dsm and Fkh2-dsm
strains to determine whether they exhibit pseudohyphal growth resulting from
deregulation of the CLB2-cluster in fkh1Δfkh2Δ cells
61
. Two major morphological defects of
pseudohyphal growth caused by FKH1 and FKH2 deletion in liquid media are cellular
104

elongation and formation of chains of cells as cell division proceeds. We used light field
microscopy to observe wild type, fkh1Δfkh2Δ, Fkh1-dsm, and Fkh2-dsm cells. Critically, we
used strains that did not contain pFKH2ΔC, as we have previously shown that C-terminally
truncated Fkh2 is sufficient to rescue the morphological defects, but not the replication
defect, caused by deletion of FKH1 and FKH2
72
. We show that cells containing Fkh1-dsm
and Fkh2-dsm display wild type morphology, indicating that both Fkh1-dsm and Fkh2-dsm
are competent in their roles as regulators of the CLB2-cluster (Fig. 4.8).

Figure 4.8. Forkhead mutant morphology. Light field microscopy images of wild type
and mutant Forkhead strain morphology are shown. Fkh-dsm strains are single Fkh
mutants as described previously. All strains shown lack the plasmid pFKH2ΔC.

105

Additionally, as previously demonstrated in agar scarring assays, cells exhibiting
pseudohyphal morphology caused by deletion of FKH1 and FKH2 invade agar when grown
on solid media
61
. We confirm the finding that fkh1Δfkh2Δ leads to agar scarring, and further
show that Fkh1-dsm and Fkh2-dsm, with or without pFKH2ΔC, suppress this morphological
defect (Fig 4.9). These morphological analyses of Fkh1-dsm and Fkh2-dsm show that
although both proteins fail to rescue the replication initiation defect caused by fkh1Δfkh2Δ,
they are able to properly regulate transcriptional activity relating to their main role as
transcription factors, regulation of the CLB2 gene cluster. Additionally, along with our
studies showing near abolishment of dimerization of Fkh1-dsm, this indicates that
dimerization is not a requirement for regulation of CLB2-cluster genes, and therefore fkh1-
dsm and fkh2-dsm represent separation of function alleles of FKH1 and FKH2 (see
Discussion).

Figure 4.9. Agar scarring assay of wild type and Forkhead mutants. Cells were gently
streaked onto agar plates and grown for ~5 days, after which time cells were gently
washed off with a stream of water. Cells exhibiting pseudohyphal growth invade agar
surrounding streaks (highlighted in red), while cells exhibiting normal growth phenotype
are removed after washing.
106

Chromatin binding of Forkhead domain swap mutants: After demonstrating
functionality of Forkhead domain swap mutants in transcriptional regulation of CLB2-
cluster genes, we sought to determine Fkh1-dsm binding specificities genome-wide
compared with wild type Fkh1. To do so, we employed chromatin immunoprecipitation
analyzed by microarray chip (ChIP-chip) of Myc-tagged Fkh1-dsm and Fkh1 (Fkh1-dsm-
Myc and Fkh1-Myc, respectively), both in the absence of FKH2. We previously showed that
Fkh1 binding to chromatin is cell cycle-regulated. Certain features, including early origins,
exhibit greatest levels of Fkh1 binding during late G1, before replication initiates but after
the replication timing program is established
39,40,99
. We found high similarity between
Fkh1-Myc and Fkh1-dsm-Myc chromatin association in late G1 with a number of important
exceptions. Specifically, observation of the Fkh-activated early origin ARS1307 and adjacent
Fkh-regulated CLB2-cluster member TEM1 on chromosome XIII reveal differential binding
of Fkh1-Myc and Fkh1-dsm-Myc in late-G1 arrested cells. ChIP-chip reveals enrichment of
Fkh1-Myc at both loci, and robust origin activity at ARS1307 measured by signal from
BrdU-IP-Seq. Interestingly, in the same assays, Fkh1-dsm-Myc signal is enriched TEM1, but
not ARS1307, and replication activity of this origin is nearly abolished (Fig. 4.10).
107

Figure 4.10. Forkhead binding and BrdU incorporation at TEM1 and ARS1307. Fkh1
binding (top) and BrdU incorporation (bottom) are shown for Fkh1-wt control (left) and
Fkh1-dsm (right). Fkh-activated origin ARS1307 and adjacent Fkh-regulated CLB2-cluster
gene TEM1 are indicated. The strain used for BrdU-IP-Seq contains pfkh2ΔC.

Observation of Fkh1-Myc and Fkh1-dsm-Myc chromatin binding genome-wide reveals the
same trend among other early origins (Fig. 4.11A) and most CLB2-cluster genes (Fig.
4.11B), with insubstantial enrichment of either Fkh1-Myc or Fkh1-dsm-Myc at Fkh-
repressed origins, consistent with previous findings with the wild type protein (Fig. 4.11A
and Fig. 4.11B, left panels)
99
. These results imply a mechanism in which Fkh1 dimerization
is involved in binding of Fkh1 in vivo at certain regulatory elements such as binding motifs
at early origins, but not transcriptional regulatory targets of Forkhead proteins, suggesting
that differences in Forkhead binding motifs or presence of binding partners such as ORC
may cause preferential binding of Fkh1 in either its dimerized or monomeric state.

108

Figure 4.11. ChIP-chip Origin Classes and CLB2 Cluster Heat Map. (A) Global ChIP-chip
signal for Fkh1-Myc (left) and Fkh1-dsm-Myc (right) is averaged 5 kb around Fkh-activated
origins, Fkh-unaffected origins, Fkh-repressed origins. (B) Median ChIP-chip enrichment of
Fkh1-Myc and Fkh1-dsm-Myc in a 5 kb window centered at ORF start are shown for CLB2-
cluster genes. Binding of Fkh1 or fkh1-dsm are statistically different at p < 0.05 for Fkh-
activated and Fkh-unaffected origins.

From our chIP-chip study of Fkh1-Myc and Fkh1-dsm-Myc binding in late G1 phase, we
chose four Fkh-regulated genomic targets that showed enrichment in either or both strains,
and on each performed an electrophoretic mobility shift assay (EMSA) to confirm and
109

further investigate our chIP results (Fig. 4.12). These targets included two early, Fkh-
activated origins, ARS305 and ARS607 that we identified as enriched for Fkh1-Myc, but not
Fkh1-dsm-Myc, chIP signal, and CLB2-cluster gene promoters TEM1 and BUD4, which we
identified as enriched for both Fkh1-Myc and Fkh1-dsm-Myc chIP signal. Additionally, Fkh1
binding at both ARS305 and ARS607 is cell cycle-regulated, peaking in late G1, whereas
Fkh1 occupation levels at both TEM1 and BUD4 persist throughout the cell cycle
99
. EMSA
analysis confirms that Fkh1 binds at each of the four loci, and replicates the differential
binding of Fkh1-dsm; ARS305 and ARS607 bind Fkh1 Fkh-DBD, and Fkh1-dsm Fkh-DBD
binding is nearly abolished. In contrast, TEM1 and BUD4 show binding of both Fkh1 and
Fkh1-dsm Fkh-DBDs, though binding affinity of Fkh1-dsm is reduced ~4-fold compared
with Fkh1. As these EMSAs do not likely reflect binding of Fkh1 dimers, these results
suggest that the modifications made to create the domain swap mutant strains alter
binding specificities in a manner consistent with predicted effects of removing the ability of
Fkh1 to dimerize on replication timing. Further testing is required to investigate in vivo
dimerization and the role it plays in establishing replication timing (see Discussion).
Additionally, post-translational modification of Fkh1 may play a role in vivo in cell cycle-
specific binding at features; for example, in nocodazole-arrested cells, there is a depletion
of Fkh1 binding at early origins
99
. Multiple sites near Fkh1 N335 and Q314 are potential
phosphorylation sites, for example (data not shown) which could serve to regulate
dimerization through electrostatic repulsion, as is mimicked in fkh1-dsm, or Fkh1 binding
to DNA through allosteric alterations to the DNA binding interface (see Discussion)
100
. A
similar mechanism of cell cycle-specific action has been demonstrated in Fkh2, which is
temporally phosphorylated to regulate expression of target genes
62,65
.
110

Figure 4.12. EMSA of Fkh1-DBD and fkh1-dsm-DBD at Fkh-regulated loci.
Electrophoretic mobility shift assay (EMSA) for Forkhead regulated origins ARS305 and
ARS607 are shown along with promoters of Forkhead-regulated CLB2-cluster genes TEM1
and BUD4. Assays were performed with Fkh-DBDs of both wild type Fkh1 and Fkh1-dsm.

Forkhead binding motifs differ at various Forkhead-regulated loci: Previous EMSA
analysis of Fkh1 and Fkh2 DNA binding at the CLB2-cluster gene SWI5 promoter revealed
the presence of multiple variations of the Forkhead DNA binding motif that bind both Fkh1
and Fkh2 with different affinities. The variations of the Forkhead motif RYMAAYA
identified were a “strong” binding site beginning with GG at the R position, and a “weak”
binding site beginning with TA at the R position
57
. Analysis of both BUD4 and TEM1
promoter regions reveals the presence of a strong Forkhead binding motif proximal to the
111

open reading frame. By contrast, ARS305 lacks a strong binding site altogether, and ARS607
contains a weak binding site within the ACS, and a more distal strong binding site.
Together, these analyses led us to consider that the specific binding motif or motifs present
at a Fkh-regulated locus may be involved in determination of cell cycle-dependency of that
locus. To address the differential binding of Fkh1 and Fkh1-dsm at certain loci, we
generalized the strong and weak binding sites to GGYMAAYA and TAYMAAYA, respectively,
and measured their presence at Fkh-activated, Fkh-unaffected, and Fkh-repressed origins.
We found that a similar proportion of each class contain one of either motif; however, Fkh-
activated origins are enriched for multiple weak binding motifs, and entirely depleted of
multiple strong binding sites (Fig. 4.13). Additionally, we find that DNA shape of Fkh
binding motifs closest to an origin’s ACS differs at Fkh-activated and Fkh-repressed origins
(data not shown). These distinctions provides insight into a possible mechanism by which
dimerization of Forkhead proteins can allow cell cycle-dependent binding of Fkh1 to
specific features such as early origins.

112

Figure 4.13. Strong and weak Forkhead motifs at origins by Fkh-regulation. Presence
of multiple strong or weak Forkhead motifs, GGYMAAYA and TAYMAAYA, respectively, are
measured at origins and evaluated by Forkhead-regulation class.

113

4.3 Discussion

Fkh1/2 and epigenetic regulation: It has been suggested that replication timing not only
correlates to the epigenetic landscape, but is an epigenetic marker itself
101
. Global
examination of replication timing in the mouse genome during embryonic stem cell
differentiation revealed that significant alterations occur in the replication timing program
as cells differentiate, and that different cell types have specific replication timing programs
associated with changes in genome architecture
102
. S. cerevisiae Forkhead proteins are its
involved in the regulation of genome architecture, imparting early firing on certain origins
and establishing the replication timing program
72
. Replication timing, as an epigenetic
feature, is related to long term chromosome structure which persists over multiple cell
cycles. Thus, in principle, Forkhead proteins serve to regulate genome architecture on
multiple levels: first to alter the architectural landscape in a cell, while in doing so affecting
heritable, long term alterations in chromosome structure. Evaluating the long term effects
of FKH1 and FKH2 deletion on chromosomal structure, including the ability for FKH1 and
FKH2 to rescue the deletion replication phenotype after many generations, will address this
concept.
Binding specificities of Forkhead proteins are altered spatially and temporally: A key
finding of this work is in furthering our understanding of how a protein exhibits cell cycle-
dependent binding at certain regulatory classes and cell cycle-independent binding at
others. Fkh1 has the predominant role in regulation of replication timing compared with
Fkh2, as Fkh2 deletion leads to no substantial deregulation of replication timing, yet Fkh2
114

is able to partially complement in the absence of FKH1
72
. This is partially explained by
examination of Fkh1 and Fkh2 binding; Fkh1 binds early origins independently of Fkh2, yet
Fkh2 only binds origins that also exhibit binding of Fkh1, implying that in wild type cells
Fkh2 is dependent on Fkh1 for binding
99
. We show here that the ability of Fkh1 to dimerize
is related to its binding at early origins required of Fkh1 to regulate early origin timing. The
requirements for Fkh2 binding at early origins are unknown, particularly as to why Fkh2
does not bind independently of Fkh1. It is possible that in wild type cells, Fkh2 alone is
sufficient to impart early initiation of origins, or that dimerization is a key requirement of
Forkhead binding at origins and Fkh2 preferentially forms heterodimers with Fkh1 over
homodimers. Furthermore, it is possible that cell cycle-regulated binding of Forkhead
proteins to early origins is regulated by cell cycle machinery; thus, we are investigating the
dependence of Fkh1 and Fkh2 binding on kinase activity. A plausible explanation is that cell
cycle-regulated modification of Fkh1 and Fkh2 is required for dimerization which either
allows the proteins to bind early origins or stabilizes certain interactions, such as that with
ORC, once bound. Answers to these questions will yield valuable insight into the molecular
mechanisms that control cell cycle-specific chromatin binding.
Fkh-dsm as a tool for fundamental investigations: The alleles fkh1-dsm and fkh2-dsm
are new tools with which to investigate replication initiation and other fundamental
cellular processes. Both alleles are loss of function, deficient in establishing early firing
(and in the case of Fkh1-dsm we also show a lack of binding at these origins), but both are
able to perform their transcriptional functions of regulating the CLB2-cluster. These
proteins can be further modified, for example by tethering to other proteins, and used for
additional studies and characterizations.
115

Residual activities of Fkh-dsm in replication timing: Replication timing in fkh1-dsm and
fkh2-dsm are not entirely deficient, coming close but ultimately failing to reach the level of
deregulation seen with complete deletion of FKH1 and FKH2, indicating that a certain
amount of residual Forkhead-mediated regulation of replication initiation may still occur.
This could be representative of residual Forkhead dimerization function, as the domain
swap mutants exhibit greatly reduced, but not entirely abolished, ability to dimerize when
analyzed in pull-down assays. This could also represent additional unknown stimulatory
function, or functions, of Forkhead proteins in regulation of replication initiation, perhaps
relating to its ability to physically interact with ORC
72
. In either or both cases, binding of
Fkh1-dsm is not detected at origins, but it remains a possibility that whatever functions
Fkh1 performs can be completed rapidly enough at a subset of origins in a subset of cells
that transient association of Fkh1-dsm is sufficient to cause initiation at detectable levels,
however low.
Alternative mechanism of Forkhead regulation of replication timing or genome
architecture: The possibility remains that Forkhead proteins serve not to recruit
replication origins into multi-origin clusters, but rather serve to stabilize
interchromosomal interactions established by an unknown mechanism. In this scenario, in
G1 phase Forkhead protein dimerization may be induced by cell cycle or replication
machinery, bind multiple origins and lock them in place. Additionally, they may serve to
recruit initiation factors once bound, potentially either as dimers or transiently-bound
monomers.
116

Future directions: Additional experiments will provide more mechanistic details of
Fkh1/2 binding at early origins and formation of long range replication clusters. Replacing
a low-affinity Fkh1 binding site at an early origin to a higher-affinity site found at a CLB2-
cluster promoter may lead to increased binding, in which case we can assess replication
activity and determine if binding of fkh1-dsm is sufficient to cause early firing of the origin.
A lack of an increase in binding may indicate a separate defect, for example, a failure of
fkh1-dsm to interact with pre-RC factor(s) that may be responsible for origin-specific Fkh1
binding in late G1 and S phase.
Additionally, replacement of Fkh1 binding sites at an early origin with LexA binding sites is
expected to delay replication timing of the origin. Introduction of LexA/Fkh1 or LexA/fkh1-
dsm fusion proteins (LexA-Fkh1 and LexA-fkh1-dsm, respectively) will also address the
question of whether Fkh1 binding alone is sufficient to advance origin timing. Performing
this experiment in a Fkh1-Myc, fkh2Δ and performing a trans-chromatin
immunoprecipitation against the Myc epitope will potentially lead to evidence of in vivo
Fkh1 dimerization.
As mentioned in Results, the discrepancy between in vivo chIP data and in vitro EMSA data
showing fkh1-dsm binding could be attributed to a lack of post-translational modifications
in the EMSA experiment. This can readily be addressed by one of two methods. First, EMSA
could be repeated as described with an added first step of incubating His-Fkh1-DBD in
yeast whole cell extract to allow other factors to act upon the protein before isolation,
incubation with DNA and analysis by PAGE. Second, EMSA could be performed with Fkh1
isolated from yeast as done in the pull-down assay. Either of these approaches can be
117

performed with whole cell extract from asynchronous or synchronized culture to address
cell cycle-specific binding of Fkh1 at, for example, early origins.
Finally, in collaboration with Carolina Dantas and Remo Rohs, we are investigating in
greater detail Fkh1 binding motifs at early origins and elsewhere. Preliminary work
indicates significant differences at Fkh1 binding sites at early origins.

118

4.4 Methods

Yeast strains and methods: Domain swap mutations were introduced to plasmid-borne
FKH1 and FKH2 (and flanking genomic regions) using QuickChange Lightning Mutagenesis
Kit to produce fkh1-dsm and fkh2-dsm, respectively. These alleles were cloned into an
integrating pRS405 vector and transformed into a fkh1Δfkh2ΔC, BrdU-integrating, bar1Δ
background. The same was done for wild type FKH1 and FKH2 to be used as controls.
BrdU-IP-Seq: BrdU-IP-Seq in hydroxyurea (HU) was performed as previously described
(Knott et al., 2012). 50 mL cells in YEPD were grown to an OD600 of 0.5 and arrested with
(X) α-factor for 4 hours. Cells were released from G1 arrest with (X) pronase into 0.2 M HU
and (X) BrdU for 1 hour, then treated with (X) sodium azide. Cells were washed with cold
TBS and pelleted. DNA was extracted, sonicated with a Covaris S2 to ~300 bp, and
immunoprecipitated in 500 µL for two hours with 1:100 anti-BrdU; IP DNA was isolated
with Dynabeads and prepared for Illumina 50-read paired end sequencing. Quality control
was performed using a Bioanalyzer and qPCR.
Preprocessing of sequencing data: Sequencing reads were aligned to the S. cerevisiae
genome using Bowtie2 allowing up to 2 mismatches. Multiply aligned reads and those
mapping to repetitive regions were filtered out, and remaining reads were binned into non-
overlapping 50-bp bins. Reads across experiments were median smoothed over (X) kb and
quantile normalized. Smoothing was repeated.
Agar Scarring Assay: Cells were grown until stationary phase. Drop-out media was used
to prevent plasmid loss and integrated gene pop-out. 10 µL cells were placed on agar plates
119

and allowed to grow for (X) days at 30°C. An image of the plate was captured. Cells were
then gently rinsed off the plate with water, and a second image was captured.
ChIP-chip: ChIP-chip was performed as described previously (Ostrow et al., 2014).
Preprocessing of microarray data: IP and total DNA were labeled with Cy5 and Cy3,
respectively. 1 µg each were hybridized to Nimblegen microarrays (~60 bp oligonucleotide
for every ~80 bp of unique genomic sequence) over ~72 hours. Microarrays were washed
and scanned using an Axon 4100A Scanner. Hybridization and washing steps were carried
out according to Nimblegen protocol. Nimblegen software was used to preprocess captured
array images, and data was processed using MA2C software. Output was binned over 50-
bp, median smoothed over 1 kb, quantile normalized, and re-smoothed.
Analysis of global replication and Fkh1 occupation at features: Two dimensional
matrices were constructed in which columns were centered around a specific genomic
feature and rows represent individual occurrences of the feature. Columns were summed
and averaged and plotted as heat maps showing mean signal per 50 bp bin over 5 KB
regions.
Peak calling and differential origin firing: Peak calling was performed using MACS.
Resulting peaks were analyzed by DiffBind (R BioConductor) for differential peak heights.
Analysis of Fkh1 dimerization using pull-down assay: Whole cell extract (WCE)
containing either Fkh1-Myc or fkh1-dsm-Myc was subjected to anti-Myc
immunoprecipitation. Purified Fkh1-Myc or fkh1-dsm-Myc was incubated with Fkh1 DNA
Binding Domain (Fkh1-DBD) or fkh1-dsm DBD. Samples were again immunoprecipitated
120

using an antibody against anti-Myc, and the resulting purified samples were run on an SDS-
PAGE gel and blotted with anti-Fkh antibody.

121

Supplemental Figures

Figures S3.1.1 – 3.1.16. Genome-wide analysis of Fkh1 and Fkh2 chromatin
binding. Plots show averaged ChIP-chip signal from three experimental replicates along
each chromosome, with enriched regions plotted in purple. The antibody and strain
genotype used for each experiment are indicated to the left of each panel. The
corresponding strains from top to bottom are: CVy43, ZOy1, CVy138, CVy139, ZOy3, ZOy4,
and CVy43. Triangles on the bottom panel indicate the position of determined binding sites
as described in the text, color-coded by classification.
Figure S3.1.1

122

Figure S3.1.2

123

Figure S3.1.3

124

Figure S3.1.4

125

Figure S3.1.5

126

Figure S3.1.6

127

Figure S3.1.7

128

Figure S3.1.8

129

Figure S3.1.9

130

Figure S3.1.10

131

Figure S3.1.11

132

Figure S3.1.12

133

Figure S3.1.13
134

Figure S3.1.14

135

Figure S3.1.15

136

Figure S3.1.16

137

Figures S3.2.1 – 3.2.16. Cell cycle binding of Fkh1/2 genome-wide. Plots show
averaged ChIP-chip signal from three experimental replicates along each chromosome,
with enriched regions plotted in purple. The cell cycle arrest for each experiment is
indicated to the left of each panel.
Figure S3.2.1

138

Figure S3.2.2

139

Figure S3.2.3

140

Figure S3.2.4

141

Figure S3.2.5

142

Figure S3.2.6

143

Figure S3.2.7

144

Figure S3.2.8

145

Figure S3.2.9

146

Figure S3.2.10

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Figure S3.2.11

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Figure S3.2.12

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Figure S3.2.13

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Figure S3.2.14

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Figure S3.2.15

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Figure S3.2.16

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Abstract (if available)

Abstract Forkhead box (FOX) transcription factors regulate a wide variety of cellular functions in higher eukaryotes, including cell cycle control and developmental regulation. In Saccharomyces cerevisiae, Forkhead proteins Fkh1 and Fkh2 perform analogous functions, regulating genes involved in cell cycle control, while also regulating mating-type silencing and switching involved in gamete development. As described here, we revealed a novel role for Fkh1 and Fkh2 in the regulation of replication origin initiation timing, which, like donor preference in mating-type switching, appears to involve long-range chromosomal interactions, suggesting roles for Fkh1 and Fkh2 in chromatin architecture and organization. To elucidate how Fkh1 and Fkh2 regulate their target DNA elements and potentially regulate the spatial organization of the genome, we undertook a genome-wide analysis of Fkh1 and Fkh2 chromatin binding by ChIP-chip using tiling DNA microarrays. Our results confirm and extend previous findings showing that Fkh1 and Fkh2 control the expression of cell cycle-regulated genes. In addition, the data reveal hundreds of novel loci that bind Fkh1 only and exhibit a distinct chromatin structure from loci that bind both Fkh1 and Fkh2. The findings also show that Fkh1 plays the predominant role in the regulation of a subset of replication origins that initiate replication early, and that Fkh1/2 binding to these loci is cell cycle-regulated. Finally, we demonstrate that Fkh1 and Fkh2 bind proximally to a variety of genetic elements, including centromeres and Pol III-transcribed snoRNAs and tRNAs, greatly expanding their potential repertoire of functional targets, consistent with their suggested role in mediating the spatial organization of the genome. ❧ Sequence comparison and structural modeling of Fkh1 and Fkh2 with mammalian FOXP family proteins reveals key homologies that suggest Fkh1 and Fkh2 are able to form domain swapped dimers, providing a possible mechanism by which Fkh1 and Fkh2 bind distal origins and sustain replication foci through dimerization in trans. Altering residues thought to be involved in dimerization, we created domain swap mutants (dsm) Forkhead proteins. We show that wild type Fkh1 dimerizes, and that this is diminished in fkh1-dsm. BrdU-IP-Seq analysis of global replication in fkh-dsm reveals a null-like phenotype, yet Forkhead regulation of transcription appears intact. Assessment of chromatin binding using ChIP-chip shows that fkh1-dsm sufficiently binds at the promoters of CLB2-cluster promoters, while binding at early origins is ablated, indicating a role of the mutated residues in cell cycle-specific binding of Fkh1 at replication origins. Finally, we find that the Forkhead binding motifs found at early origins differ from those found elsewhere, supporting a role of alternative sequence specificities in Forkhead binding and regulation of replication origins.

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Creator Ostrow, Andrew Zachary (author)

Core Title Forkhead transcription factors regulate replication origin firing through dimerization and cell cycle-dependent chromatin binding in S. cerevisiae

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publication Date 06/03/2015

Defense Date 10/13/2014

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

Tag chromatin,chromatin organization,dimer,dimerization,dimers,Fkh1,Fkh2,forkhead,genome architecture,genome structure,OAI-PMH Harvest,replication foci,replication origins,replication timing,transcription factors

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Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Aparicio, Oscar M. (committee chair), Chen, Lin (committee member), Michael, Matthew (committee member), Rohs, Remo (committee member)

Creator Email aostrow@usc.edu,zacostrow@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c3-520750

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Legacy Identifier etd-OstrowAndr-3093.pdf

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