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Subnuclear localization of replication origins is controlled by Fkh1-dependent recruitment of DDK to origins in S. cerevisiae

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Subnuclear localization of replication origins is controlled by Fkh1-dependent recruitment of DDK to origins in S. cerevisiae

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Content Copyright 2021 Haiyang Zhang
Subnuclear localization of replication origins is controlled by Fkh1-
dependent recruitment of DDK to origins in S. cerevisiae
by
Haiyang Zhang
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)
May 202 1
ii

Acknowledgements

First and foremost, I sincerely thank my amazing advisor Dr. Oscar Aparicio. Having the
honor to join his lab is truly the luckiest thing that happened in my life. Pursuing a Ph.D. in
graduate school is much more difficult than I imagined it would be. But with his incredible
mentoring, the last six years have become a productive and rewarding enjoyable journey.
Dr. Oscar Aparicio is a perfect role model in both the scientific field and daily life for me.
For doing research, he taught me how to develop a habit of critical thinking and conduct
demonstrations rigorously before making any steady statements. For instance, 'no controls,
no experiments'; 'probably ok is not ok'. These are the inspiring philosophies that I can
benefit from in my life not only in research. Dr. Aparicio's passionate attitude for research
study also inspires me significantly all the time. His impressive perspectives on
interpreting data results and those million-dollar ideas of investigating for more
explorations helped me to develop the mind of how to think about science. He encourages
me to move out of my comfort zone and challenge myself to explore more possibilities in
research directions. I certainly will not be able to find my real interest without his generous
supports and suggestions. Being a member of Dr. Oscar Aparicio group is the most precious
experience for me that I will appreciate for my entire life. I also thank the other member of
my committee, Dr. Irene Chiolo, Dr. Matthew Michael, and Dr. Sean Curran. They provided
me numerous constructive suggestions and feedback, which has been most appreciated. It
is a pleasure working with everyone. Dr. Chiolo has been instrumental in Chapter 2 in this
dissertation and taught me how to analyze imaging data appropriately. I would also like to
thank my lab mates and I enjoy every minute of being with them in our lab over the years.
Yiwei He, Meghan Petrie, Dr. Zac Ostrow, Dr. Sandra Villwock, Dr. Jared Peace, Dr. Joanna E.
iii

Haye, Dr. Kankana Ghoshal. Our lab manager Yan Gan treats me as a family member. I am
really grateful for this cherished friendship with her. Lastly, I would like to thank my
family. They have been very supportive of my pursuits. I would like to thank my wife
Shiyue Huang and her company for these years in the US. She makes me believe that we can
conquer any difficulties together especially this year with the pandemic. I enjoy seeing both
of us moving forward to a better future together.
iv

TABLE OF CONTENTS
Acknowledgements ................................................................................................................................................. ii
List of Figures ............................................................................................................................................................. v
Abstract ........................................................................................................................................................................ vi
Introduction ................................................................................................................................................................ 1
Chapter I: Dynamic relocalization of replication origins by Fkh1 requires execution of DDK
function and Cdc45 loading at origin ............................................................................................................... 7
Introduction ...................................................................................................................................................... 7
Materials and Method ................................................................................................................................... 9
Results .............................................................................................................................................................. 15
Discussion ....................................................................................................................................................... 35
Chapter II: Fkh1 domain-swapping motif regulates subnuclear localization of replication
origin suggesting a role of recruitment of Dbf4 ....................................................................................... 41
Introduction .............................................................................................................................................. - 41 -
Materials and Methods .............................................................................................................................. 44
Results .............................................................................................................................................................. 45
Discussion ....................................................................................................................................................... 53
Appendix ................................................................................................................................................................... 56
References ................................................................................................................................................................ 64
Supplemental Figures .......................................................................................................................................... 79

v

List of Figures
Figure 1 Fkh1-induced origin activation re-positions a sub-telomeric origin in G1 phase…16
Figure 2 Normal dosage of Fkh1 is sufficient to relocalize ARS305V-R and advance its firing
time…………………..…………………………………………………………………………………………………………20
Figure 3 Fkh1 determines early origin positioning globally……………………………………………22
Figure 4 Origin localization in G1 is DDK regulated………………………………………………………..26
Figure 5 DDK regulation of origin localization reflects its phosphorylation of Mcm4 and
consequent Cdc45 loading………………………………………………………………………………………….…29
Figure 6 Origin mobility increases with origin relocalization………………………………….………34
Figure 7 Model of origin localization linked to initiation…………………………………………….…..38
Figure 2.1 Domain swap mutant of Fkh1 fails to relocalize ARS305 to the center of the
nucleus in G1………………………………………………………………………………………………………………..46
Figure 2.2 DBF4-FKH fkh1Δ but not DBF4-fkh-dsm phenocopies WT in both replication
timing and subnuclear localization………………………………………………………………………….…….48
Figure 2.3 Deletion of CTF19 in fkh1Δ background shows an additive replication
phenotype………………………………………………………………………………………………………………..…..52
Figure 1-figure supplement 1. Fkh1-induction is required to re-position a sub-telomeric
origin in G1 phase………………………………………………………………………………………………………...79
Figure 2-figure supplement 1. Normal dosage of Fkh1 is sufficient to advance firing time of
ARS305
V-R
…………………………………………………………………………………………………………………...80
Figure 3-figure supplement 1. Fkh1 determines early origin positioning globally……….….81
Figure 4-figure supplement 1. Fkh1 binds origins independently of Cdc7 function………...83
Figure 5-figure supplement 1. CDK activity is dispensable for origin localization in G1…..84
vi

Abstract
Chromosomal DNA elements are organized into spatial domains within the eukaryotic
nucleus. Sites undergoing DNA replication, high-level transcription, and repair of double-
strand breaks coalesce into foci, although the significance and mechanisms giving rise to
these dynamic structures are poorly understood. In S. cerevisiae, replication origins exhibit
characteristic subnuclear localizations prior to S phase that anticipate their initiation
timing and/or efficiency during S phase: origins found within the nuclear interior in G1
phase initiate early and efficiently in S phase while origins found associated with the
nuclear periphery in G1 phase initiate later and less efficiently. Here, we link localization of
replication origins in G1 phase with Fkh1 activity, which is required for their early
replication timing. Using a Fkh1-dependent origin relocalization assay, we determine that
execution of Dbf4-dependent kinase function, including Cdc45 loading, results in dynamic
relocalization of a replication origin from the nuclear periphery to the interior in G1 phase.
Origin mobility increases substantially with Fkh1-driven relocalization. Specifically,
domain-swapping motif of Fkh1 is crucial for the regulation of both replication timing and
subnuclear location, which is required for Dbf4 recruitment. The replication timing of
centromeric origins is controlled by Ctf19 but independent of Fkh1. These findings provide
novel molecular insight into the mechanisms that govern dynamics and spatial
organization of DNA replication origins and possibly other functional DNA elemen
- 1 -

Introduction
An overview of eukaryotic replication and dynamics of replication origin

DNA replication is highly conserved from budding yeast to mammalian cells in
terms of the initiation of eukaryotic replication origins. Precise and complete DNA
replication maintains the genome stability and high fidelity of cell division and
proliferation (Cvetic and Walter, 2005). The process, faithful duplication of the
chromosome, is regulated temporally and spatially to minimize mutagenesis. During G1
phase, the replication program is established through the recruitment of initiation factors.
In Saccharomyces cerevisiae, autonomously replicating sequences (ARSs) -100-200 base
pair (bp) chromosomal DNA sequence- is bound by Origin Recognition Complex (ORC)
initially, which is considered as the first step of origin initiation (Bell and Dutta, 2002).
Subsequently, ORC serves as a scaffold to stimulate the recruitment of following initiation
factors such as Cdc6 and Cdt1 with inactive minichromosome maintenance (Mcm2-7, or
MCM) helicase. This assembly of the pre-replication complex (pre-RC) on the origin is
referred to as a ‘licensing’ step. Then, the licensed origin is activated by Dbf4-dependent-
kinase (DDK) and cyclin-dependent-kinase (CDK) in G1-to-S-phase transition (Labib,
2010). DDK phosphorylates the MCM, particularly the Mcm4 and Mcm6 subunit, and
recruits initiation factors Cdc45 and Sld3 (Deegan et al., 2016). Then, CDK phosphorylates
Sld2 and Sld3 to load GINS, which activates MCM helicase and completes the replisome
components assembly (Heller et al., 2011). Ultimately, DNA unwinding proceeds
- 2 -

bidirectionally along the chromosome until it encounters neighboring replication fork
(Machida et al., 2005).

Although replication origins share the pre-RC assembly process, the replication
timing and activities of different origins are distinct along the whole genome. Chromosomal
structure and subnuclear localization are highly correlated with the regulation of
replication origin initiation timing (Aparicio, 2013; Rhind and Gilbert, 2013). Importantly,
the series of events after the establishment of replication timing program do not occur
simultaneously among all origins. In addition, many groups found that some regions such
like centromeres and telomeres showed a remarkable and specific replication efficiency
and timing in the genome (Pohl et al., 2012). Based on whole-genome studies, early firing
origins are often found near the centromere (Heun et al., 2001a; Heun et al., 2001b). In
contrast, telomeric and subtelomeric regions are late replicating or dormant because of the
negative contribution from nearby heterochromatic structure. Not only the various local
chromatin environment, but also epigenetic regulation of gene expression significantly
influences the replication timing (Hiratani et al., 2009). With the regulation of histone
modifiers and nucleosome remodelers, chromatin architecture is subject to covalent
modifications leading to the changes in chromatin dynamics and its biophysical properties
such as mass density and diffusion rate (van Steensel et al., 2011). Thus, replication timing
can be affected by post-translational modifications which are able to alter both local
chromatin folding and 3D genome organization.
- 3 -

(Heller et al., 2011)
Studies of Rif1 (Rap1-interacting factor 1) showed that Rif1 is important in
regulating replication origin timing in both yeast and mammalian cells (Hayano et al., 2012;
Cornacchia et al., 2012; Lian et al., 2011; Yamazaki et al., 2012). Rif1 binds telomeres and
subtelomere regions by direct interaction with Rap1, a telomere sequence binding protein,
in S. cerevisiae (Hardy et al., 1992). In S. pombe, Rif1 interacts with Taz1 to bind to the
telomeres (Kanoh and Ishikawa, 2001). In the fission yeast, deletion of rif1 results in an
- 4 -

early initiation pattern of many late origins in subtelomeric as well as some internal
chromosomal loci. Meanwhile, a delayed initiation was observed in many early replication
origins, including pericentric origins (Hayano et al., 2012). Many research groups
discovered that S. cerevisiae Rif1 protein controls DNA replication genome-wide through
PP1-mediated dephosphorylation of the MCM complex early in the cell cycle (Hiraga et al.,
2012; Peace et al., 2014). Therefore, Rif1-PP1 counteracts the process of DDK-mediated
phosphorylation of Mcm4, which determines initiation globally in the genome. Our lab
showed that Rif1 plays a global role in the regulation of late/dormant origins by analyzing
replication genome-wide in cells lacking Rif1 function (Peace et al., 2014). The results are
consistent with the previous conclusion that Rif1 globally regulates replication timing and
is independent of telomere proximity.

Both centromeres and telomeres have characteristic replication timing patterns,
early and late, respectively. Specifically, the deletion of a centromere in Candida albicans
resulted in a formation of a new centromere which is away from the original CEN site
(Koren et al., 2010). Relocating a centromere into a normally late-firing region caused an
early firing of this region (Pohl et al., 2012). The data is consistent with the idea that
centromeric factors impart early timing to the regions in which they reside. Recently,
studies in both Schizosaccharomyces pombe and S. cerevisiae showed that the early firing of
the CEN region is because of the DDK-recruitment to centromeric origins (Hayashi et al..,
2009). For instance, the live cell imaging in budding yeast revealed that both Dbf4 and Cdc7
accumulate at kinetochores and spindle pole bodies in early G1 phase (Natsume et al.,
2013). Both replication timing and origin subnuclear localization were significantly
- 5 -

affected by the deletion of the chromosome kinetochore protein Ctf19. Their data indicate
that Ctf19 facilitates the Dbf4-dependent kinase (DDK) accumulation at kinetochores in G1
and stimulates the early initiation of the pericentromeric region in S phase.

Our lab recently discovered that Fkh1/2 establishes origin timing program and
regulates long-range clustering of origins in S. cerevisiae (Knott et al., 2012). BrdU-IP-seq
was conducted for replication timing analysis. In addition, the deletion of FKH2 alone
shows no replication effect, while fkh1Δ cells showed a slightly less intense replication
phenotype when compared to the fkh1Δfkh2Δ double mutant. Therefore, Fkh1 exhibits the
predominant role in replication timing regulation and Fkh2 partially compensates in the
absence of Fkh1. Furthermore, out of 352 total origins, 106 were defined Fkh-activated as
their early initiation became later; while 82 were termed Fkh-repressed as these late
origins became earlier firing. Gene expression analysis by RNA-Seq has exhibited that the
Fkh replication regulation is not caused by local changes in transcription near origins. This
was further supported by the result that overexpression of C-terminally truncated Fkh2 in
the double mutant rescued most of the transcriptional deregulation but did not rescue
origin timing deregulation.

Chromosome conformation capture analyses showed that early-firing origins cluster
spatially in G1-phase. (Duan et al., 2010) Our lab discovered a correlated result and found
the clustering is regulated by Fkh1/2 (Knott et al., 2012). Meanwhile, early and late origins
tend to occupy in different locations in the nucleus. Specifically, early-firing origins
typically are enriched in the interior of the nucleus; while late-firing origins are usually
- 6 -

crowded around the nuclear periphery (Heun et al., 2001a; Heun et al., 2001b). Together,
Fkh1/2 is considered to contribute to both replication timing and subnuclear location of
origins. Recently, My research in the Aparicio lab examined how origin stimulation by Fkh1
determines subnuclear origin localization. Chapter 1 presents the results of this
investigation, which found that origin relocalization from the nuclear periphery is
regulated by Fkh1 through recruitment of DDK.
The Aparicio lab previously identified a domain-swapping motif in Fkh1 and Fkh2 is
critical for replication timing regulation (Ostrow et al., 2017). Mutations at specific residues
were introduced to create “domain-swap minus” (dsm) alleles of FKH1. BrdU-IP-seq
analysis was conducted and showed a significant delay in replication timing fkh1-dsm
mutant cells. Therefore, the data further supports the mechanism that Forkhead proteins
facilitate the establishment of physical communications between origins for clustering in
advancing replication. Thus, we examined the function of the domain-swapping motif in
Fkh1 on regulating origin subnuclear localization in G1 phase, in relation to its function in
replication timing regulation. Chapter 2 presents the results of this investigation, which
showed that domain-swapping motif in Fkh1 is required for the recruitment of Dbf4 based
on the origin repositioning assay conducted on fkh1-dsm mutant cells.

- 7 -

Chapter I

Dynamic relocalization of replication origins by Fkh1 requires
execution of DDK function and Cdc45 loading at origins
From Zhang et al., 2019

Introduction
The spatial organization of chromosomal DNA elements within the nucleus is thought to
derive from and contribute to the regulation of their activity (reviewed in (Shachar and
Misteli, 2017). For example, euchromatin and heterochromatin represent distinct forms of
chromatin that are distinguished by their levels of transcriptional activity, replication
timing, and subnuclear localization (reviewed in (Caridi et al., 2017). Chromosomes
partition into subdomains ranging from hundreds to thousands of kilobases in length that
preferentially self-associate and are consequently referred to as topologically associated
domains (TADs) (reviewed in (Zhao et al., 2017). TAD boundaries correlate closely with
replication timing domains, suggesting that replication timing is determined or influenced
by this domain structure and/or vice-versa.

In budding and fission yeast, specific mechanisms defining replication timing are linked
with chromosomal domain organization (reviewed in (Aparicio, 2013; Yamazaki et al.,
2013). Rif1, which is highly enriched at telomeres, is globally responsible for delayed
- 8 -

replication timing of subtelomeric domains as well as internal late-replicating domains
(Hafner et al., 2018; Hayano et al., 2012; Peace et al., 2014; Tazumi et al., 2012). Rif1 acts
by directly antagonizing replication initiation triggered by Dbf4-dependent kinase (DDK)
phosphorylation of MCM helicase proteins (Dave et al., 2014; Hiraga et al., 2014; Mattarocci
et al., 2014). Against this inhibitory backdrop, specific origins are selected for early
activation by mechanisms involving recruitment of Dbf4 (Dfp1 in fission yeast), which is
one of several initiation proteins present in limited abundance and thus rate-limiting for
origin firing (Mantiero et al., 2011; Patel et al., 2008; Tanaka et al., 2011; Wu and Nurse,
2009). In S. pombe, Dfp1 is recruited to kinetochores through heterochromatin protein
Swi6 (Hayashi et al., 2009), and in S. cerevisiae, kinetochore protein Ctf19 recruits Dbf4 to
stimulate firing of origins within ~25kb of the centromere (Natsume et al., 2013), thus
ensuring early centromere replication by distinct mechanisms regulating DDK activity. In
S. cerevisiae, Fkh1 and/or Fkh2 (Fkh1/2) recruits Dbf4 to many origins distributed
throughout chromosome arms, thereby ensuring timely and complete chromosome
replication, particularly of centromere-distal regions (Fang et al., 2018; Knott et al., 2012;
Looke et al., 2012; Ostrow et al., 2014).

Chromosome conformation capture experiments suggest that early-firing origins cluster
spatially in G1-phase prior to initiation and this clustering is dependent on Fkh1/2 (Duan
et al., 2010; Knott et al., 2012). These studies also indicated that early origins generally
occupy a distinct space than late origins. Further studies suggest that Fkh1/2 are enriched
at TAD boundaries and control contacts among origins within TADs (Eser et al., 2017). The
distinct spatial distributions suggested by these recent studies are in accord with earlier
- 9 -

studies that examined the subnuclear distribution of individual origins by fluorescence
microscopy. These seminal studies from Heun and Gasser showed that late-firing origins
typically associate with the nuclear periphery during G1 phase whereas early-firing origins
typically are found in the nuclear interior during G1 (Heun et al., 2001a; Heun et al.,
2001b). Despite the observed correlations between origin localization in G1 and firing time
in S, the main origin timing determinants mentioned above had not been elucidated and
have not been examined for their impact on subnuclear localization of replication origin. In
this study, we examined how origin stimulation by Fkh1 determines subnuclear origin
localization. Our results suggest that origin relocalize from the nuclear periphery upon
execution of the DDK-dependent step of replication initiation, which is stimulated by Fkh1.
This may represent the initial stages in the coalescence of replication origins into clusters
that will become replication factories.

Materials and Method
Plasmid constructions Plasmids were constructed using Gibson Assembly kit (SGI
cat#GA1200) unless otherwise indicated. Restriction enzymes, T4 DNA ligase, and Klenow
were from New England Biolabs and used according to their protocols. Mutagenesis was
carried out using QuikChange Lightning Multi kit (Agilent cat#210515); sequence changes
were confirmed by DNA sequencing (Retrogen Inc.). STBLII cells were used for maintenance
of plasmids containing tandem repeats (Invitrogen cat#10268019). Primer sequences for
plasmid constructions are given in Table 1. NUP49-GFP was PCR-amplified from pUN100-
GFP*-Nup49 (from V. Doye) using primers Nup49-GFP-F and Nup49-GFP-R and subcloned
- 10 -

into XhoI+SacI-digested vectors pRS403 and pRS404 (Sikorski and Hieter, 1989) to yield
p403-Nup49-GFP and p404-Nup49-GFP, respectively. Primers ADE2-up-F and ADE2-int-R
and separately ADE2-farup-F and ADE2-up-R were used to amplify sequences of ADE2 for
targeting and as a selectable marked; these were inserted into pbluescriptKS+ to create
pblueKS-ADE2target. TetR-Tomato was PCR-amplified from plasmid p402-TetR-Tomato
(from S. Sabatinos) using primers TetR-Tom-F and TetR-Tom-R and inserted into PacI-
digested pblueKS-ADE2target to generate pTetR-Tom-ADE2. 2.1kbp KpnI-SacI fragment
containing LacI-GFP was subcloned from pAFS135 (from J. Bachant) into pRS404 digested
with same enzymes to create p404-LacI-GFP. Plasmids containing tetO (pGS004 from J.
Bachant) or lacO (pJBN164 from J. Bachant) arrays were modified by introduction of
genomic sequences to target integration near different origins. The following primer pairs
were used to generate sequences adjacent to the indicated origins (with the corresponding
chromosomal coordinates given in parentheses): primers ARS501-tetO-F and ARS501-tetO-
R for ARS501 (V:547812-548329), primers ARS1103-tetO-F and ARS1103-tetO-R for
ARS1103 (XI:54673-54996) and primers ARS1303-tetO-F and ARS1303-tetO-R for ARS1303
(XIII:31983-32247), and these were inserted into KpnI+ClaI-digested pGS004 yielding
pARS501-tetO, pARS1103-tetO, and pARS1303-tetO, respectively. Likewise, the following
primer pairs were used to generate sequences adjacent to the indicated origins: primers
ARS710-lacO-F and ARS710-lacO-R for ARS710 (VII:204305-204831), primers ARS718-
lacO-F and ARS718-lacO-R for ARS718 (VII:422375-423281), and primers ARS1018-lacO-F
and ARS1018-lacO-R for ARS1018 (X:539662-540395), and these were inserted into
XhoI+KpnI-digested pJBN164 yielding pARS710-lacO, pARS718-lacO, and pARS1018-lacO,
respectively. Two adjacent regions near ARS305 were PCR-amplified using primer pair NotI-
- 11 -

ARS305-5  and XhoI-ARS305-5  (III:37283-37778) and primer pair NotI-ARS305-3  and
KpnI-ARS305-3  (III:37779-38282) and digested with NotI and XhoI and NotI and KpnI,
respectively; these fragments were ligated into pRS404 digested with XhoI and KpnI. The
XhoI-KpnI fragment was subcloned by digestion and ligation into pJBN164 digested with
same enzymes to yield pARS305-lacO. Plasmids p501Δ-ARS305-ΔACS and p501Δ-ARS305-
∆2BS were created by mutagenesis of p501Δ-ARS305 (Peace et al., 2016) with primers
ARS305-∆ACS-mut1, ARS305-∆ACS-mut2 and ARS305-∆2BS-mut1, and ARS305-∆2BS-
mut2, respectively. Plasmid p404-ars305∆-BInc was constructed as described for p306-
ars305∆-BrdU-Inc (Zhong et al., 2013) except that p404-BrdU-Inc (Viggiani and Aparicio,
2006) was used instead of p306-BrdU-Inc. p404-ars305∆-BInc was digested with PmlI and
KpnI, blunted-ended with Klenow, and ligated with T4 DNA ligase to remove the TRP1
selectable marker, yielding p400-ars305∆-BInc. The 1.5 kb SalI-SpeI fragment containing
the KanMx cassette from pFA6-KanMx (Longtine et al., 1998) was ligated into SalI-SpeI-
digested p400-ars305∆-BInc, creating pKanMx-ars305∆-BInc. The cdc45-1 allele was PCR-
amplified from strain YB298 (from B. Stillman) with primers Cdc45-F and Cdc45-R and
inserted into SacI+KpnI-digested pRS406 (Sikorski and Hieter, 1989) to create p406-cdc45-
1.

Yeast strain constructions All strains are congenic with SSy161, derived from W303-1a
(RAD5) (Viggiani and Aparicio, 2006); complete genotypes are given in Table 2. Strain
constructions were carried out by genetic crosses or lithium acetate transformations with
linearized plasmids or PCR products generated with hybrid oligonucleotide primers having
- 12 -

homology to target loci (Ito et al., 2001; Longtine et al., 1998); primer sequences for strain
constructions are given in Table 1. Genomic alterations were confirmed by PCR analysis or
DNA sequence analysis as appropriate.

FKH1 was deleted using primers Fkh1-up and Fkh1-down to amplify KanMx selectable
marker from pFA6-KanMx (Longtine et al., 1998). FKH2 was replaced by fkh2-dsm in two
steps: first, FKH2 was entirely replaced with URA3 (C. albicans) using pAG61 (Addgene), and
the resulting strain was transformed with fkh2-dsm DNA from p405-fkh2-dsm (Ostrow et al.,
2017) followed by selection on 5-FOA. GAL-FKH1 was introduced using p405-GAL-FKH1
and FKH1 was FLAG-tagged as described previously (Peace et al., 2016). ARS501 was
replaced by ARS305 or mutant versions of ARS305 by transformation with p501∆-ARS305,
p501∆-ARS305-∆ACS, or p501∆-ARS305-∆2BS as described previously (Peace et al., 2016).
BrdU incorporation cassette was introduced, replacing ARS305, by transformation with
BglII-digested p404-ARS305-BrdUInc. CLB5 and CLB6 deletion alleles were introduced by a
series of crossings to previously described strains (Gibson et al., 2004). The cdc7-as3 allele
was introduced as described previously (Zhong et al., 2013); cdc7-4 was back-crossed from
H7C4A1 (from L. Hartwell) into the W303 background four times, with the final cross to
HYy151. MCM4-DD/E+DSP/Q (referred to in text as MCM4-14D) was introduced by
transformation with PacI-digested pJR179 (from S.P. Bell). The cdc45-1 allele was
introduced by crossing with strain YB298 (from B. Stillman) or by pop-in/pop-out with BglII-
digested p406-cdc45-1. The dbf4∆C allele was constructed by insertion of a non-sense codon
with the KanMx cassette from pFA6-KanMx (Longtine et al., 1998) using primers Dbf4-up
and Dbf4-down. TetR-Tomato was introduced by transformation with PacI-digested pTetR-
- 13 -

Tom-ADE2. LacI-GFP was introduced by transformation with HindIII-digested p404-LacI-
GFP. The tetO or lacO arrays were introduced by transformation with pARS501-tetO,
pARS1103-tetO, pARS1303-tetO, pARS305-lacO, pARS710-lacO, pARS718-lacO and
pARS1018-lacO digested with PacI, PshAI, BlpI, NotI, PshAI, SnaBI, and BlpI, respectively.

Cell growth and synchronization Cells were grown at 25°C unless otherwise indicated. For
microscopy, cells were grown in complete synthetic medium supplemented with 15 g/mL
adenine (CSM+ade) +2% dextrose, unless otherwise indicated (raffinose or galactose); for
QBU and ChIP-seq, cells were grown in YEP +2% dextrose, unless otherwise indicated
(raffinose or galactose). G1 arrest was achieved by incubation with 2.5nM (1x) –factor
(Sigma T6901); for most extended arrests, a fresh or additional dose of –factor was added
at time of induction/non-induction or at time of temperature shift as indicated in figure
legend. PP1 (Tocris Biosciences) was added to 25 M at the time of initial –factor
incubation.

Live-cell fluorescence microscopy and image analysis ~5x10
6
cells were harvested by
centrifugation and spread on agarose pads made of CSM+ade +4% dextrose. A DeltaVision
wide-field deconvolution microscope was used to capture 28 Z-stacks in 0.25μm increments
for each image. SoftWorX software (Applied Precision/GE Healthcare) was used for
deconvolution and three-dimensional reconstruction of nuclei, and for measuring the
distance between replication origins and nuclear periphery. For experiments with mutant
strains having irregularly shaped nuclei (e.g.: fkh1∆ fkh2-dsm), measurements were made in
- 14 -

three-dimensions; otherwise, measurements were made in two dimensions using a few
middle sections as previously described (Ryu et al., 2015). A z-test was applied to compare
the distribution of measured distances. Images are max intensity projections of 2-4 middle
Z-stacks.

Quantitative BrdU Immunoprecipitation (QBU) QBU and analysis of sequencing reads
was performed as described previously using KAPA Hyper Prep Kit (KK8504) (Haye-
Bertolozzi and Aparicio, 2017). Data analysis was performed using 352 replication origins
classified as Fkh-activated, Fkh-repressed, or Fkh-unregulated (Knott et al., 2012); the latter
two classes are grouped together as “other origins” in Figure S2.

Chromatin Immunoprecipitation analyzed by sequencing (ChIP-seq) ChIP-seq and
analysis of sequencing reads was performed as described previously using KAPA Hyper Prep
Kit (KK8504) (Ostrow et al., 2015). Data analysis was performed using 352 replication
origins classified as Fkh-activated, Fkh-repressed, or Fkh-unregulated (Knott et al., 2012);
the latter two classes are grouped together as “other origins” in Figure S3.

Time-lapse video and MSD analysis A DeltaVision wide-field deconvolution microscope
was used to capture 20 Z-stacks in 0.30μm increments for each time point. GFP signals were
imaged every 12 sec for 5 min, with 0.1 sec exposure for each Z-stack and 32% of transmitted
light using an LED source. All time-lapse movies were deconvolved using SoftWoRx. At least
20 individual cells with nearly stationary nuclei were used to track the trajectory of origin
- 15 -

focus for each strain using Imaris (Bitplane), and MSD curves, Rc, and volumes were derived
as previously described (Caridi et al., 2018); the error bars represent standard error.

Results
Fkh1-induced origin activation re-positions a subtelomeric origin in G1 phase

The association between replication timing and subnuclear localization of replication origins
and the requirement of Fkh1/2 for the clustering of early-firing replication origins according
to chromosome conformation capture studies led us to examine whether Fkh1 has any role
in establishing the spatial positioning of origins within the nucleus. We adapted a system
that we recently engineered that restores origin timing by induction of FKH1 expression in
G1-arrested FKH1/2 mutant cells (Peace et al., 2016). This system has Fkh-activated origin
ARS305 moved into a well characterized, late-replicating, subtelomeric region of
chromosome V-R, replacing the endogenous, late-firing ARS501 (Fig. 1A). In this context, we
showed previously that ARS305
V-R
fails to replicate early in fkh1∆ fkh2∆ cells. However,
induction of FKH1 expression in these cells in G1-phase results in early-firing of ARS305
V-R
in
the ensuing S-phase. In the current study, we used fkh1∆ fkh2-dsm instead of fkh1∆ fkh2∆
cells; fkh1∆ fkh2-dsm cells are essentially null for replication timing control, but exhibit more
normal growth and particularly, more normal cell and nuclear morphologies favorable for
cytological analysis (Ostrow et al., 2017). To locate ARS305
V-R
(or ARS501) in vivo, we
introduced tandem repeats of tetO binding sites adjacent to the origin and expressed TetR-
Tomato protein (Fig. 1A); we also expressed Nup49-GFP (Nup49 is a nuclear pore protein)
to illuminate the nuclear envelope (Belgareh and Doye, 1997).
- 16 -

Figure 1. Fkh1-induced origin activation re-positions a sub-telomeric origin in G1
phase. (A) Schematic of chromosome V-R showing tetO repeats inserted adjacent to the
- 17 -

ARS501 locus, which has been replaced with ARS305 (designated ARS305
V-R
); TetR-Tomato
binds to and illuminates the locus as a single focus. (B) Images of cells from an
unsynchronized culture of strain HYy132 (fkh1∆ fkh2-dsm ARS305
V-R
-Tomato NUP49-GFP
GAL-FKH1) are shown sorted according to cell morphology; all images are at the same
magnification: scale bar = 0.5 m. (C) FKH1 induction scheme: HYy132 cells grown at 25°C
in raffinose medium were arrested in G1 phase with 1x -factor 2.5 hours, incubated an
additional 2 hours in raffinose (Non-induction) or galactose (Fkh1-induction) with 1.7x -
factor, and images of live cells captured, examples of which are shown; scale bar = 0.5 m.
(D) The shortest distance from the ARS305
V-R
-Tomato focus to the nuclear periphery
(Nup49-GFP) in each cell was measured and plotted as quartile boxplots (median shown as
thick black segment) for non-induction and FKH1-induction; the result of a z-test
comparing the two distributions is given as P. (E) Cells of fkh1∆ fkh2-dsm GAL-FKH1
NUP49-GFP strains HYy119 (ars305-∆ACS
V-R
-Tomato) and HYy120 (ars305-∆2BS
V-R
-
Tomato) were treated and analyzed as above.

Microscopic examination of cells showed a single Tomato focus per undivided nucleus (Fig.
1B). Images of cells from an unsynchronized population were sorted according to budding
morphology, which is reflective of cell cycle progression. The localization of the ARS305
V-R
-
Tomato focus correlates with cell cycle stage, showing primarily peripheral localization in
unbudded and small-budded cells and interior localization in larger-budded cells (Fig. 1B).
This is consistent with previous studies showing peripheral localization of
subtelomeric/late-firing origins in G1 followed by relocalization to the interior during S
- 18 -

phase (Heun et al., 2001a). Because origin timing is normally established in G1, we focused
further analysis on origins in G1 phase cells.

In G1-arrested fkh1∆ fkh2-dsm cells, almost all cells exhibited peripheral localization of
ARS305
V-R
(Fig. 1C, left panel). Induction of FKH1, however, resulted in an increase in the
proportion of cells with non-peripheral positioning of ARS305
V-R
(Fig. 1C, right panel),
suggesting that origin relocalization is associated with initiation timing re-programming by
FKH1. We confirmed that re-localization is a direct result of FKH1 induction by
demonstrating that neither the induction scheme (raffinose -> galactose) with a strain
lacking inducible FKH1 nor a non-inducing change to a more favorable carbon source
(raffinose -> dextrose) resulted in origin re-localization (Figure 1-figure supplement 1). To
confirm the change in origin localization resulting from FKH1 induction, we created three-
dimensional image reconstructions from confocal z-stacks and measured the shortest
distance in three dimensions from the ARS305
V-R
focus to Nup49-GFP signal in the nuclear
envelope amongst populations of cells. Statistical analysis of these measurements shows a
significant increase in the distances associated with FKH1-induction versus non-induction
(Fig. 1D).

We tested whether the function of ARS305 is required for relocalization by introducing into
the V-R locus ARS305 bearing a mutation of the ARS consensus sequence (ACS) (ars305-
∆ACS), which is essential for ORC binding and origin function. Disruption of ARS305
function not only eliminated its relocalization in response to FKH1 induction but also
- 19 -

resulted in an even more peripheral distribution, suggesting that a functional origin is
required for relocalization away from the periphery (Fig. 1E). We also tested ARS305 with
mutations of two proximal Fkh1/2 binding sites (ars305-∆2BS), which retains origin
function but is delayed in activation at its normal locus (Knott et al., 2012); ars305-∆2BS
V-R

did not relocalize upon FKH1 induction, confirming that Fkh1 acts through direct binding in
cis to ARS305
V-R
(Fig. 1E). Using chromatin immunoprecipitation, we verified that Fkh1
binding was eliminated specifically at ars305-∆2BS
V-R
(Fig. A1). Mutation of the ACS also
resulted in loss of Fkh1 binding specifically at ars305-∆ACS
V-R
(Fig. A1), consistent with a
previous report that origin licensing is required for Fkh1-origin binding (Reinapae et al.,
2017), leaving it unclear whether an additional function in origin firing downstream of
Fkh1/2 binding is required for re-localization.

In the experiments above, relocalization of ARS305
V-R
involved induction of FKH1 from the
GAL1/10 promoter, which results in higher than normal levels of Fkh1 protein (Peace et al.,
2016). To determine whether this overabundance of Fkh1 was required for the origin
relocalization, we compared localization of ARS305
V-R
with ARS501 in cells with native FKH1
(and FKH2) expression (Fig. 2A). The analysis showed that ARS305
V-R
was significantly more
distant from the nuclear periphery than ARS501 (Fig. 2B). We also analyzed origin timing of
ARS305
V-R
in FKH1 (fkh2-dsm) versus fkh1∆ (fkh2-dsm) cells by quantitative BrdU
immunoprecipitation (QBU) of cells released from G1 phase into hydroxyurea (HU), in which
early but not late origins fire efficiently. We found that ARS305
V-R
fired efficiently in HU in
FKH1 but not fkh1∆ cells (Fig. 2C and Figure 2-figure supplement 1). Thus, normal Fkh1
- 20 -

levels are able to overcome the effect that subtelomeric location has on subnuclear
localization and initiation timing of ARS305
V-R
.

- 21 -

Figure 2. Normal dosage of Fkh1 is sufficient to relocalize ARS305
V-R
and advance its
firing time. (A) HYy160 (ARS501-Tomato NUP49-GFP) and HYy157 (ARS305
V-R
-Tomato
NUP49-GFP) cells were arrested in G1 phase at 25°C with 1x -factor 2 hours and images
were collected; scale bar = 0.5 m. (B) Distances from origin foci to nuclear periphery were
determined, plotted as quartile boxplots, and analyzed by a z-test. (C) Quantitative BrdU-
IP-Seq (QBU) analysis was performed with ARS305
V-R
-bearing strains HYy113 (fkh2-dsm)
and HYy38 (fkh1∆ fkh2-dsm) after G1 block-and-release into hydroxyurea in the presence
of BrdU; averaged data from three experimental replicates was plotted for the V-R region
with the positions of several replication origins indicated; ARS305
V-R
resides at the ARS522
(aka: ARS501) locus.

Fkh1 globally regulates subnuclear positioning of early origins in G1 phase

We tested whether the Fkh1-dependent localization of ARS305
V-R
is also responsible for
ARS305 localization when residing at its native locus more distal from the telomere. We
inserted a lacO array near ARS305 and expressed LacI-GFP and Nup49-GFP; imaging
showed that the LacI-GFP focus is clearly distinguishable from the more diffuse Nup49-GFP
signal (Fig. 3A). Consistent with previous analysis, ARS305 was non-peripheral in most G1-
arrested WT cells (Fig. 3A) (Heun et al., 2001a), however, deletion of FKH1 significantly
increased the proportion of cells in which ARS305 was closer to the periphery (Fig. 3A).
Consistent with this requirement for FKH1, elimination of the Fkh1/2 binding sites in
ARS305 also resulted in peripheral localization (Figure 3-figure supplement 1A). Moreover,
- 22 -

ars305-∆ACS, which lacks origin function (and Fkh1/2 binding) exhibits an even more
peripheral distribution (Figure
3-figure supplement 1A), suggesting that origin function is required for interior localization
in G1 and that Fkh1/2 stimulates this localization. Previous analysis showed that ARS305
initiation timing was significantly delayed in the absence of FKH1 or Fkh1/2 binding sites
(Knott et al., 2012), so once again we observe a FKH1-dependent relationship between
subnuclear localization in G1 phase with replication initiation timing in S phase.

- 23 -

Figure 3. Fkh1 determines early origin positioning globally. Diagrams of
chromosomes with replication origins labeled with lacO/LacI-GFP (green-filled segment)
or tetO/TetR-Tomato (red-filled segment) are shown above the corresponding images.
Distances between origins (black-filled spheres) and telomeres, and in some cases
centromeres (ovals), are indicated and include ~14kb or 16kb added by lacO or tetO
repeats, respectively; elements are not drawn to scale. Cells of WT and fkh1∆ strains with
ARS305-GFP (HYy151, HYy147) in (A), ARS1303-Tomato (HYy166, HYy173) and ARS1103-
Tomato (HYy165, HYy172) in (B), ARS710-GFP (MPy6, MPy10), ARS718-GFP (MPy20,
MPy21), and ARS1018-GFP (MPy19, MPy22) in (C), all expressing NUP49-GFP, were
arrested in G1 phase at 25°C with 1x -factor 2 hours and live images were captured; scale
bar = 0.5 m. Distances from origin foci to nuclear periphery were determined, plotted as
quartile boxplots, and analyzed by a z-test.

To determine whether other Fkh-activated origins’ localizations are also determined by
FKH1, we performed similar tests by inserting a tetO array adjacent to a few additional
Fkh-activated origins, expressing TetR-Tomato and Nup49-GFP, and deleting FKH1. Like
ARS305, ARS1303 and ARS1103 were located closer to the periphery in G1-arrested fkh1∆
versus WT cells (Fig. 3B). ARS305, ARS1303 and ARS1103 are relatively telomere proximal,
residing 39, 32, and 56kb from the nearest telomere respectively (the lacO or tetO arrays
add ~14 or 16kb, respectively to these distances), which might constrain the extent to
which FKH1 influences their positioning. To address this possibility, we tested localization
of several additional Fkh-activated origins that are more distal from telomeres, including:
ARS710, ARS718, and ARS1018, residing at 204, 421, and 205kb from the nearest telomere.
- 24 -

All of these origins show significant reduction in distance from the nuclear periphery upon
deletion of FKH1 (Fig. 3C). We also observed similar results with ARS305 and ARS710 in G1
cells from an unsynchronized population (Figure 3-figure supplement 1B). In contrast, the
peripheral localization in G1 of late origin ARS501, which is not Fkh-activated, was not
altered by deletion of FKH1 (Figure 3-figure supplement 1C). These results suggest that
FKH1 plays an expansive role in relocalizing replication origins from the nuclear periphery
to the nuclear interior in G1 phase.

DDK- but not CDK-dependent step of replication initiation drives origin
relocalization

Because our previous study indicated that Fkh1/2 was required for origin recruitment of
Cdc45 in G1 phase (Knott et al., 2012), we tested the requirement for CDC7, which encodes
the catalytic subunit of DBF4-dependent kinase (DDK), and is required for Cdc45 origin-
loading (reviewed in (Tanaka and Araki, 2013). We introduced the cdc7-as3 allele, the
kinase activity of which is inhibited by ATP analog PP1 (Wan et al., 2006), and tested
whether FKH1-induced origin relocalization occurs with inhibition of CDC7 function.
Remarkably, ARS305
V-R
relocalization was eliminated by inhibition of Cdc7-as3 kinase with
PP1 (Fig. 4A, also compare with non-induction in Fig. 1D). These results suggest that DDK
activity is required for origin relocalization in G1-arrested cells.
To test whether the same result of the endogenous ARS305 can be observed in a wild type
Fkh1 background, we performed the re-positioning assay on cdc7-as3 single mutant cells in
G1 phase. However, ARS305 maintains its interior location in the nucleus in G1 with
inhibited DDK kinase function by PP1 (data not shown). Then, we compared the conditions
- 25 -

of cell cycle progression between fkh1Δ fkh2dsm and WT cells with inhibited DDK function
by using cdc7-as3 allele. We synchronized cell with -factor for 3.5h plus PP1 and then
released cells in S phase while maintaining PP1 treatment. Based on the FACS results, we
discovered a significant delay S phase entry in cdc7-as3 fkh1Δ fkh2-dsm cells compared to
cdc7-as3 single mutant cells (Fig. A2). While fkh1Δ fkh2dsm cells did not show differences
on cell cycle progression with WT cells when DDK’s function is normal. This near 40min-
delay indicates cdc7as3 mutant is very sensitive to Fkh1 background with PP1. The residual
of Fkh1 function plays a positive role in maintaining ARS305 at the center of the nucleus.
Additionally, cdc7as3 allele, as a relative less stringent mutant, is not sufficient enough to
affect the subnuclear location of ARS305 by inhibiting DDK function in wild type Fkh1
background.
- 26 -

Figure 4. Origin localization in G1 is DDK regulated. (A) HYy186 (fkh1∆ fkh2-dsm GAL-
FKH1 ARS305
V-R
-Tomato NUP49-GFP cdc7-as3) cells were subjected to FKH1-induction
scheme as described in Figure 1C legend except that PP1 or DMSO (vehicle) was included
with -factor, and images were captured. (B) Cells of ARS305-GFP NUP49-GFP strains
HYy151 (WT) and HYy191 (cdc7-4) were arrested in G1 with 1x -factor 2 hours at 25°C
followed by 1 hour incubation at 37°C with 2x -factor, and images were captured. (C)
HYy181 (ARS305-GFP NUP49-GFP dbf4∆C) cells were arrested in G1 phase with 1x -factor
2 hours at 25°C and live images were captured. The control experiment with WT cells
- 27 -

(HYy151) is shown in Figure 3A. (A, B, C) Scale bar = 0.5 m. Distances from origin foci to
nuclear periphery were determined, plotted as quartile boxplots, and analyzed by a z-test.

A role for DDK in G1 phase was unexpected as DDK activity has been reported to be low in
-factor-arrested G1 cells due to instability of Dbf4 (Nougarede et al., 2000; Oshiro et al.,
1999). To provide further evidence for DDK’s role, we tested a native origin without FKH1
overexpression, and to inactivate CDC7, we chose the temperature-sensitive cdc7-4 allele
(Hereford and Hartwell, 1974). For this experiment, G1-arrested WT and cdc7-4 cells
bearing a lacO array inserted near ARS305 and expressing LacI-GFP were shifted to the
non-permissive temperature and ARS305 location was determined. Compared to WT cells,
cdc7-4 cells at the non-permissive temperature showed a significant increase in the
proportion of cells with ARS305 near the nuclear periphery (Fig. 4C). This result supports
the conclusion that DDK activity is required for origin re-positioning in G1 phase cells.

Fkh1-origin binding is cell cycle-regulated, occurring in G1 and S phases (Ostrow et al.,
2014), suggesting that the requirement for DDK activity in Fkh1-stimulated origin
relocalization might be due to dependence of Fkh1 origin-binding on DDK. We tested this
possibility by performing chromatin immunoprecipitation analysis of Fkh1 comparing WT
and cdc7-as3 cells. The results showed that binding of Fkh1 to ARS305 and other Fkh-
activated origins was largely unaffected by Cdc7 inhibition (Figure 4-figure supplement 1).
Thus, Fkh1 origin binding appears to be independent of DDK activity, and, by inference, of
the subcellular change in localization resulting from DDK inhibition. Alternatively, the
requirement for DDK activity in Fkh1-stimulated origin relocalization may reflect Fkh1
- 28 -

acting upstream of DDK, which would comport with a recent report that a critical role of
Fkh1 in origin stimulation is DDK recruitment through direct physical interaction with
Dbf4 (Fang et al., 2017). We tested whether the same mechanism is responsible for Fkh1-
induced origin re-positioning by testing the effect on ARS305 positioning in cells expressing
Dbf4 lacking its C-terminus (dbf4∆C), which is required for interaction with Fkh1 (Fang et
al., 2017). Deletion of DBF4’s C-terminus had a similar effect on origin localization as FKH1
deletion, with greater enrichment of ARS305 near the nuclear periphery (Fig. 4B. compare
with Fig. 3A), consistent with Fkh1 and Dbf4 acting in the same pathway.

The essential function of DDK in origin firing is phosphorylation of MCM helicase subunits,
particularly Mcm4, resulting in removal of auto-inhibition and enabling recruitment of
helicase accessory protein Cdc45 through its loading factor Sld3 (reviewed in (Tanaka and
Araki, 2013). To test whether the requirement for Cdc7 kinase activity in origin
relocalization reflects its function in Mcm4 helicase phosphorylation, we introduced into
the cdc7-as3 strain an allele of MCM4, MCM4-DD/E(7)+DSP/Q(7) abbreviated herein as
MCM4-14D, which contains 14 S/T->D substitutions that mimic critical DDK-
phosphorylated residues in Mcm4, and suppresses reduced Cdc7 kinase activity (Randell et
al., 2010). The presence of MCM4-14D restores ARS305
V-R
relocalization upon FKH1
induction in the cdc7-as3 strain inhibited by PP1 (Fig. 5A). This supports the conclusion
that the function of Cdc7 kinase required for origin relocalization is phosphorylation of
Mcm4.
- 29 -

Figure 5. DDK regulation of origin localization reflects its phosphorylation of Mcm4
and consequent Cdc45 loading. (A) Fkh1-induction scheme with PP1 as described in
- 30 -

Figure 4A legend was carried out with fkh1∆ fkh2-dsm GAL-FKH1 ARS305
V-R
-Tomato
NUP49-GFP strains HYy186 (cdc7-as3) and HYy177 (cdc7-as3 MCM4-14D), and images
captured. (B) ARS305-GFP NUP49-GFP strains HYy151 (WT) and HYy184 (cdc45-1) were
arrested in G1 with 1x -factor 1 hour at 30°C followed by 2 hour incubation at 16°C with
1x -factor, and images were captured. (C) Cells of strain HYy177 harboring no plasmid or
high-copy plasmid expressing CDC45 were arrested in G1 with 1x -factor 2 hours at 25°C
and images captured. (D) ARS305-GFP NUP49-GFP strain HYy197 (cdc28-as1) cells were
arrested in G1 phase with 0.5x -factor 2 hours at 25°C, PP1 or DMSO was added and
incubated one additional hour with 0.5x -factor, and images were captured. (A-D) Scale
bar = 0.5 m. Distances from origin foci to nuclear periphery were determined, plotted as
quartile boxplots, and analyzed by a z-test.

We tested whether completion of the DDK-dependent step, that is Sld3 and Cdc45 loading,
is required for origin relocalization by testing the effect of inactivation of CDC45 function.
The cold-sensitive cdc45-1 allele exhibits interdependence with heat-sensitive alleles cdc7-
4 and dbf4-1 in reciprocal temperature-shift experiments, tightly inhibits replication
initiation, and reduces Sld3-origin association in G1 phase (Aparicio et al., 1999; Kamimura
et al., 2001; Owens et al., 1997). We synchronized WT and cdc45-1 cells in G1 phase at the
permissive temperature and shifted the cultures to semi-permissive temperature while
maintaining the G1 arrest. Analysis showed that ARS305 was more peripherally localized
in cdc45-1 cells at the semi-permissive temperature in G1 phase (Fig. 5B). As origin
binding of Cdc45 and Sld3 are interdependent and Cdc45-1 inactivation reduces Sld3-
- 31 -

origin binding (Kamimura et al., 2001), these results suggest that assembly of Sld3-Cdc45
onto origins is required for origin relocalization.

Cdc45 is incorporated into replisomes as a component of the active helicase complex
together with MCM and GINS. However, Cdc45 is present in low abundance and is likely
limiting for the total number of active replisomes that may be simultaneously active
(Mantiero et al., 2011; Tanaka et al., 2011). We noticed that the presence of Mcm4-14D
was not sufficient to relocalize ARS305
V-R
in fkh1∆ fkh2-dsm cells in the absence of Fkh1
induction (Fig. 5C), which might be contrary to expectations if the only function of Fkh1 is
to physically recruit DDK, which has been rendered dispensable by Mcm4-14D. We note,
however, that MCM4-14D does not suppress a deletion of CDC7 (S.P. Bell, personal
communication) suggesting that residual DDK activity is required for sufficient origin
firing, and hence, Fkh1 may act to target this residual activity to specific origins. Absent
this targeting, we postulated that the limited abundance of Cdc45 would be further diluted
amongst all licensed origins due to potentiation by Mcm4-14D. To test this idea, we
introduced a high-copy plasmid expressing Cdc45 from its native promoter into the fkh1∆
fkh2-dsm MCM4-14D strain, and examined origin location. Consistent with the notion that
Cdc45 is limiting for execution of the DDK-dependent step, expression of high copy Cdc45
significantly increased the frequency of ARS305 more distal from the periphery (Fig. 5C).
This finding supports the conclusion that full execution of the DDK-dependent step in the
form of Cdc45 loading, as opposed to Mcm4 phosphorylation itself or phosphorylation of
other targets is required for origin relocalization.
- 32 -

As the interior localization of early origins occurs in -factor-arrested, G1 phase cells,
cyclin-dependent kinase (CDK) activity would appear to be dispensable because G1 phase
cells have very low levels of S/G2/M-CDK activities, and G1-CDKs, which are required for
passage through Start, are inhibited by -factor (reviewed in (Mendenhall and Hodge,
1998). Nevertheless, low levels of S/G2/M-CDK activities in G1 phase cannot be ruled out,
and indeed, it appears that low levels of DDK are involved. Thus, to address the possibility
that CDK activity might be contributing to G1 phase origin dynamics, we tested the
requirement for CDC28, the Cdk1 kinase, using analog-sensitive cdc28-as1 cells (Bishop et
al., 2000). In G1-arrested cells, inhibition of Cdc28-as1 activity with PP1 did not alter
localization of ARS305 (Fig. 5D), although budding was inhibited upon release from -
factor arrest indicating effective inhibition of Cdc28-as1 (Figure 5-figure supplement 1A).
Similarly, PP1 treatment of cycling cdc28-as1 cells did not alter distribution of ARS305 in
G1 phase cells (Figure 5-figure supplement 1B), while DNA content analysis showed
delayed entry of cells into S phase indicating effective inhibition of Cdc28-as1 (Figure 5-
figure supplement 1C). Thus, CDK activity appears to be dispensable for normal, Fkh1-
dependent positioning of ARS305. Overall, our findings indicate that DDK but not CDK
activity stimulates origin relocalization in G1 phase.

Origin mobility increases with origin relocalization

Fkh1 might facilitate origin relocalization by promoting origin mobilization (release from
the periphery or movement per se), or by increasing the stability of origin-origin
- 33 -

interaction after relocalization. In addition to changes in location, replication origins
exhibit decreased rate of mobility during progression into S phase (Heun et al., 2001b). We
directly investigated how Fkh1 affects origin mobility by tracking the locations of ARS305
and ARS718 in individual WT and fkh1∆ cells over time, and applying mean square
displacement (MSD) analyses (Marshall et al., 1997). The analysis shows significantly
lower plateau of MSD curves in fkh1∆ cells (Fig. 6A), consistent with less nuclear space
explored. Calculation of the radius of constraint (Rc) and the corresponding volume of
space explored reveals that ARS305 explores about 2.5-fold more volume and ARS718
explores about 3.8-fold more volume in WT than fkh1∆ cells. Tracings of the paths of origin
foci show confinement proximal to the nuclear periphery in fkh1∆ cells (Fig. 6B). Together,
these data show that Fkh1 stimulates origin mobilization.
- 34 -

Figure 6. Origin mobility increases with origin relocalization. (A) Mean-squared-
displacement (MSD) analysis of tracking data for ARS305-GFP strains HYy151 (WT) and
HYy147 (fkh1∆) and ARS718-GFP strains MPy20 (WT) and MPy21 (fkh1∆). Radius of
constraint (Rc) and volume searched (V) are given, and statistical significance comparing
WT and fkh1∆ was estimated by two-tailed Mann-Whitney test. (B) Images (left) and 3D
reconstructions with Imaris (right) showing examples of tracks of origin focus over time
(color corresponding to time progression); scale bar = 0.4 m.
- 35 -

Discussion
This study reveals new links between key molecular interactions in replication initiation
and the localization and mobility of replication origins within the nucleus. In particular, we
show that early origin specification in G1 phase by Fkh1 induces a change from peripheral
to interior nuclear localization of Fkh1-activated origins. Quite remarkably, we find that
origin relocalization requires execution of the DDK-dependent step of origin firing that
loads Cdc45. That the DDK requirement reflects the key, recognized function of DDK in
replication initiation, that is, phosphorylation of MCM proteins leading to Sld3-Cdc45
origin-loading, is demonstrated by the bypass of CDC7 requirement by phosphomimetic
mutations in MCM4-14D as well as the dependence on CDC45 function. This early
execution of the DDK step was unexpected because DDK levels have been reported to be
very low in a-factor-arrested G1 cells due to Dbf4 instability (Nougare`de et al., 2000;
Oshiro et al., 1999). Our findings provide direct evidence that DDK is active in G1 phase and
has already established origin timing by late G1 phase in a-factor arrest. This finding
explains previous observations that Sld3 and Cdc45 associate with early replication origins
in G1 phase (Aparicio et al., 1999; Kamimura et al., 2001). Our findings are also consistent
with a more recent study showing that Sld3- and Cdc45-origin association in G1 phase is
DDK-dependent and CDK-independent, as well as the conclusion that DDK acts prior to and
independently of S-CDK (Heller et al., 2011; Yeeles et al., 2015), the latter of which is
dispensable for the observed origin relocalization.

- 36 -

The finding that Fkh1 and DDK are required for origin relocalization fits well with the
recent finding that Fkh1 acts to stimulate origin firing by directly recruiting Dbf4 through
physical interaction (Fang et al., 2017), and extends our understanding of the significance
of this interaction to replication initiation via nuclear positioning of replication sites. As
predicted by this interaction model, inactivation of DDK activity should phenocopy deletion
of FKH1, as we have herein demonstrated with depletion of CDC7 function. Moreover,
specific deletion of Dbf4’s C-terminus, which is required for interaction with Fkh1, also
phenocopies deletion of FKH1. Furthermore, the absence of FKH1 and FKH2 function is
bypassed by the MCM4-14D allele in the presence of increased levels of Cdc45. Together,
these results support a mechanism involving Fkh1 recruitment of DDK activity to load
Cdc45 at a subset of origins in G1, corresponding with a change in subnuclear positioning
of these origins, and early firing in the subsequent S phase.

Chromosome conformation capture (Hi-C) experiments have indicated that early firing
replication origins preferentially interact with each other, or ‘cluster’ in G1 phase (Duan et
al., 2010). Related studies have shown that Fkh1/2 is required for these spatial interactions
amongst early origins (Eser et al., 2017; Knott et al., 2012; Ostrow et al., 2017). We propose
that the origin clustering interactions revealed by Hi-C experiments directly reflect origin
localization to distinct nuclear territories as observed microscopically. Thus, localization to
the nuclear interior might increase the likelihood for physical interaction amongst this
subset of origins. Such interactions may be driven by cooperative interactions between
Fkh1-bound origins recruiting limiting initiation factors such as Dbf4, Sld3 and Cdc45. This
aggregation of origins selected for early/efficient activation has the inevitable consequence
- 37 -

that replication initiation transforms these origin clusters into replication foci, which have
been observed as concentrations of DNA synthesis and replication factors (Berezney et al.,
2000; Frouin et al., 2003; Hoza´k et al., 1994; Kitamura et al., 2006; Nakamura et al., 1986;
Newport and Yan, 1996). These assemblages may contribute to efficient chromosomal
replication initiation and elongation in multiple ways, such as accretion of activities and co-
factors directly required for DNA synthesis (e.g.: dNTP production), and scaffolding to
colocalize and coordinate replication with related activities like chromatin assembly,
cohesion establishment, topological resolution, and DNA repair.

It remains to be determined exactly what maintains either the peripheral or interior
localization of origins or what drives relocalization between different subnuclear zones.
While telomere tethering to the nuclear envelope has been assumed to cause the peripheral
localization of telomere-proximal origins, we find that early origins distal from telomeres
that are normally enriched in the nuclear interior, are closer to the nuclear periphery in
cells lacking Fkh1, suggesting that perinuclear localization represents a default state for
most origins irrespective of telomere tethering (Figure 7A). It is unclear what promotes
this origin localization. Complete elimination of origin function results in even more
peripheral distribution of the locus suggesting that peripheral localization is independent
of an origin tethering mechanism, and that interior localization is linked to activation of
origin function, which may occur, with less efficiency, in the absence of Fkh1/2. There may
be passive exclusion from the interior where other activities like transcription may
predominate in early G1, or there may be a dedicated tethering mechanism, though origin
- 38 -

association with the periphery does not appear to be as stringently localized or as stable as
that of telomeres (Hediger et al., 2002; Heun et al., 2001a; Heun et al., 2001b).

Figure 7. Model of origin localization linked to initiation. (A) Absent Fkh1, most
replication origins are enriched at nuclear periphery, however, Fkh1 binding to a subset of
origins allows execution of the DDK-dependent step of initiation, resulting in release from
the nuclear periphery and/or capture in the nuclear interior to form early origin clusters.
(B) Hypothetical mechanism for origin tethering to the nuclear periphery regulated by
Rif1-PP1 versus Fkh1-DDK-Cdc45 activities. Rif1 associates with inner nuclear membrane
and with licensed replication origins, while associated PP1 antagonizes execution of the
- 39 -

DDK-dependent step. Fkh1-dependent recruitment of DDK results in phosphorylation of
MCM, Cdc45 loading and local release from Rif1 and PP1. See text for further discussion.

In addition to subtelomeric origins, Rif1 regulates and associates with origins distal from
telomeres and with the nuclear envelope, and therefore could potentially tether origins to
the periphery (Figure 7A) (Hafner et al., 2018; Park et al., 2011; Peace et al., 2014). Rif1
interacts with Dbf4 and with the counteracting PP1 phosphatase, suggesting that the Rif1-
origin interaction may be down-regulated by DDK-dependent phosphorylation of MCM
proteins and/or Rif1 (Dave´ et al., 2014; Hiraga et al., 2014; Mattarocci et al., 2014). Thus,
Fkh1-mediated, origin-specific recruitment of DDK may overwhelm Rif1-mediated PP1
inhibition locally and thereby release the origin from peripheral tethering (Figure 7B).
Consequent Cdc45 loading might effectively prevent reversal of MCM phosphorylation and
fully disrupt interaction with Rif1-PP1. Alternatively, MCM phosphorylation and/or Cdc45
loading might change the licensed origin’s biophysical properties, thereby forcing the
origin to occupy and search different space and/or capture scaffolding factors, which may
themselves be localized to the interior and thus stabilize interior localization. Future
studies aimed at more detailed examination of how individual factors affect origin mobility
should provide further insights.
Previous studies have concluded that peripheral localization of origins is neither necessary
nor sufficient to regulate initiation timing. In one study, subtelomeric origin ARS501
remained late firing following excision (in a-factor-arrested G1 cells) from the
chromosome, which allowed its diffusion away from the nuclear periphery, leading the
authors to suggest that peripheral localization might promote a chromatin mark that
- 40 -

maintains late timing (Heun et al., 2001a). However, we have shown that induction of Fkh1
(in a-factor-arrested cells) can reprogram timing of a Fkh-activated origin inserted into the
ARS501 locus (Peace et al., 2016). We propose that the relevant chromatin mark is MCM
phosphorylation, removal of which is promoted by peripheral localization and addition by
DDK recruitment. Thus, excised ARS501 remains late despite its mobilization because DDK
is limiting and already recruited by Ctf19 and Fkh1/2 to other origins. Other studies have
shown that tethering of early origins to the nuclear periphery does not delay their
activation (Ebrahimi et al., 2010; Zappulla et al., 2002). However, both origins in these
previous studies, ARS305 and ARS607, are Fkh-activated origins that we have shown can
overcome the replication initiation delay associated with peripheral localization. Overall,
these previous findings fit neatly into our model, which suggests that interior origin
localization is a consequence rather than a cause of early timing.

- 41 -

Chapter II
Fkh1 domain-swapping motif regulates subnuclear localization
of replication origin suggesting a role of recruitment of Dbf4

Introduction
Fkh1 and Fkh2 binding near origins stimulate the interactions among origins, which
presumably occurs through binding to origin recognition complexes (ORCs) at origins
(Knott et al., 2012). Our lab proposed that origin clustering facilitates the interaction
among origins to recruit the limiting initiation factors, which eventually results in early
firing (Aparicio OM, 2013). With a similar mechanism, Fkh1 is also able to regulate mating-
type switching. Mating-type switching is a homologous recombination process between
distal chromosomal loci which involves long-term interactions among Fkh proteins (Haber
JE, 2012). However, how exactly they promote clustering is not clear. We have recently
found that the domain-swapping motif in Fkh1 and Fkh2 is crucial for replication timing
regulation (Ostrow et al., 2017). Mutations at specific residues were introduced to create
“domain-swap minus” (dsm) alleles of FKH1. The results showed that the dimerization
ability was abolished in fkh1-dsm mutant cells and caused a significant delay in replication
timing especially among Fkh-activated origins. Therefore, the data further supports the
mechanism that Forkhead proteins facilitate the establishment of physical communications
between origins for clustering in advancing replication.

- 42 -

The winged-helix DNA binding domain of the Forkhead Box (FOX) family of transcription
factors is conserved in eukaryotes from yeast to humans (Lalmansingh et al., 2012). A
conserved proline is replaced with alanine in the region of the winged-helix DNA binding
domain in FoxP. This insertion of alanine at this position permits domain swapping or
helical extension, which is usually halted by the rigid backbone of proline (Lalmansingh et
al., 2012; Stroud et al., 2006; Bandukwala et al., 2011). This shared feature, conserved
alanine, among FOXP family members correlates their ability to form domain-swapped
dimers with functional importance (Stroud et al., 2006; Bandukwala et al., 2011). It has
been shown that Fkh1 domain-swapping motif has a strong regulatory ability on
replication timing. However, whether it is also important for origin subnuclear localization
regulation has not previously been tested.

A recent study found that there are direct interactions between Fkh1 and Dbf4, the
regulatory subunit of DDK (Fang et al., 2017). This data suggest that DDK is controlled by
Fkh1/2 to a subset of replication origins to promote early initiation. Given that DDK-
dependent activation of replication origin is a universal step in the genome, this additional
piece of evidence unveils that the dynamic mechanism of replication timing regulation by
Fkh1/2 to licensed origins is because of the competition of limiting Dbf4. Therefore, Fkh
proteins play an important role in regulating replication timing globally through
interacting with DDK. In Schizosaccharomyces pombe, DDK recruitment is regulated by the
interaction between Swi6 and Dfp1 (the fission yeast Dbf4 orthologue) (Hayashi et al.,
2009). Interestingly, subtelomeres remain late firing despite having Swi6 bound. Once DDK
was artificially recruited to the subtelomeric region, an advanced replication timing was
- 43 -

observed. However, whether the low activities and concentration of DDK or DDK
counteracting activity of Rif1-PP1 at subtelomeres are responsible for late firing of
subtelomeric origins is still obscure.

Early studies on replication timing analysis showed that centromeres present early
initiation and usually stay near other early-replicating factories in the nucleus (Pohl et al.,
2012; Knott et al., 2012). Later studies in both Schizosaccharomyces pombe and S.
cerevisiae showed that the early firing of the CEN region is because of the DDK-recruitment
to centromeric origins (Hayashi et al.., 2009; Natsume et al., 2013). In particular, Ctf19, a
kinetochore complex, can recruit the Scc2–Scc4 complex to the centromere (Eckert et al.,
2007; Fernius and Marston et al., 2009; Ng et al., 2009). Specifically, a new function of Ctf19
in facilitating the Dbf4-dependent kinase (DDK) accumulation at kinetochores in telophase
of G1 phase was discovered by the Tanaka group in 2013. The deletion of CTF19 showed a
considerably reduced Dbf4 association with centromeres. Additionally, a reduction of the
initial formation of replication factories at centromeric regions was observed too.
Therefore, the Ctf19 complex promotes DDK association with centromeres and regulates
DNA replication in this region. Considering the robust ability of recruiting Dbf4 for both
Fkh and Ctf19 to Fkh-act origins and CEN-proximal origins respectively, a competing
relationship between these two recruiters of accessing limiting Dbf4 is proposed. In
Chapter 2, we also found that Fkh1 domain-swapping domain is required for Dbf4
recruitment.

- 44 -

Materials and Methods
Plasmid constructions Plasmids were constructed using Gibson Assembly kit (SGI
cat#GA1200). Restriction enzymes were from New England Biolabs and used according to
their protocols. Sequence was confirmed by DNA sequencing (Retrogen Inc.). Primer
sequences for plasmid constructions are given in Table 1. Fkh1-Myc and fkh1-dsm-Myc was
PCR-amplified from p403-Fkh1-Myc and p403-fkh1-dsm-Myc respectively (from Ostrow et
al., 2017) using primers Fkh-Myc-F and Fkh-Myc-R and subcloned into EcoRI+MscI-digested
vectors p306-ΔNdbf4-FKH to yield p306-ΔNdbf4-FKH-Myc and p306-ΔNdbf4-fkh-dsm-Myc
respectively.

Yeast strain constructions All strains are congenic with SSy161, derived from W303-1a
(RAD5) (Viggiani and Aparicio, 2006); complete genotypes are given in Table 2. Strain
constructions were carried out by genetic crosses or lithium acetate transformations with
linearized plasmids or PCR products generated with hybrid oligonucleotide primers having
homology to target loci (Ito et al., 2001; Longtine et al., 1998); primer sequences for strain
constructions are given in Table 1. Genomic alterations were confirmed by PCR analysis or
DNA sequence analysis as appropriate.

HYy222 was generated by crossing HYy180 with OAy1108. HYy192 was made from HYy147
followed by selection on 5-FOA. HYy210 was made by crossing HYy143 with JPy19 and
CTF19 was deleted from the diploid strain by using primers Ctf19-F and Ctf19-R to amplify
- 45 -

TRP1 selectable marker from pFA6-TRP1 (Longtine et al., 1998). Dbf4-Fkh and Dbf4-fkh-dsm
were introduced by transformation with EcoNI-digested p306-ΔNdbf4-FKH484-MYC and
p306-ΔNdbf4-fkh484-dsm-MYC respectively.

Results
Domain-swapping motif in Fkh1 is required to relocalize ARS305 to the interior of
the nucleus in G1

Based on the previous discoveries from our group, domain-swapping motif in Fkh1 is
essential for replication timing regulation (Ostrow et al., 2017). In particular, the
interaction between Fkh proteins was affected and further showed significant late firing in
Fkh1 domain swap mutant (fkh1-dsm) cells. To determine whether the domain-swapping
motif in Fkh1 was also required for the origin relocalization in G1, the origin re-positioning
assay was conducted and compared the localization of ARS305 in WT and fkh1-dsm in G1
phase arrested cells. The result revealed that ARS305 lost its interior nuclear localization
but moved to the nuclear periphery, which indicates that the domain-swapping motif of
Fkh1 is required for the origin subnuclear relocalization in G1 (Fig 2.1). This result support
the mechanism that early origin clustering in G1 facilitated by Fkh1 stimulates early
initiation in S phase. Together, the Fkh1 domain-swapping motif is critical for regulating
subnuclear localization.
- 46 -

Figure 2.1. Domain swap mutant of Fkh1 fails to relocalize ARS305 to the center of
the nucleus in G1. (A) ARS305-GFP NUP49-GFP strains HYy147 (fkh1Δ) and HYy222
(fkh1-dsm) were arrested in G1 with 1x -factor 2 hours at 25°C and images captured.
Distances from origin foci to nuclear periphery were determined, plotted as quartile
boxplots, and analyzed by a z-test.

Fkh1 and DBF4 act in the same pathway to regulate replication timing temporally
and spatially

It has been shown that Fkh1 plays an upstreaming role by recruiting Dbf4 to early
replication origins (Fkh-act origins) to advance replication initiation (Fang et al., 2017). In
addition, the interaction between Fkh1 and Dbf4 is also critical for origin subnuclear
localization in G1 phase (Zhang et al., 2019). To be specific, the Fkh1-induced origin re-
positioning assay was conducted by testing the effect on ARS305 positioning in cells
expressing Dbf4 lacking its C-terminus (dbf4∆C). The result showed that the deletion
- 47 -

of DBF4’s C-terminus phenocopies the origin localization in fkh1Δ cells which is consistent
with the mechanism that Fkh1 and Dbf4 are in the same pathway.

However, it cannot rule out the possibility that DDK and Fkh1 play independent roles in
regulating replication origins both temporally and spatially. To address this, we tagged
Fkh1 DNA-binding domain (Fkh-DBD) to DBF4 at its endogenous locus so that the full
function of Fkh1 was eliminated except its DNA binding ability. To determine whether the
Dbf4-Fkh can rescue the subnuclear location ARS305 from nuclear periphery in fkh1Δ cells,
the origin positioning assay was carried out in WT and Dbf4-Fkh fkh1Δ cells. The result
showed that the fusion protein Dbf4-Fkh was able to maintain the interior location of
ARS305 in the nucleus in fkh1Δ background, which reinforces the previous conclusion that
Fkh1 and Dbf4 are in the same pathway and Fkh1 acts an upstreaming role (Fig 2.2A).

Meanwhile, to examine whether the fusion protein Dbf4-Fkh was able to advance origin
replication, BrdU-IP-Seq was conducted on WT and Dbf4-Fkh fkh1Δ strains. In this case, the
early initiation of ARS305 was rescued by Dbf4-Fkh compared from fkh1Δ cells and
reached the WT level (Fig 2.2B). Strikingly, in the whole genome level analysis, not only
ARS305, but the replication timing defect caused by deleting FKH1 was fully rescued by
Dbf4-Fkh in the genome (Fig 2.2C left). The distinction of replication timing between WT
and Dbf4-Fkh fkh1Δ has no significant difference. Based on the scatter plot of WT and Dbf4-
Fkh fkh1Δ, the pattern of replication timing of Dbf4-Fkh fkh1Δ cells phenocopy WT cells (Fig
2.2C left). We grouped the genomic origins into Fkh-act, centromeric, and other origins
subgroup. Surprisingly, tagging Fkh-DBD to endogenous DBF4 did not affect the normal
- 48 -

distribution of Dbf4 to the genome inferred by their WT replication timing phenotype.
Therefore, the data support the mechanism that Fkh1 recruits Dbf4 through the same
pathway and impact both replication timing and subnuclear localization.

Figure 2.2 DBF4-FKH fkh1Δ but not DBF4-fkh-dsm phenocopies WT in both
replication timing and subnuclear localization. (A) ARS305-GFP NUP49-GFP strains
HYy192 (fkh1Δ), HYy195 (Dbf4-Fkh fkh1Δ) and HYy202 (Dbf4-fkh-dsm fkh1Δ) were
arrested in G1 with 1x -factor 2 hours at 25°C and images captured. Distances from origin
foci to nuclear periphery were determined, plotted as quartile boxplots, and analyzed by a
- 49 -

z-test. (B) Quantitative BrdU-IP-Seq (QBU) analysis was performed with strains CVy63
(wt), HYy192 (fkh1Δ), HYy195 (Dbf4-Fkh fkh1Δ) and HYy202 (Dbf4-fkh-dsm fkh1Δ) after G1
block-and-release into hydroxyurea in the presence of BrdU; averaged data from two
experimental replicates was plotted for Chromosome III. (C) A scatter plot shows the
average of two replicates. The results show expected global effects on Fkh-activated
origins, CEN-proximal origins and other origins.

Domain swap mutant of Dbf4-Fkh maintained the origin interior location in G1 and
partially rescued replication timing

To further demonstrate the function of domain-swapping motif in Fkh1 in regulating both
replication timing and subnuclear location, we introduced the domain swap mutations into
Dbf4-Fkh fusion protein in fkhΔ cells.
Both the origin re-positioning localization assay of ARS305 and BrdU-IP-seq experiment
were performed on Dbf4-fkh-dsm fkh1Δ and WT cells. The result displayed that Dbf4-fkh-
dsm was still able to be relocalized ARS305 to the interior of the nucleus in fkh1Δ
background compared to fkh1Δ single mutant cells (Fig 2.2A). In this case, introducing dsm
mutation into Dbf4-Fkh fusion protein is not sufficient to affect its ability on subnuclear
localization regulation. Meanwhile, in the BrdU-IP-seq analysis, ARS305 showed early
firing in Dbf4-fkh-dsm fkh1Δ cells compared to fkh1Δ despite its replication timing was not
rescued back to WT. Consistent with the previous study, this partial improvement of the
origin replication suggests the domain-swapping motif of Fkh1 plays an important role in
replication timing regulation. The fusion protein Dbf4-fkh-dsm is still able to bypass the
suppression of Fkh1 deletion on replication timing regulation despite introducing the
- 50 -

domain swap mutations. Therefore, the domain-swapping motif in Fkh1 is not an absolute
requirement for replication timing regulation in the fusion protein.

To further explore the effects of dsm mutations in Dbf4-Fkh on replication timing globally,
the initiation of replication origins in the whole genome was analyzed based on the
sequencing results. Specifically, not only ARS305, but also many other origins exhibited a
delayed firing in S phase as compared to WT and Dbf4-Fkh fkh1Δ cells (Fig. 2.2C right). The
Fkh-act origin group reveals an obvious late firing pattern compared to WT cells (red dots).
In addition, a significantly delayed replication timing was observed in the centromeric
origin group (green dots). Particularly, since the recruitment of Dbf4 is not affected by
tagging Fkh-DBD based on its WT timing phenotype in Dbf4-Fkh fkh1Δ cells, we propose
that the domain-swapping motif of Fkh1 is required for the recruitment of Dbf4.
Presumably, the defect of Dbf4 recruitment is mainly due to the inefficient interaction
between Dbf4 and fkh1-dsm. The dsm mutations in Forhead DNA binding domain result in
an undesirable protein structure, which negatively affects its interaction with Dbf4. The
incomplete recruitment of Dbf4 by fkh-dsm was not fully recovered by fusing these two
proteins together, which down-regulated origin initiation in later S phase. However, we
cannot rule out the possibility that this replication downregulation might due to the defect
of DNA binding in fkh1-dsm mutant. Previously, our lab showed Fkh-dsm showed relatively
less robust DNA binding as WT based on the result of ChIP-seq analysis (Ostrow et al.,
2017).

- 51 -

Replication timing regulation on centromeric origins by Ctf19 is independent of
Fkh1

Intriguingly, the notable late firing of centromeric origin subgroup showed a much more
severe delay than Fkh-act origins subgroup due to the effect of the domain swap mutant in
Dbf4-Fkh. Thus, the replication timing regulation of centromere-proximal origin (CEN-
origins) may be independent of Fkh1. Previously, we defined most of CEN-origins as Fkh-
repressed (Fkh-rep) origins based on the earlier initiation in fkhΔ compared to WT cell.
Together, origins in cells lacking Fkh1/2 may result from reduced competition from Fkh-
act origin subgroup for limiting initiation factors, rather than a direct repressive function of
Fkh1/2. Further supporting this was the study done by Natsume et al, which found that
Dbf4 accumulates at kinetochores through facilitation by Ctf19 in budding yeast.

Therefore, to examine whether the regulation of origin initiation regarding CEN-origins is
independent of Fkh1, a replication timing analysis was conducted (BrdU-IP-seq) by
deleting CTF19 in the fkhΔ background. The results showed that the CEN-origins showed
significant late firing while Fkh-act origins maintained their delayed replication timing as
in fkh1Δ single mutant (Fig 2.3A). Some representative CEN-origins and Fkh-act origins
were highlighted in the chromosome IX plot (Fig 2.3B). The delayed firing of Fkh-act
origins in fkhΔ cells remained late in ctf19ΔfkhΔ cells as additional CEN-origins displayed a
late firing pattern. So, this additive down-regulated replication pattern indicates that the
Ctf19 and Fkh1 act independently in CEN-origins and Fkh-act origins regulation
respectively.

- 52 -

Figure 2.3 Deletion of CTF19 in fkh1Δ background shows an additive replication
phenotype.
(A) Quantitative BrdU-IP-Seq (QBU) analysis was performed with strains CVy63 (wt),
HYy217 (fkh1Δ), HYy210 (ctf19Δ) and HYy208 (fkh1Δ ctf19Δ) described in Fig.2.1 legend. A
scatter plot shows the average of two replicates. The results show expected global effects
on Fkh-activated origins and CEN-proximal origins; (B) Averaged data from two
experimental replicates was plotted for Chromosome IX.
- 53 -

Discussion
Our results revealed a novel function of domain-swapping motif of Fkh1 on origin
subnuclear localization regulation beyond its replication timing control discovered in a
previous study (Ostrow et al., 2017). To dissect which domain of Fkh protein is required for
both regulations, we started from the domain-swapping motif of Fkh1 which is important
for the formation of Fkh protein dimerization. These interactions between Fkh protein
facilitate the early origin clustering in G1 phase based on chromosome conformation
capture analysis (Knott et al., 2012). In particular, our result shows that introducing dsm
mutations into domain-swapping motif restrains the interior location of early replication
origins in G1 phase. Consistent with previous discovery, the peripheral subnuclear location
in G1 links to late firing of S phase regulated by Fkh1.

In addition, Dbf4-fkh-dsm fkh1Δ did not suppress the interior location of early origin
ARS305 in G1, and partially up-regulated its replication timing compared to fkh1Δ cells.
With further exploration of different categories of replication origins in the genome, we
found Fkh-act origins and centromeric origins are the two groups indicating significantly
delayed replication timing. The late initiation of Fkh-act origin group is expected since the
Fkh dimerization is required for early origin clustering in G1 which is critical for early
initiation in later S phase (Knott et al., 2012; Ostrow et al., 2017). However, the replication
defect on CEN-proximal origins in Dbf4-fkh-dsm fkh1Δ cells is surprising especially since
tagging wild type Fkh-DBD to Dbf4 did not impair the normal recruitment of Dbf4 to other
loci in the genome, which phenocopies WT in terms of replication timing. In addition,
centromeric region dominates the early replication origins in cells lacking Fkh1/2,
- 54 -

consistent with the results that CENs facilitate origin initiation through Ctf19-Dbf4
interaction. Centromeres commonly cluster at an interior position in the nucleus. It is
considered that CENs overlap with the replication factory pools (Jin et al., 1998). Therefore,
the mechanism of early initiation of centromeric origin is independent of Fkh1 but mainly
regulated by Ctf19.

Thus, we propose that the domain-swapping motif of Fkh is necessary for Dbf4
recruitment. Presumably, Fkh-dsm results in a poor interaction between Forkhead protein
and Dbf4 due to the unfavorable protein structure of Fkh-dsm. In this case, even though the
Dbf4 was fused with fkh-dsm, the recruitment of Dbf4-fkh-dsm to genomic loci may be
weakened due to the undesirable protein structure or protein folding of the fusion protein.
In this scenario, the domain-swapping motif of Fkh plays a crucial role in both spatial and
temporal regulation on replication. To demonstrate and test the function of domain-
swapping motif of Fkh on Dbf4 recruitment, ChIP-seq of Dbf4 is planned to explore the DNA
binding condition in fkh1-dsm mutant in G1 phase. However, given that Fkh-dsm showed a
slight DNA binding defect compared to WT cells based on the result of ChIP-seq analysis
(Ostrow et al., 2017). Therefore, we cannot rule out the possibility that this replication
defect might be because of the weak DNA binding in fkh1-dsm mutant compared to wild
type domain-swapping motif.

The replication defect observed in both Fkh-act and CEN-proximal origin subgroups in
Dbf4-fkh-dsm fkh1Δ presumably was caused by the undesirable protein structure after
tagging fkh-dsm to the endogenous Dbf4. However, CEN-proximal origins revealed a more
- 55 -

significant delayed replication timing than Fkh-act origins in Dbf4-fkh-dsm fkh1Δ cells
compared to WT cells. These differences on origin initiation reveals the distinct efficiencies
of Dbf4 requirement to Fkh-act and CEN-proximal origin specifically. Thus, the interaction
of Dbf4 with Ctf19 and Fkh1 was affected by different degrees after introducing the dsm
mutation in Dbf4-Fkh. Therefore, the additive late firing replication pattern in the genome
after deleting of CTF19 in fkh1Δ cells indicates the independent pathways in timing
regulation of Ctf19 and Fkh1.

In the lack of Fkh1 and Fkh2, CEN-proximal origins dominate the early replication in the
genome, implying that CEN architecture can advance replication intrinsically. The early
origins clustered in the interior of the nucleus in G1 phase mediated by Fkh1 (Zhang et al.,
2019). Meanwhile, CENs also commonly occupy at the center of the nucleus, which is close
or overlaps with the other replication factories (Jin et al., 1998). Our findings are consistent
with the mechanism that the early initiation of centromeric origins is independent of
Fkh1/2. Thus, as Dbf4 recruiters, Ctf19 and Fkh are competing, and therefore limiting Dbf4
availability at Fkh-act and CEN origins respectively in G1-S transition. The CEN-proximal
early origins were defined as Fkh-repressed origins based on BrU-IP-seq experiment
(Knott et al, 2012). In this scenario, rather than a direct repressive function of Fkh1/2, the
advanced replication timing of CEN-proximal origins in cells lacking Fkh1/2 actually
results from the decreased competition from Fkh-act origins subgroup for limiting
initiation factors.

- 56 -

Appendix
Table 1. List of oligonucleotide DNA primer sequences

Primer Name Oligonucleotide sequence
NotI-ARS305-5' GGAATTGTGAGCGGATAACAATTTGTGGAATTGCGGCCGCcagtaatgaatat
tccaagt
XhoI-ARS305-5' GGAATTGctcgagTGAGAACAAGTTCGGATGTGGAATCgatagaagtaatttctat
at
NotI-ARS305-3' GTTGTAAAACGACGGCCAGTGAATTCGAGCTCGCGGCCGCagactatgtaatg
gtaaaga
KpnI-ARS305-3' GTGGAATTGTGAGCGGATAACAATTTGTGGAATTGGTACCatttaaatacata
tatatac
Nup49-GFP F atacgactcactatagggcgaattgggtaccgggcccccc
Nup49-GFP R caagcgcgcaattaaccctcactaaagggaacaaaagctg
TetR-Tom F ATCATGACGTAAGAAATGTATCTTAATTAAcgcgcgtatacgactcactatagggc
gaattgggtccc
TetR-Tom R ACACTGACATCTTTAACAACTTTTAATTAAtccaccgcggtggcggccgc
ARS501-tetO F tatagggcgaattgGGTACCCCAAGAACGATCCAATTGC
ARS501-tetO R agcttatcgataccgtcgacGGGTCAACTACCCTTCCC
ARS1103-tetO F gcgcgcgtaatacgactcactatagggcgaattgGAAACTAATTCAGATTTGGGTAAA
AG
ARS1103-tetO R TGATACGGATCCcccgggctgcaggaattcgatatcaagcttGGATGCTGGGTAAAT
GCC
- 57 -

ARS1303-tetO F gccagtgagcgcgcgtaatacgactcactatagggcgaattgCTCTGTGCCGTCTTCTCT
ARS1303-tetO R TACGGATCCcccgggctgcaggaattcgatatcaagcttGTAAGTAGTCTGCTTCAA
CCA
ARS710-lacO F acatgtggaattgtgagcggataacaatttgtggaattAAAGAGAGGCTGGTGACTTTT
C
ARS710-lacO R gacgttgtaaaacgacggccagtgaattcgagctcTGTAATCTCTAGTTATTAAGGAC
GC
ARS718-lacO F catgtggaattgtgagcggataacaatttgtggaattCTTATGGGAAACTGGTTACATT
C
ARS718-lacO R gtcacgacgttgtaaaacgacggccagtgaattcgagctcGAGCTGATGAACCTTCTGT
T
ARS1018-lacO F gccacatgtggaattgtgagcggataacaatttgtggaattTTGAAGTCATCTACTGCCC
ARS1018-lacO R acgacgttgtaaaacgacggccagtgaattcgagctcACATTACTATAATGAAAGCCG
AG
ARS305-ΔACS-
mut1
GGGAAAATAAACAATACATAACAAAcgAggTAAAAACCAACACA
ARS305-ΔACS-
mut2
GGGAAAATAAACAATACATAACcctcgAggTAAAAACCAACACA
ARS305-Δ2BS-
mut1
GTGTTGGTTTTTATATGTTTTGCTCGAGATTGTTTATTTTCCC
ARS305-Δ2BS-
mut2
GCTTTAAGAACTACAAAGTAGGTACCAAATAATAAATCACACCG
- 58 -

FKH1-up CAGAAACGGTATAGAGAGAACAGG
FKH1-down CACAGAGGGTACAGAAGTCATAAAG
DBF4-up CAGCCACTATAGCAACTACTGC
DBF4-down ATTAACCGCGGTGGGTACTC
Cdc45-F cagtgagcgcgcgtaatacgactcactatagggcgaattgATGTATTATGGAATCAGCC
A
Cdc45-R aagcgcgcaattaaccctcactaaagggaacaaaagctgTTATAACAATCCACTCAAGG
T
ADE2-up-F AACTTTTAATTAAGATACATTTCTTACGTCATGATTGATTATTACAGCT
ATGC
ADE2-int-R aatacgactcactatagggcgaattggagctcCTTTACAACGAAGTTACCTCTTCCAT
CG
ADE2-farup-F ccctcactaaagggaacaaaagctgggtaccCCTTTTGATGCGGAATTGACTTTTTC
TTG
ADE2-up-R GACGTAAGAAATGTATCTTAATTAAAAGTTGTTAAAGATGTCAGTGTTA
TGTTGGTG
Fkh-Myc-F GCCCCGCAATTACAAAGAACTCAACTCACT
Fkh-Myc-R AATTCATTCAATAAAAAGATATAAATCGAGATGATCGTTCCACTTTTTA
GCTAGTGGATC
Ctf19-F GTGTGATCTTGTTGATACTAGGTCGGCAAAGAACGCAAATCGGATCCCC
GGGTTAATTAA
- 59 -

Ctf19-R GTTTAAGCAAGCCGTCCAGTTGGCAATGGCAAATGGAACAGAATTCGAG
CTCGTTTAAAC

Table 2. List of yeast strains

Strain Genotype (strains share the SSy161 genotype except as noted)
SSy161 MATa, ade2-1, leu2-3,112, his3-11,15 trp1-1, ura3-1, can1-100, bar1∆::hisG
HYy38 fkh1∆::KanMx, fkh2-dsm, leu2::GAL-FKH1::LEU2, ars501∆::ARS305-tetO-URA3,
ade2::TetR-Tomato::ADE2, ars305∆::BrdU-Inc-TRP1
HYy82 FKH1-3XFLAG-TRP1, leu2::GAL-FKH1::LEU2, ars501∆::ars305::tetO-URA3,
ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1
HYy85 FKH1-3XFLAG-TRP1, leu2::GAL-FKH1::LEU2, ars501∆::ars305-∆ACS::tetO-
URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1
HYy108 FKH1-3XFLAG-TRP1, leu2::GAL-FKH1::LEU2, ars501∆::ars305-∆2BS::tetO-
URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1
HYy156 fkh1∆::KanMx, fkh2-dsm, cdc7-as3, leu2::GAL-FKH1::LEU2,
ars501∆::ARS305::tetO-URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-
GFP::TRP1
HYy158 cdc7-as3, leu2::GAL-FKH1::LEU2, ars501∆::ARS305::tetO-URA3, ade2::TetR-
Tomato::ADE2, trp1::NUP49-GFP::TRP1
HYy108 FKH1-3XFLAG-TRP1, leu2::GAL-FKH1::LEU2, ars501∆::ars305-∆2BS::tetO-
URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1
- 60 -

HYy147 ARS305::lacO-LEU2, trp1::LacI-GFP::TRP1, ARS607::tetO-URA3, ade2::TetR-
Tomato::ADE2, his3::NUP49-GFP::HIS3, fkh1∆::KanMx
HYy151 ARS305::lacO-LEU2, trp1::LacI-GFP::TRP1, ade2::TetR-Tomato::ADE2,
his3::NUP49-GFP::HIS3
HYy157 ars501∆::ARS305::tetO-URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-
GFP::TRP1
HYy160 ARS501::tetO-URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1,
leu2::GAL-FKH1::LEU2
HYy165 ARS1103::tetO-URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1,
leu2::GAL-FKH1::LEU2
HYy166 ARS1303::tetO-URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1,
leu2::GAL-FKH1::LEU2
HYy172 ARS1103::tetO-URA3, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1,
leu2::GAL-FKH1::LEU2, fkh1∆::KanMx
HYy173 ARS1303::tetO-URA, ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1,
leu2::GAL-FKH1::LEU2, fkh1∆::KanMx
HYy177 fkh1∆::KanMx, fkh2-dsm, leu2::GAL-FKH1::LEU2, ars501∆::ARS305::tetO-URA3,
ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1, cdc7-as3, mcm4::NatMX4-
pMCM5-MCM4[D(D/E)+DS(P/Q)]
HYy181 ARS305::lacO-LEU2, trp1::LacI-GFP::TRP1, ade2::TetR-Tomato::ADE2,
his3::NUP49-GFP::HIS3, dbf4∆C::KanMX
- 61 -

HYy183 ARS305::lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3,
clb5∆::URA3, clb6∆::KanMX
HYy184 ARS305::lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3, cdc45-1
HYy186 fkh1∆::KanMx, fkh2-dsm, leu2::GAL-FKH1::LEU2, ars501∆::ARS305::tetO-URA3,
ade2::TetR-Tomato::ADE2, trp1::NUP49-GFP::TRP1, cdc7-as3
HYy191 ARS305::lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3,
bar1∆::KanMX, cdc7-4
MPy6 ARS710-lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3
MPy10 ARS710-lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3,
fkh1∆::KanMx
MPy19 ARS1018-lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3
MPy20 ARS718-lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3
MPy21 ARS718-lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3,
fkh1∆::KanMx
MPy22 ARS1018-lacO-LEU2, trp1::LacI-GFP::TRP1, his3::NUP49-GFP::HIS3,
fkh1∆::KanMx
OAy110
2
FKH1-3XFLAG-TRP1

- 62 -

Figure A1. Fkh1 binding was eliminated at ars305- ∆2BS
V-R

FKH1-FLAG-tagged strains HYy82 (ARS305
V-R
), HYy85 (ars305-∆ACS
V-R
), and HYy108
(ars305-∆2BS
V-R
) were treated as above and subjected to ChIP-seq analysis; averaged data
from duplicate experiments was plotted for the V-R region.

- 63 -

Figure A2. cdc7as3 allele is not stringent enough to affect the subnuclear location of
ARS305 by inhibiting DDK function.

FACS analysis of DNA content of HYy151(WT), HYy158(cdc7as3) and HYy156 (fkh1Δ
fkh2dsm cdc7as3) cells synchronized in G1-phase with -factor and released synchronously
into S-phase.

- 64 -

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Zhang, H., Petrie, M. V., He, Y., Peace, J. M., Chiolo, I. E., & Aparicio, O. M. (2019). Dynamic
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- 79 -

Supplemental Figures

Figure 1-figure supplement 1. Fkh1-induction is required to re-position a sub-
telomeric origin in G1 phase. (A) HYy80 (fkh1∆ fkh2-dsm ARS305
V-R
-Tomato NUP49-GFP)
cells were treated and analyzed as in Figure 1C and D legends. (B) HYy132 (fkh1∆ fkh2-dsm
GAL-FKH1 ARS305
V-R
-Tomato NUP49-GFP) cells were treated as in Figure 1C legend, except
- 80 -

that dextrose was substituted for galactose, and analyzed as in Figure 1D legend. The
galactose data are the same as in Figure 1D.

Figure 2-figure supplement 1. Normal dosage of Fkh1 is sufficient to advance firing
time of ARS305
V-R
. (A) QBU analysis was performed as described in Figure 3C legend and
presented as a scatter plot showing average of three replicates. The results show expected
- 81 -

global effects on Fkh-activated origins while highlighting ARS305
V-R
. (B) Scatter plots show
comparison of individual experimental replicates for the above experiment.

Figure 3-figure supplement 1. Fkh1 determines early origin positioning globally. (A)
Cells with ARS305-GFP (HYy151), ARS305-∆2BS-GFP (MPy46), and ars305-∆ACS-GFP
- 82 -

(MPy43) all expressing NUP49-GFP, were treated and analyzed as in Figure 3 legend; scale
bar = 0.5 m. (B) Cycling cultures of WT and fkh1∆ strains with ARS305-GFP (HYy151,
HYy147) and ARS710-GFP (MPy6, MPy10) were imaged and G1 phase (unbudded) cells
were analyzed as described in Figure 3 legend. (C) Cells of WT and fkh1∆ strains with
ARS501-Tom (HYy198, HYy201) were arrested in G1 phase and analyzed as described in
Figure 3 legend.
- 83 -

Figure 4-figure supplement 1. Fkh1 binds origins independently of Cdc7 function.
FKH1-FLAG-tagged strains OAy1102 (WT) and HYy123 (cdc7-as3) were arrested in G1 at
23°C and treated with PP1 or DMSO (vehicle), and subjected to ChIP-seq analysis. (A)
Averaged data from duplicate experiments is plotted for the ARS305 chromosomal region.
A
B
10
5
0
5 10 0
10
5
0
5 10 0
10
5
0
5 10 0
10
5
0
5 10 0
10
5
0
5 10 0
WT + DMSO
cdc7-as3 + DMSO
WT#1 + DMSO
WT#2 + DMSO
WT#1 + PP1
cdc7-as3#2 + DMSO
WT#2 + PP1
cdc7-as3#1 + DMSO cdc7-as3#1 + PP1
cdc7-as3#2 + PP1
Fkh-activated origins
10
5
0
5 10 0
WT + PP1
cdc7-as3 + PP1
cdc7-as3 + PP1
3.5 4.0 4.5
7.5
15
Chr.III Coord (bp/10
4
)
0
3.5 4.0 4.5
7.5
15
0
WT + DMSO
3.5 4.0 4.5
7.5
15
0
3.5 4.0 4.5
7.5
15
Chr.III Coord (bp/10
4
)
0
ARS305
ARS305
ARS305
ARS305
cdc7-as3 + DMSO
WT + PP1
FigureS4
- 84 -

(B) Data for Fkh-activated origins are plotted. The upper panels present averaged data
from the individual replicates presented in the middle and lower panels.

Figure 5-figure supplement 1. CDK activity is dispensable for origin localization in
G1. (A) Cells treated as described in Figure 5D legend were released from -factor block
with PP1 or DMSO and budding morphology was quantified and plotted. (B) ARS305-GFP
cdc28-as1 cells (HYy197) were treated with PP1 or DMSO for two hours at 25°C and origin
localization was analyzed in G1 phase (unbudded) cells as described in Figure 5 legend. (C)
Cells treated as in (B) were subjected to DNA content analysis; the brackets estimate the
proportion of cells in S phase based on DNA content between 1C and 2C.

Abstract (if available)

Abstract Chromosomal DNA elements are organized into spatial domains within the eukaryotic nucleus. Sites undergoing DNA replication, high-level transcription, and repair of double-strand breaks coalesce into foci, although the significance and mechanisms giving rise to these dynamic structures are poorly understood. In S. cerevisiae, replication origins exhibit characteristic subnuclear localizations prior to S phase that anticipate their initiation timing and/or efficiency during S phase: origins found within the nuclear interior in G1 phase initiate early and efficiently in S phase while origins found associated with the nuclear periphery in G1 phase initiate later and less efficiently. Here, we link localization of replication origins in G1 phase with Fkh1 activity, which is required for their early replication timing. Using a Fkh1-dependent origin relocalization assay, we determine that execution of Dbf4-dependent kinase function, including Cdc45 loading, results in dynamic relocalization of a replication origin from the nuclear periphery to the interior in G1 phase. Origin mobility increases substantially with Fkh1-driven relocalization. Specifically, domain-swapping motif of Fkh1 is crucial for the regulation of both replication timing and subnuclear location, which is required for Dbf4 recruitment. The replication timing of centromeric origins is controlled by Ctf19 but independent of Fkh1. These findings provide novel molecular insight into the mechanisms that govern dynamics and spatial organization of DNA replication origins and possibly other functional DNA element.

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Asset Metadata

Creator Zhang, Haiyang (author)

Core Title Subnuclear localization of replication origins is controlled by Fkh1-dependent recruitment of DDK to origins in S. cerevisiae

School College of Letters, Arts and Sciences

Degree Doctor of Philosophy

Degree Program Molecular Biology

Publication Date 02/23/2021

Defense Date 12/10/2020

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

Tag Cdc45,Dbf4-dependent kinase,DNA replication,Fkh1,mobility,OAI-PMH Harvest,subnuclear localization

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor Aparicio, Oscar (committee chair), Chiolo, Irene (committee member), Curran, Sean (committee member), Michael, Matthew (committee member)

Creator Email zhan731@usc.edu,zhanghy0516@gmail.com

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c89-421555

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Legacy Identifier etd-ZhangHaiya-9286.pdf

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

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...

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