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The role of Cdc7 in replication fork progression in response to DNA damage
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The role of Cdc7 in replication fork progression in response to DNA damage
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Content
THE ROLE OF CDC7 IN REPLICATION FORK PROGRESSION IN RESPONSE TO
DNA DAMAGE
by
Yuan Zhong
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 2012
Copyright 2012 Yuan Zhong
ii
Acknowledgements
I would like to thank my advisor Dr. Oscar Aparicio who constantly encourages and
motivates me to pursue this study. From him, I learned not only a great amount of
knowledge and critical thinking but also the spirit that one should not give up no matter
how frustrated you are. There were so many times that he really could calm me down and
went back to the basic problems trying to help me to solve the problems without losing
time and energy on criticizing me and showing his upsets on my stupid mistakes. His
love to science and curiosity to wonder really affected the way I am thinking. I also
would like to thank my committee members, especially Dr. Susan Forsburg and Dr.
Matthew Michael who gave me valuable suggestions in our group meetings. I also would
like to thank our current and past lab members who created a wonderful environment for
me to study science and meanwhile to enjoy the life of graduate school at some relaxing
moments so that I could refresh myself to carry on. Especially I would like to thank
Simon Knott and Tittu Thomas for their work for analyzing the microarray data and their
patience to explain the computational analysis to me. I also would like to thank Sandra
Villwock who showed me how to do 2-D gel analysis. Jared Peace and Zack Ostrow have
been encouraging me when I am so down and cheered me up when I needed. Also I
would like to thank Lin Ding who is one of the true friends of mine, who is always there
to watch me trying to figure out what I need and gave me a hand to reach. I also would
like to thank all the undergraduates who worked with me, especially Janis Yee and
Sanket Rege who brought lots of fun and lots of help to me. More importantly, I would
iii
like to thank my husband Yunxiang Mu who tolerated my bad temper when I was
stressed out and has been taken lots of family responsibilities to free me to focus on my
own work. My daughter, Nova Mu, helps me to relax by playing with her. I also would
like to thank my parents who came to help me with Nova and my brother who has been
taking care of my parents at home to let me concentrate on the study here. I would like to
thank WiSE program, which has supported me both financially and emotionally during
my pregnancy and the childcare.
iv
Table of Contents
Acknowledgements ii
List of Figures vii
Abstract x
Chapter 1: Introduction 1
1.1 The initiation of replication 1
1.2 DNA Replication in response to MMS 2
1.3 Checkpoint activation 3
1.4 Post-Replication Repair Pathways (PRR) 4
1.5 Dbf4-dependent kinase 4
Chapter 2: Cdc7 functions in replication fork progression
In response to MMS 9
Chapter 3: Characterization of the relationship between
checkpoint activation and fork rate in response to MMS 18
3.1 Cdc7 is required for proper checkpoint activation
in response to replication stress 18
3.2 Pph3 deletion can partially antagonize the deregulated fork
in MMS in cdc7-1/mcm5-bob1 21
3.3 DDK regulation by Rad53 in response to DNA damage
does not play a role in fork progression 23
3.4 Other checkpoint deficiency strains cannot
phenocopy the faster fork movement in MMS 25
Chapter 4: The choice of Post Replication repair
pathways and fork progression rate in
response to MMS 29
4.1 Deletion of TLS polymerases has no effect
on the replication fork rate in MMS
and on the faster fork in MMS in cdc7-1/mcm5-bob1 30
4.2 Rad5 is indispensable for replication fork progression
in MMS in the absence of Cdc7 33
v
Chapter 5: The mechanism for the synergistic defect
between Rad5 and DDK in response to MMS 36
5.1 CDC7 has synergistic defect with both the helicase
and E
3
ligase function in RAD5 36
5.2 Elimination of TLS and Rad5 pathways showed
no synergistic defects in replication fork progression
in response to MMS 39
5.3 cdc7-1/mcm5-bob1 rad5∆ double mutant cannot recover
after MMS exposure 42
5.4 rad5∆ is not synergistic with mec1-100 to 0.033%MMS 44
Chapter 6: Detecting ssDNA gaps behind
replication forks in Cdc7 mutant 47
Chapter 7: ORC depletion deregulates replication fork progression in MMS 50
Chapter 8: DDK regulation on MCM2 in response to MMS 52
Chapter 9: Material and Methods 55
9.1 Plasmid and strain construction 55
9.2 DNA content analysis 56
9.3 BrdU-IP-chip 56
9.4 Viability assay 57
9.5 Pulse Field Gel Electrophoresis 58
9.6 Western blot 59
Chapter 10: Discussion 60
10.1 Replication fork progression is regulated in response
to DNA damage 60
10.2 Loss of Cdc7 function creates a critical dependence on Rad5
to complete DNA replication 63
10.3 Cdc7 functions in checkpoint activation in response
to DNA damage 67
Bibliography 70
Appendices
Appendix A: Cdc7 is required for fork stalling in response to HU 77
Appendix B: Post-replication Repair Pathway and
replication fork progression rate in MMS 79
Appendix C: PFGE analysis for chromosome replication
in constant MMS treatment in WT and cdc7-1/mcm5-bob1 80
Appendix D: The synergist between Rad5 deletion and cdc7as3 81
vi
Appendix E: Pph3 deletion can not rescue the sensitivity of
cdc7-1/mcm5-bob1 rad5∆ to MMS 82
Appendix F: Fork progression and checkpoint activation after
cdc7as3 inactivation at latter S Phase 83
Appendix G: List of Strains 85
Table G Strains used in this study 85
vii
List of Figures
Figure 2.1 FACS analysis for DNA content
in WT and cdc7as3 without PP1 10
Figure 2.2 FACS analysis for DNA content in WT and cdc7as3
in the presence of PP1 with and without MMS 11
Figure 2.3 Replication fork failed to slow down in response
to MMS in cdc7as3 strain 13
Figure 2.4 FACS analysis for S phase progression in YEPD
in WT, and cdc7-1/ mcm5-bob1 15
Figure 2.5 Replication fork failed to slow down in response to
MMS in cdc7-1/mcm5-bob1 strain 16
Figure 3.1 Rad53 activation in Cdc7 mutant during S phase
in response to DNA damage 19
Figure 3.2 Cdc7 is needed for maintaining checkpoint activation 20
Figure 3.3 Pph3 deletion can partially antagonize
the deregulated faster fork in MMS in cdc7-1/mcm5-bob1 22
Figure 3.4 pph3∆ can restore checkpoint activation deficiency
in cdc7-1/mcm5-bob1 mutant 23
Figure 3.5 Deregulation of Dbf4 by Rad53
does not affect the fork rate in MMS 24
Figure 3.6 Fork progression in mrc1-AQ is normal in response to MMS 26
Figure 3.7 mec1-100 has delayed checkpoint activation
but normal fork rate in MMS 27
Figure 4.1 Genetic interaction between Cdc7 and different components
of post replication repair pathways in response to MMS 30
viii
Figure 4.2 Deletion of TLS polymerases has no effect
on the replication fork rate and also has no effect
on the faster fork in MMS in cdc7-1/mcm5-bob1 32
Figure 4.3 Rad5 is required for DNA synthesis
by FACS analysis in the absence of Cdc7 34
Figure 4.4 Rad5 is required for DNA synthesis in the absence of Cdc7 35
Figure 5.1 CDC7 has synergistic defect with both the helicase
and E3 ligase function in RAD5 38
Figure 5.2 Genetic interaction between Cdc7
and Rad5 GAA (helicase-) and Rad5I916A (E3 ligase-) 39
Figure 5.3 TLS∆ and Rad5∆ show no synergistic defects
in response to MMS 41
Figure 5.4 Rad53 activation in rad5∆
and in cdc7-1/mcm5-bob1 rad5∆ double mutant 43
Figure 5.5 cdc7-1/mcm5-bob1 rad5∆ double mutant could not recover
after 1 hr 0.033% MMS exposure 44
Figure 5.6 rad5∆ did not show synergistic defect with mec1-100 45
Figure 5.7 Rad5 deletion doesn’t show synergistic defect
with mec1-100 to 0.033% MMS 46
Figure 6.1 RPA-ChIP-Chip in WT and cdc7-1/mcm5-bob1 47
Figure 6.2 cdc7-1/mcm5-bob1 is defective in recovery
after MMS exposure by PFGE 49
Figure 7.1 orc1-161 also showed accelerated fork rate in MMS 51
Figure 8.1 The mcm2AA mutant showed
normal replication fork progression in MMS 53
Figure A.1 Cdc7 is needed for maintaining checkpoint activation
in response to HU 77
Figure A.2 Cdc7 is required for proper fork stalling
in 0.2M HU 77
ix
Figure A.3 RPA-ChIP-Chip in WT and cdc7-1/mcm5-bob1
in response to 0.2M HU 78
Figure B.1 rad18∆ siz1∆ does not affect fork rate
in response to 0.033% MMS 79
Figure C.1 PFGE analysis for chromosome replication
in constant MMS treatment in WT and cdc7-1/mcm5-bob1 80
Figure D.1 The synergist between Rad5 deletion and cdc7as3 81
Figure E.1 Deletion of Pph3 does not rescue
the sensitivity of cdc7-1/mcm5-bob1 to MMS 82
Figure E.2 pph3∆ could not rescue the synergistic defect
of rad5 and cdc7-1/mcm5-bob1 82
Figure F.1 Fork progression and checkpoint activation after
inactivation of cdc7as3 at later S Phase 83
x
Abstract
Cdc7-Dbf4 is an essential protein kinase complex required for every single origin
firing. As a target of the intra-S checkpoint, Cdc7 kinase activity has also been implicated
in the response to replication fork stress, with a role in translesion DNA synthesis (TLS).
We have examined the role of Cdc7 in the regulation of replication forks, particularly in
response to MMS, which normally stalls replication forks and inhibits late origin firing.
We find that replication forks proceed as fast as with no damage along an MMS-damaged
template both in cdc7as3 and cdc7-1/mcm5-bob1 cells. However the DNA synthesis in
cdc7-1/mcm5-bob1 in MMS is defective, indicated by the slower recovery after MMS by
PFGE, suggesting the replication is incomplete. These deregulated forks did not rely on
TLS pathway but are dependent on both helicase and E3 ligase function of Rad5 for
continued fork progression along MMS-damaged DNA, demonstrating a role for Rad5 at
the replication fork. Phosphorylation of MCM2 by DDK was not sufficient for slowing
down Mcm2-7 helicase activity in vivo. Temperature-sensitive mutant orc1-161, which is
defective in pre-replication complex (pre-RC) assembly, phenocopies the defects in
origin firing and faster fork progression in MMS in Cdc7 mutants, suggesting that
decreased origin firing is the common source for the deregulated fork progression. We
attribute the effect of Cdc7 depletion on replication fork progression in MMS to the
reduced origin-firing, which leads to less established forks and in turn the deficient
checkpoint activation in MMS. The mec1-100 cells, which initiate many origins but also
have compromised checkpoint activation, fail to phenocopy the faster fork movement in
xi
MMS, suggesting that the number of active replication forks also influences fork rate,
perhaps due to competition for limiting factors for DNA replication. These findings
provide new insights into how Cdc7, besides its essential role in replication initiation,
could imply a function in the regulation of replication fork progression in response to
DNA damage possibly through checkpoint activation and replication fork restart in MMS.
1
Chapter 1: Introduction
1.1 The initiation of replication
The replication of chromosomes in S phase in Saccharomyces cerevisiae relies on the cell
cycle-regulated activation of numerous replication origins throughout the genome
following a strict timing profile with some origins firing early and some origins firing late.
The components of replication machinery in budding yeast are well characterized. From
late M to G1 phase, the ORC
1-6
complex, CDC6, CDT1 and the MCM
2-7
complex are
sequentially recruited onto replication origins to form the pre-replicative complex (pre-
RC) (Robinson et al., 2005). An important step in replication initiation to convert pre-RC
to pre-IC (pre-initiation complex) is the conversion of MCM2-7 into the active helicase,
resulting in DNA unwinding, replisome assembly and DNA synthesis. Once cells enter S
phase, Dbf4-dependent kinase (DDK) phosphorylates MCM2-7 facilitating the
recruitment of Sld3 and Cdc45. Then S-phase cyclin-dependent kinase (S-CDK)
phosphorylates Sld3 to recruit Sld2, Dpb11, GINS and Pol ε and Mcm10. The formation
of Cdc45-Mcm2-7-GINS complex activates the helicase which triggers the unwinding of
origin DNA. Pol α and Pol δ are loaded on the unwound DNA in a Mcm10-dependent
process to complete the replisome assembly (Heller et al., 2011). So the activation of pre-
RC into pre-IC to become active replisome involves two highly conserved kinases, S-
CDK and DDK to coordinately regulate this process, both of which become active at the
G1-S transition. DNA unwinding helicase activity is not activated until the
phosphorylation events catalyzed by CDK and DDK are achieved.
2
1.2 DNA Replication in response to MMS
Methyl-methanesulfonate (MMS) is an alkylating reagent which modifies both guanine
(to 7-methylguanine) and adenine (to 3-methlyladenine) to cause replication blocks
(Beranek, 1990). When cells enter S phase in the presence of DNA damage reagent, the S
phase is dramatically slowed down. In Saccharomyces cerevisiae, the slowed S phase
resulting from MMS treatment depends on checkpoint signaling controlled by Mec1 and
Rad53, indicating that slowing of DNA synthesis is a regulated process (Paulovich et al.,
1995), The prolonged S phase in cells treated with MMS is the result of checkpoint-
dependent regulation of late origin firing and slower fork movement (Tercero et al.,
2001). The checkpoint-dependent late origin firing inhibition is supported by the de-
suppression of late origin firing in checkpoint mutants. Furthermore, the inhibition of late
origin firing is bypassed by mutation of the Rad53 phosphorylation sites on Sld3 and
Dbf4 (Lopez-Mosqueda et al., 2010, Zegerman et al., 2010). In addition to the inhibition
of late origin firing, the fork progression rate in the presence of MMS is dramatically
slowed down. It has been a question whether this fork slowing is regulated process or
simply the result of the passive physical obstacle to the replication machinery from the
lesions on the templates. It is very hard to distinguish between an active slowing of
replication fork progression and a passive effect of lesions blocking the forks, because the
intra-S checkpoint is required for stabilization of stalled forks. The forks stalled by the
lesion on the template collapse in checkpoint mutants (Lopes et al., 2001). However it
has been shown that deletion of Exo1 suppresses DNA replication fork instability in
3
Rad53 mutants (Segurado et al., 2008, Lopez-Mosqueda et al., 2010). Also it has been
shown that the dephosphorylation of Rad53 by Pph3-Psy2 is required for replication fork
restart during recovery from DNA damage. pph3∆ cells have extremely slower fork
movement in MMS (O'Neill et al., 2007, Szyjka et al., 2008) with highly activated
checkpoint, indicating that the process for slowing fork is a regulated process through
checkpoint activation. However, no evidence has been shown that replication fork could
actually progress faster or fail to slow down in the presence of DNA damage if there were
less checkpoint activation. So the mechanism controlling fork stability or fork movement
in MMS is not very clear.
1.3 Checkpoint activation
The DNA lesion on the template strand will activate the checkpoint generated from the
forks established from the early origins. The intra-S phase checkpoint is administered by
three classes of proteins: the sensors (eg. Mec1-Ddc2 and Tel1 and 9-1-1 complex RFC
complex) which directly sense the DNA damage displayed by RPA coated ssDNA, and
the effector protein kinases Rad53 (Chk2) and Chk1 which target the cell cycle
machinery to inhibit cell cycle progression. In between these two classes are the adaptors
(Rad9 and Mrc1) which appear to mediate the signaling from the sensors to the effectors
mainly through recruiting the effectors to the damage site (Melo et al., 2002). In addition
to all these functional proteins as checkpoint component, replication fork itself functions
as a sensor for activating the checkpoint (Tercero et al., 2003): the fewer forks
established, the less checkpoint activation.
4
1.4 Post-Replication Repair Pathways (PRR)
To avoid cell cycle arrest and replication fork collapse, cells have evolved multiple
mechanisms by which the arrested or stalled replication forks can be rescued. It involves
two main DNA damage tolerance and DNA repair pathways: the Rad6/Rad18 post
replication repair pathway (PRR) and Rad52 dependent homologous recombination (HR)
pathways (Branzei et al., 2010). In Rad6/Rad18 repair pathway, there are two modes of
PRR. One is error prone mechanism, which uses non-processive translesion polymerases
(Rad30, Rev3) to carry out nucleotide incorporation opposite a damaged base in the DNA
template that might incorporate mutations and require mono-ubiquitination of PCNA on
Lys
164
mediated by Rad6/Rad18. Another is error-free mechanism which additionally
requires MMS2, UBC13 and RAD5 mediated poly-ubiquitylation of PCNA through
Lysine
63
(Hoege et al., 2002). Inhibition of Rad52-mediated HR repair pathway is
mediated by SUMO modified PCNA by Ubc9 and Siz1 through recruiting Srs2 to disrupt
Rad51 nucleoprotein filaments (Pfander et al., 2005).
1.5 Dbf4-dependent kinase
Dbf4-dependent kinase (DDK) is composed of a catalytic subunit Cdc7 and a regulatory
subunit Dbf4. Both subunits are very well conserved from yeast to humans and are
essential for cell viability. Its activity is required throughout S phase for the firing of
individual replication origin (Bousset et al., 1998, Donaldson et al., 1998). Its function in
initiation of DNA replication has been extensively examined. The essential role for DDK
5
is to activate the helicase activity by the phosphorylation of the replicative helicase
MCM
2-7
complex. A mutant of the Mcm5 subunit (mcm5-bob1) and an N-terminal
truncation mutant of Mcm4 (mcm4
Δ74–174
) can bypass the essential function of Cdc7 in
the initiation of DNA replication (Sclafani et al., 2002, Sheu et al., 2010). Whether its
activity is continuously or constantly required during the unperturbed fork progression
after the MCMs complex is activated has not been studied.
In budding yeast, besides its role in replication initiation, it has been found that Cdc7
prevents UV sensitivity and promotes UV-induced mutagenesis (Njagi et al., 1982). In
fission yeast, mutants of hsk1(sc-cdc7) and dfp1(sc-dbf4) are also defective for MMS-
induced mutagenesis (Dolan et al., 2010). Conversely, ectopic over-expression of Cdc7
shows hyper-mutability in response to UV (Sclafani et al., 1988). DDK is also involved
in Rad6 post replication repair pathway, implying a possible role for Cdc7 in translesion
DNA synthesis and is required for induced mutagenesis (Njagi et al., 1982, Pessoa-
Brandao et al., 2004, Dolan et al., 2010). Recently, Cdc7 has been shown to
phosphorylate Rad18 to direct DNA polymerase η to sites of stalled replication (Day et
al., 2010). However the role of Cdc7 in TLS does not explain additional phenotypes
associated with its loss-of-function. Cdc7 is essential for origin firing, but neither of the
two bypass mutants, mcm5-bob1 and mcm4Δ2–174, can bypass the sensitivity of Cdc7
mutant to genotoxic reagent, nor have the proper S phase checkpoint activation (Sheu et
al., 2010). In fission yeast, a hsk1 temperature-sensitive mutant released to the restrictive
temperature after early S-phase arrests in hydroxyurea (HU) is able to complete bulk
6
DNA synthesis, but undergoes an abnormal mitosis (Snaith et al., 2000). These results
suggest that DDK plays a role in later S phase after the initiation of replication step.
Studies of hsk1
ts
cells show a defect in Cds1 checkpoint kinase activation along with
accumulation of Rad22 (Rad52 ortholog) recombinational repair foci, indicating that
Hsk1 functions in replication fork stabilization. Also it has been shown that Hsk1
physically interacts with Swi1 and Swi3, components of the replication fork protection
complex (Matsumoto et al., 2005). It has been shown that Hsk1 and Dfp1 were chromatin
bound during S and G2 phase and this interaction is disrupted in C terminal truncation of
Dfp1 (Dolan et al., 2010). All of these indicate that Cdc7 is playing an important role in
replication fork after the initiation step.
Whether Cdc7 is really involved in maintaining fork stability and checkpoint activation is
not clear. Whether Cdc7 plays a role in the replication fork progression by channeling the
replication fork restart through different post replication repair pathways in response to
DNA damage is also not known. It remains unclear whether Cdc7 regulates MCMs
complex function at the replication forks, especially when forks encounter the damage on
the template. It remains unclear whether the checkpoint defects and HU sensitivity
resulting from Cdc7/Hsk1 depletion reflect a direct role of DDK in checkpoint signaling,
or indirect effects of decreased early origin firing. Since replication forks are the sensors
for generating the checkpoint signaling (Tercero et al., 2003), decreased early origin
firing may generate fewer replication forks and hence weaker checkpoint signaling.
7
Recently, Szyjka and colleagues showed that Rad53 activation regulates the rate of
replication fork progression possibly through affecting the fork restart after fork stalling
by the damaged template (Szyjka et al., 2008). Based on the fact that Cdc7 regulatory
subunit Dbf4 is a target of Rad53 in response to DNA damage, it is reasonable to
hypothesize that Rad53 decides the choice of fork restart mechanism through regulating
DDK by phosphorylation of Dbf4 possibly through retargeting DDK from the origins to
forks.
The replication fork dynamics in MMS was examined in cells depleted of Cdc7 function
and showed that replication fork failed to slow down in MMS in Cdc7 mutants. This
effect is related with its function in initiation but not related with its possible role in
translesion DNA synthesis. It is the checkpoint activation deficiency which is caused by
less origin firing in Cdc7 mutant that makes the cells fail to slow the fork progression.
This conclusion was further supported by another initiation defect mutant orc1-161
which is only deficient in Pre-RC complex assembly. In contrast, mec1-100 cells, which
also has compromised checkpoint activation manifested by de-suppressed late origin
firing in response to MMS, doesn’t have faster fork movement as Cdc7 and Orc1 mutant,
suggesting decreased checkpoint activation alone is not sufficient to have faster fork
movement in MMS. We conclude that replication fork rate is under checkpoint-
dependent and independent regulation. Interestingly, the DNA synthesis through the
faster fork movement in Cdc7 mutant is Rad5 dependent and Rad5 maintains the viability
8
of Cdc7 in response to MMS. Also it seems that the DNA synthesis through the faster
fork in Cdc7 mutant is not fully complete, perhaps leaving gaps behinds the fork.
9
Chapter 2: Cdc7 functions in replication fork progression in
response to MMS
hsk1
cdc7
( hsk1-1312) shows synthetic lethality with loss of Rqh1, which is RecQ helicase
in fission yeast to help rescue stalled forks (Snaith et al., 2000). Also Hsk1
Cdc7
/dfp1
Dbf4
physically interacts with the Swi1
Tof1
/ Swi3
Csm3
replication fork protection complex in
fission yeast (Matsumoto et al., 2005). In addition to its role in replication initiation,
Cdc7 could also be required for stabilizing stalled forks. To test whether Cdc7 is
required for fork stability or fork restart in response to DNA damage, we monitored the
progression of replication forks in MMS treated Cdc7 mutants cells by BrdU-IP-chip.
Since forks move too fast to be tracked easily at higher temperature, working with
temperature sensitive cdc7 mutants to track normal replication fork progression is
challenging. An alternative approach to inactivate Cdc7 kinase activity quickly and
reversibly is by ATP analogue PP1 without increasing the temperature, the analogue
sensitive cdc7as3 (L120A, V181A) mutant was constructed (Wan et al., 2006). This
Cdc7 mutant is sensitive to PP1, because the ATP binding pocket is enlarged by
introducing these two point mutations. cdc7as3 allele was introduced into the strain
YZy50 which has ars608∆ ars609∆ on chromosome VI in order to have about 75kb
replicon window from the fork initiated from ARS607 to the right arm of chromosome VI.
ARS305 was also replaced on chromosome III by inserting BrdU-Inc cassette in YZy50.
This not only enables the cell to uptake BrdU for tracing the fork movement by BrdU-IP-
chip but also creates another large replicon on chromosome III to look at fork movement.
10
mins
100
80
60
40
20
0
WT cdc7as3
For working with cdc7as3 cells, the experiment was carried out at 23˚C. Cells were
arrested in G1 phase with α factor for 4 hours and then released into S-phase without
adding ATP analogue PP1, WT and cdc7as3 followed a similar kinetics by FACS
analysis in YEPD medium (Fig 2.1), indicating cdc7as3 by itself does not affect Cdc7
function in the absence of PP1.
Figure 2.1 FACS analysis for DNA content in WT and cdc7as3 without PP1. WT and cdc7as3 were
arrested at G1 phase with alpha factor for 4 hrs at 23˚C. Then cells were synchronously released into S
phase in YEPD medium. Samples were taken at the indicated time for FACS analysis.
In another set of experiments, cells were arrested in α factor and then PP1 was added 25
minutes before release to allow cells to take up PP1 and then cells were released into S
phase in the presence of PP1 treated with or without MMS at concentration of 0.033%. In
the absence of MMS, WT cells reached 2C DNA content at ~60 mins. However cdc7as3
cells did not reach 2C DNA content until ~160 mins after release, indicating PP1 can
slow down DNA synthesis, presumably by inhibiting origin firing in cdc7as3 cells (Fig
2.2A).
min
11
MMS
Figure 2.2 FACS analysis for DNA content in WT and cdc7as3 in the presence of PP1 with and
without MMS. WT and cdc7as3 were arrested at G1 phase with alpha factor for 4 hrs at 23˚C. PP1 (25μM)
was added 25mins before release into S phase. Then cells were synchronously released in to S phase in the
presence of PP1 (25μM) in YEPD medium either without MMS as in A or with MMS at 0.033% as in B.
Samples were taken at the indicated time for FACS analysis.
The inhibition of cdc7as3 allele is not complete; otherwise cells would be totally arrested
in G
1
phase. We speculate the ATP molecule originally presented in the cells could still
bind to the mutant ATP binding pocket of Cdc7as3. And the residual Cdc7 activity is
enough for the earliest origins to fire. This is an advantage for us to study the replication
fork progression in the absence of Cdc7 activity, as we must allow some forks to be
established.
A B
WT+PP1
CDC7AS3+PP1
160
140
120
100
90
80
70
60
50
40
30
20
10
0
WT
min
s
With PP1
WT in costant mms
WT
AS in constant MMS
cdc7as3
180
160
140
120
100
80
60
40
20
0
Asyn
-
MMS
+
MMS
MMS
MMS
cdc7as3
12
In the presence of MMS, WT did not reach 2C DNA content by 180mins in S phase,
showing a dramatically slowed overall S phase progression compared to WT without
MMS (Fig 2.2B). This indicates that MMS slows down S phase in WT cells presumably
by inhibiting late origin firing and slowing down replication fork progression. In cdc7as3
cells, in the presence of MMS, the kinetic of S phase progression is similar to cdc7as3
without MMS but even slower indicating less bulk DNA synthesis in MMS in cdc7as3.
In order to follow the DNA replication fork progression, cells were aliquot during the
time course and pulse labeled with thymine analogue BrdU in the designated time
interval and then BrdU incorporated DNA was immunoprecipitated using anti-BrdU
antibody. The ongoing DNA replication in each time period was traced genome wide by
BrdU-IP-chip. In the first time interval 10-30 mins, the activities of the earliest origins
(e.g. ARS606, ARS607) and middle early origins (e.g. ARS605 and ARS603.5) were
detected clearly in WT. Similar to WT, cdc7as3 cells fired ARS606 and ARS607 as
efficiently as WT, probably through sequestering normal ATP molecule. However, the
treatment of PP1 did inhibit the initiation events from middle early origins (e.g. ARS605
and ARS603.5) as BrdU signal is barely detected at those origins in the first time point
10-30 mins (Fig 2.3). Following the forks that were established from the very early
origins ARS607 and ARS306, the fork rates were dramatically slowed down in the
presence of MMS in WT. However, the forks in cdc7as3 cells moved at a much faster
rate compared to WT throughout the time course especially in the presence of MMS. In
the last time point 55-75 mins, the fork established from ARS607 reached the coordinate
13
at 2.3x10
5
on chromosome
6 in the WT, which is ~30kb from the origin ARS607.
However, in cdc7as3, MMS failed to slow down the fork, which makes the fork rate
comparable to no MMS and reached the end of Chromosome 6.
Figure 2.3 Replication fork failed to slow down in response to MMS in cdc7as3 strain. Aliquots of the
culture were exposed to BrdU for 20-min pulses as indicated in the diagram. Cells were harvested at the
end of the each pulse for DNA isolation. Replication fork progression was monitored by tracking BrdU
incorporation using BrdU-IP-chip with Nimblegen 12x135K array slide
The fork that initiated from the early origins ARS607 failed to slow down in the presence
of MMS when the middle early origins (e.g. ARS603.5, ARS605) were inhibited as
consequence of the lacking of Cdc7 activity in the presence of the inhibitor PP1. These
results indicated that Cdc7 was not required for fork stability in response to MMS but
required for fork slowing down in MMS. This result indicated that Cdc7 is not required
ChrVI
WT w/o MMS
WT with MMS cdc7as3 with MMS
ChrIII
ARS603.5
ARS605
ARS606
ARS607 ARS304
ARS306
cdc7as3 w/o MMS
10-30
25-45
40-60
55-75
mins
14
for normal replication fork movement. Once MCMs complex has been activated through
phosphorylation by Cdc7, the helicase activity does not need to be maintained by
consistent activation by Cdc7 kinase activity. In the absence of MMS, cdc7as3 also
showed a moderately faster fork rate.
In order to confirm this result using another Cdc7 mutant, a temperature-sensitive allele,
cdc7-1 allele, was introduced to our strain background. In order to bypass the essential
function of Cdc7, we also introduced mcm5-bob1 allele into cdc7-1 mutant. The cells
were arrested in α factor for 3 hrs at 23˚C and then shifted to 32˚C for 1hr to inactivate
Cdc7 activity and then released either into YEPD or 0.033% MMS at 32˚C. As shown in
Fig 2.4, cdc7-1/mcm5-bob1 cells showed slower S phase progression in YEPD
supposedly from the deficiency of middle-early origin firing which could not be bypassed
by mcm5-bob1 allele.
15
Figure 2.4 FACS analysis for S phase progression in YEPD in WT, and cdc7-1/ mcm5-bob1.
Asynchronous WT (YZy50) and cdc7-1 (1298) and cdc7-1/mcm5-bob1 (1307) were arrested in G1 phase at
23˚C with α factor for 3h and then transferred to 32°C for 1hr before release into S phase in YEPD. And
then cells were released into YEPD at 32°C. DNA content by FACS analysis samples were collected at
time points indicated.
In the presence of MMS, cdc7-1 cells were arrested in G1 DNA content because of the
inhibition of origin firing. However cdc7-1/mcm5-bob1 strain could still enter S phase
and had similar kinetic in terms of S phase progression in MMS based on the FACS
analysis compared to WT (Fig 2.5A).
16
Figure 2.5 Replication fork failed to slow down in response to MMS in cdc7-1/mcm5-bob1 strain. A. S
phase progression in 0.033% MMS in WT, cdc7-1, cdc7-1/ mcm5-bob1. Asynchronous WT (YZy50)
and cdc7-1 (1298) and cdc7-1/mcm5-bob1 (1307) were arrested in G1 phase at 23˚C with α factor for 3h
and then transferred to 32°C for 1hr before release into S phase. Then cells were released synchronously
into YEPD medium containing 0.033% MMS at 32°C (Time=0). DNA content by FACS analysis samples
were collected at time points indicated. B. Replication fork failed to slow down in cdc7-1/mcm5-bob1
strain in response to MMS. Aliquots of the culture were exposed to BrdU for 15-min pulses as indicated
in the diagram. Cells were harvested at the end of the each pulse for DNA isolation. Replication fork
progression was monitored by tracing BrdU incorporation using BrdU-IP-chip on Nimblegen 12x135K
array slide.
A
B +MMS
17
BrdU incorporation initially occurred at the early origins ARS606 and ARS607 region
during 5-25 mins pulse period in both wild-type and cdc7-1/mcm5-bob-1 cells. Thus
replication initiation at early origins appears normal in the absence of Cdc7. However, in
cdc7-1/mcm5-bob1 cells, the mid-early origin ARS603.5 and ARS605, fired at lowered
efficiency, consistent with the cdc7as3 result. Also this is consistent with the data which
has been shown previously by 2-D gel analysis that the bypass of mcm5-bob1 for DDK
activity results in reduced intrinsic firing efficiency at some of the origins. (Hoang et al.,
2007). In cdc7-1/mcm5-bob1 cells, the fork moved at a dramatically faster rate compared
to WT at later time points (Fig 2.5 B). These results indicated that Cdc7 was not required
for fork stability in response to MMS but required for fork slowing down in MMS.
18
Chapter 3: Characterization of the relationship between
checkpoint activation and fork rate in response to MMS
3.1 Cdc7 is required for proper checkpoint activation in response to replication
stress
To test whether Cdc7 is involved in checkpoint activation in response to DNA damage
during replication, we synchronized the cells in G1 phase using alpha factor for 3 hrs at
23˚C and then shifted to 32˚C for one more hour before releasing the cells into MMS at
32˚C. We checked the checkpoint activation by detecting Rad53 phosphorylation using
western blot through the time course. Consistent with other groups’ results, we found that
checkpoint activation is compromised in response to DNA damage (Ogi et al., 2008,
Sheu et al., 2010). During the time course, we saw about equal amount of Rad53
phosphorylation by comparing the unphosphorylated and phosphorylated bands.
However in the cdc7-1/mcm5-bob1 cells, the majority of Rad53 remained
unphosphorylated (Fig 3.1)
19
Figure 3.1 Rad53 activation in Cdc7 mutant during S phase in response to DNA damageYZy50 (WT)
and (RSy1307) cdc7-1/mcm5-bob1were arrested in G1 phase at 23˚C with α factor for 3hrs and then Tm
was shifted to 32°C to inactivate Cdc7 activity for 1hr before release into S phase. Then cells were released
synchronously from G1 arrest into YEPD medium containing 0.033% MMS at 32°C (Time=0). Cell
samples were collected for TCA preps at indicated time points for looking at Rad53 activation by western
blot.
Replication forks are the sensors to activate checkpoint (Tercero et al., 2003), so less
established forks which could sense the damages could result in less checkpoint
activation. The mcm5-bob1 (P83L) mutation partially bypasses DDK requirement for
origin firing. Because deletion of Cdc7 even in mcm5-bob1 background results in about
55-75% of WT firing efficiency even at early origins (Hoang et al., 2007). The
compromised checkpoint activation in cdc7-1/mcm5-bob1 is supposedly due to the
limited number of origins fired in the mutant. In order to test whether Cdc7 really plays a
role in checkpoint activation and to exclude its indirect effect on the firing to allow
enough replication forks to sense the damages, cells were released into S phase in the
presence of 0.033% MMS at permissive temperature at 23˚C for one hour to allow
origins to be fired and the replication forks established comparable to WT. Then cells
were brought to restrictive temperature at 32˚C to go through the rest of S phase. Again
0 α factor arrested
1 WT
2 cdc 7-1 mcm5bob1
α factor arrested 3hrs at 23˚C 32˚C for 1hr release from α factor arrest into 0.033%MMS at 32˚C
20
the checkpoint activation was checked by detecting Rad53 phosphorylation using western
blot. As shown in Fig 3.2, in the first hour after releasing to S phase before Cdc7 activity
is inactivated, Rad53 could be phosphorylated at similar level in cdc7-1/mcm5-bob1
mutant compared to WT. However, the phosphorylated Rad53 started to decrease and
finally was barely detectable at the end of the time course after temperature had been
shifted to restrictive temperature at 32˚C.
Figure 3.2 Cdc7 is needed for maintaining checkpoint activation. YZy50 (WT) and RSy1307 (Cdc7-
1/mcm5-bob1) were arrested in G1 phase at 23˚C with α factor for 4hrs. Then cells were released
synchronously from G1 arrest into YEPD medium containing 0.033% MMS at 23°C (Time=0). Then Tm
was shifted to 32°C to inactivate Cdc7 activity after an hour at 23˚C. Cell samples were collected for TCA
preps at indicated time points for looking at Rad53 activation by western blot.
We observed the same result in response to HU treatment. This indicates that Cdc7 is
required for proper activation of checkpoint in response to MMS. Although we allowed
the origins to fire in the first hour at permissive temperature, presumably there are still
some late or dominate origins needed to be fired later than that in order to finish the
replication of the genome that did not replicate in the cdc7 mutant. So this phenotype of
cdc7-1/mcm5-bob1 could involve in maintaining checkpoint activation either by the
WT cdc7-1/mcm5-bob1
α factor arrested 3 hrs at 23°C release at 23°C into 0.033%MMS for 1hr raise Tm to 32°C for
2hrs
21
direct phosphorylation of Rad53 in response to DNA damage or indirectly through the
activation of the origin firing to establish the forks to sense the damage in S phase.
3.2 Pph3 deletion can partially antagonize the deregulated fork in MMS in cdc7-
1/mcm5-bob1
We observed the faster fork progression in MMS and the compromised checkpoint
activation phenotype in cdc7 mutant. It is reasonable to draw the relationship between the
failure to slow down the fork movement on damage template and the failure to activate
the checkpoint. Pph3-Psy2 is a phosphatase-complex required for Rad53
dephosphorylation and replication fork restart during recovery (O'Neill et al., 2007,
Szyjka et al., 2008). Since cdc7-1/mcm5-bob1 strain has compromised checkpoint
activation in response to MMS, we asked whether the faster fork in cdc7-1/mcm5-bob1
strain could be slowed by deleting the phosphatase Pph3 which deactivates Rad53. WT
and cdc7-1/mcm5-bob1 and cdc7-1/mcm5-bob1 pph3∆ cells were arrested in G1 phase at
23˚C for 3 hrs and then shifted to 32˚C to inactivate Cdc7 activity before release into S
phase for 1 hr; cells were released into S phase in the presence of 0.033% MMS at 32˚C
for 3hrs. We monitored fork progression in MMS by BrdU-IP-chip and found that the
faster forks in cdc7-1/mcm5-bob1 could be slowed dramatically in the cdc7-1/mcm5-bob1
pph3∆ strain (Fig 3.3). In cdc7-1/mcm5-bob1 strain, the peak of BrdU incorporation
signal for the replication fork progression has reached the end of the chromosome at
75mins after release. However in the cdc7-1/mcm5-bob1 pph3∆ strain, the faster fork in
cdc7-1/mcm5-bob1 was slowed down dramatically but still a little faster than WT.
22
Figure 3.3 Pph3 deletion can partially antagonize the deregulated faster fork in MMS in cdc7-
1/mcm5-bob1. Aliquots of the culture were exposed to BrdU for 20-min pulses as indicated in the diagram.
There is a 5 min overlap between each time point to allow the cells to take up BrdU between the time
points. Cells were harvested at the end of the each pulse for DNA isolation. Replication fork progression
was monitored by tracing BrdU incorporation using BrdU-IP-chip using Nimblegen array.
Analysis of Rad53 phosphorylation by western blot showed that the diminished
checkpoint activation indicated by the weak signal of phosphorylated Rad53 in cdc7-
1/mcm5-bob1 is partially rescued to some extent by Pph3 deletion but still not equal to
α factor arrested 3hrs at 23˚C 32˚C for 1hr release from α factor arrest into 0.033%MMS
at 32˚C
23
the WT level as shown in Fig 3.4. This suggests that there is a correlation between the
Rad53 phosphorylation state and fork progression rate in MMS.
Figure 3.4 pph3∆ can restore checkpoint activation deficiency in cdc7-1/mcm5-bob1 mutant. TCA
prep was collected at 60mins and 120 mins after release into MMS for detecting Rad53 phosphorylation
through western blot.
3.3 DDK regulation by Rad53 in response to DNA damage does not play a role in
fork progression
Dbf4, the regulatory subunit of Cdc7, is under direct regulation of checkpoint signaling
through phosphorylation by Rad53. The phosphorylation of Dbf4 has been shown to
inhibit late origin firing (Lopez-Mosqueda et al., 2010, Zegerman et al., 2010) in
response to DNA damage, whereas the function of the phosphorylation of Dbf4 on the
replication fork is unclear. It is known that DDK remains active during replication stress
(Tenca et al., 2007). To test whether the phosphorylated Dbf4 functions at forks
progressing through a damaged template, dbf4-4A allele which carries four mutations that
abolish the Rad53 phosphorylation sites was introduced into our strain background and
the replication fork progression was traced by BrdU-IP-chip. Fork progression is not
α factor arrested 3 hrs at 23°C 32°C for 1hr release from α factor at 32°C in 0.033%MMS
1 WT
2 cdc7-1/mcm5-bob1
3 pph3 Δ
4 cdc7-1 mcm5-bob1 pph3∆
24
affected in the dbf4-4A cells, which indicates that the regulation of Dbf4 by Rad53 is not
involved with fork progression rate (Fig 3.5).
Figure 3.5 Deregulation of Dbf4 by Rad53 does not affect the fork ratein MMS. A.Asynchronous WT
(YZy50) dbf4-4A (YZy55) were arrested in G1 phase at 23˚C with α factor for 4hrs and then cells were
released synchronously into YEPD medium containing 0.033% MMS at 23°C (Time=0). Samples were
taken for FACS analysis at indicated time points. B. Aliquots of the culture were exposed to BrdU for 20-
min pulses as indicated in the diagram. There is a 5 min overlap between each time point to allow the cells
to take up BrdU between the time points. Cells were harvested at the end of the each pulse for DNA
isolation. Replication fork progression was monitored by tracing BrdU incorporation using BrdU-IP-chip
with Nimblegen array.
A
B
WT dbf4
B
B
25
3.4 Other checkpoint deficiency strains cannot phenocopy the faster fork movement
in MMS
Since there appears to be a correlation between decreased checkpoint activation and the
faster fork movement, we checked other checkpoint deficient mutants to see whether they
have a faster fork movement in MMS or not. Mrc1, which is required for normal DNA
replication and checkpoint activation as a checkpoint mediator, moves with the fork as a
fork component. When replication fork stalls, Mec1 is recruited to the replication fork
where it phosphorylates Mrc1, facilitating the phosphorylation of Rad53 by Mec1 via
promotion of a stronger enzyme-substrate interaction (Chen et al., 2009). Mutation of 17
canonical Mec1 phosphorylation sites on Mrc1 (mrc1AQ) prevents Mrc1 phosphorylation
and blocks Rad53 activation, but does not alter the role of Mrc1 in DNA replication
(Osborn et al., 2003) . So we analyzed the fork rate in MMS in mrc1AQ cells in Fig 3.6.
The results show that the late origins are fired at 20-45mins in mrc1AQ cells indicating
that the late origins are not inhibited due to the compromised checkpoint activation. Fork
movement in mrc1AQ cells is a little slower than the WT strain, which means that the
compromised checkpoint activation does not necessarily leads to faster fork.
26
Figure 3.6 Fork progression in mrc1-AQ is normal in response to MMS. Aliquots of the culture were
exposed to BrdU for 20-min pulses as indicated in the diagram. There is a 5 min overlap between each time
point to allow the cells to take up BrdU between the time points. Cells were harvested at the end of the each
pulse for DNA isolation. Replication fork progression was monitored by chasing BrdU incorporation using
BrdU-IP-chip using Nimblegen array.
ARS605
ARS603.5 ARS606
ARS607
27
B
Figure. 3.7 mec1-100 has delayed checkpoint activation but normal fork rate in MMS A.
Asynchronous WT (YZy50) mec1-100 (YZy52) were arrested in G1 phase at 23˚C with α factor for 4hrs
and then cells were released synchronously into YEPD medium containing 0.033% MMS at 23°C
(Time=0). Then Cell samples were taken for FACS analysis at indicated time points. B. Aliquots of the
culture were exposed to BrdU for 20-min pulses as indicated in the diagram. There is a 5 min overlap
between each time point to allow the cells to take up BrdU between the time points. Cells were harvested at
the end of the each pulse for DNA isolation. Replication fork progression was monitored by tracing BrdU
incorporation using BrdU-IP-chip with Nimblegen array. C. Cells for TCA prep were collected at each time
point for analysis of Rad53 phosphorylation by western blot.
A
WT mec1-100
C
28
To characterize the relationship between checkpoint activation and fork progression rate
in MMS further, we also examined mec1-100 (F1179S and N1700S), which is deficient in
replication checkpoint but maintain a G2/M checkpoint (Paciotti et al., 2001). The forks
in mec1-100 in response to MMS are stable (Cobb et al., 2005) . Cells were arrested in α
factor at 23˚C for 4 hrs and released into 0.033% MMS for 3 hrs at 23˚C. Consistent with
previous finding, mec1-100 cells failed to slow bulk DNA replication in response to
MMS and reached 2C DNA content much earlier than WT cells based on the FACS data
(Fig 3.7A). However, the BrdU-IP-chip results did not show that the fork progressed at a
faster rate in MMS (Fig 3.7B); there was little or no difference in the fork rate between
WT and mec1-100. We did observe late origin firing in MMS due to the de-suppression of
the late origin firing resulting from compromised checkpoint activation in mec1-100.
However the checkpoint activation is compromised in mec1-100 at the first 2 hours in
MMS but started to increase in later S phase in the presence of MMS as shown in Fig
3.7C. However the fork did not show faster rate during the time when the checkpoint is
compromised, which indicates that down-regulation of checkpoint itself is not sufficient
for the faster fork movement in MMS.
29
Chapter 4: The choice of Post-replication repair pathways and
fork progression rate in response to MMS
To examine the genetic interaction between Cdc7 and genes in the post replication repair
(PRR) pathway, we test the sensitivity of Cdc7 and individual PRR pathway proteins and
different combination of them. As shown in Fig. 4.1, at 23˚C at which temperature cdc7-1
was not inactivated, cdc7-1/mcm5-bob1 behaved as WT strain. A TLS-defective strain
(rad30∆ rev3∆) and rad5∆ showed synergistic sensitivity to MMS treatment shown on
the 0.0001% and 0.0003% MMS plates. rad5∆ itself is very sensitive to MMS and starts
to show the sensitivity on 0.0003% whereas rad30∆ rev3∆ does not show sensitivity until
0.01%. At 32˚C, cdc7-1/mcm5-bob1 shows sensitivity at MMS concentration of 0.001%,
whereas rad30∆ rev3∆show sensitivity at 0.01%. The different sensitivities to MMS
between cdc7-1/mcm5-bob1 and rad30∆ rev3∆ further support the idea that the
sensitivity of cdc7-1/mcm5-bob1 to MMS is not simply due to its role in translesion
synthesis. Cdc7 maintains the viability of the cells through a function beyond TLS
pathway. This argument is also supported by the finding that cdc7-1/mcm5-bob1 rad30∆
rev3∆mutant had the same sensitivity as cdc7-1/mcm5-bob1. Interestingly, compared to
the rad5∆ grown at 23˚C, rad5∆ showed more sensitivity on 0.0003% MMS plate at
32°C. This indicates that the sensitivity of rad5∆ is related with the speed of the cell
cycle, presumably the speed of replication fork. The faster the forks progress at higher
temperature, the more indispensable role of Rad5 for the cells to maintain the viability.
30
At 32˚C, we started to see the synergistic defect of Rad5 mutant and Cdc7 mutant. The
synergistic defect is as strong as the synergism between Rad5 and rad30∆ rev3∆ mutant.
Figure 4.1 Genetic interactions between Cdc7 and different components of post replication repair
pathways in response to MMS. The different strains were spotted in 10-fold dilution series onto YEPD
plates containing the indicated concentrations of MMS and YEPD for control, as described in Materials and
Methods. Each plate series was incubated at 23°C or 32°C for 2 days.
4.1 Deletion of TLS polymerases has no effect on the replication fork rate in MMS
and on the faster fork in MMS in cdc7-1/mcm5-bob1
Cdc7 has been implicated to function in translesion syntheses and is required for
mutagenesis in response to DNA damage. To test whether the choice between TLS and
error-free bypass pathways might affect the replication fork rate in response to MMS, we
deleted the TLS polymerases Rev3 and Rad30 to trace the fork rate. Also, to test whether
31
the faster fork in cdc7-1/mcm5-bob1 in MMS is due to down-regulation of the translesion
synthesis bypass pathway in the cdc7-1/mcm5-bob1 strain, we deleted REV3 and RAD30
in cdc7-1/mcm5-bob1. WT (YZy50) and cdc7-1/mcm5-bob1 (1307) and rev3∆ rad30∆
(JPy3) and rev3∆ rad30∆ cdc7-1/mcm5-bob1 (YZy40) were arrested in G1 phase at
23˚C with α factor for 3hrs and then transferred to 32°C to inactivate Cdc7 activity for
1hr before being released into S phase. Then cells were released synchronously into
YEPD medium containing 0.033% MMS at 32°C. As show in Fig 4.2 A, all four strains
progressed through S phase normally, except that cdc7-1/mcm5-bob1 showed a little
slower S phase progression presumably from less efficient origin firing. In Fig. 4.2B,
BrdU-IP-chip results showed that there was no difference between WT and rev3∆ rad30∆
double mutant fork rates in the presence of MMS, both of which showed slow fork
movement. The results also showed that the deletion of TLS polymerases didn’t have any
effect on the faster fork in Cdc7 mutant. Cdc7 mutant again showed faster fork in MMS
and reached the end of chromosome 6 at 55-80 mins, while Cdc7 and TLS deletion
double mutants had the same faster fork as Cdc7 single mutant had. This indicates that
the possible role of Cdc7 in TLS does not account for the altered fork progression. The
DNA synthesis happening on the faster fork in cdc7-1/mcm5-bob1 is not dependent on
any of those translesion synthesis polymerases. Also the absence of translesion synthesis
in S phase in MMS does not lead to faster fork progression.
32
Figure 4.2 Deletion of TLS polymerases has no effect on the replication fork rate and also has no
effect on the faster fork in MMS in cdc7-1/mcm5-bob1 (A) Asynchronous WT (YZy50) and cdc7-
1/mcm5-bob1 (1307) and rev3∆ rad30∆ (JPy3) and rev3∆ rad30∆ cdc7-1/mcm5-bob1 (YZy40) were
arrested in G1 phase at 23˚C with α factor for 3hrs and then transferred to 32°C to inactivate Cdc7 activity
for 1hr before being released into S phase. Then cells were released synchronously into YEPD medium
containing 0.033% MMS at 32°C (Time=0). Then Cell samples were taken for FACS analysis at indicated
time points. (B) Aliquots of the culture were exposed to BrdU for 20-min pulses as indicated in the diagram.
Cells were harvested at the end of the each pulse for DNA isolation. Replication fork progression was
monitored by chasing BrdU incorporation using BrdU-IP-chip on Nimblegen 12x135K array slide.
A
α factor arrested 3 hrs at 23°C 32°C for 1hr release from α factor at 32°C
in 0.033%MMS
B
B
33
4.2 Rad5 is indispensable for replication fork progression in MMS in the absence of
Cdc7
Rad5 not only catalyzes PCNA poly-ubiquitylation by its E3 ligase activity, but also
possesses helicase activity which is involved in fork regression for template switch lesion
bypass pathway (Blastyak et al., 2007). Our result in Fig 4.1 and previous results showed
that cdc7∆ and rad5∆ had a synergistic sensitivity upon MMS treatment (Pessoa-Brandao
et al., 2004). We wanted to examine whether the faster fork movement in cdc7-1/mcm5-
bob1 is dependent on Rad5 mediated DNA synthesis. So we analyzed fork movement by
BrdU-IP-chip in rad5∆ and rad5∆ cdc7-1/mcm5-bob1 cells. In the rad5∆ cdc7-1/mcm5-
bob1 strain, The BrdU-IP-chip result showed origin ARS607 was fired like WT at 10-30
mins and the fork rate was indistinguishable compared to WT at 10-30 and 25-45 min.
However, the BrdU signal subsequently diminished (40-60 and 55-75 min) (Fig 4.3B).
Accordingly, DNA content analysis (Fig 4.3A) showed cells arrested with G1 DNA
content throughout the time course while the WT almost reached 2C DNA content by
3hrs. Notably, the signal did not disappear at the replicated region where the earlier time
points previously showed signal, suggesting that the replicated region probably was not
complete and ssDNA gaps were being repaired. Rad5 deletion did not slow the fork in
cdc7-1/mcm5-bob1 in MMS as PPH3 deletion did, but affected the ability of cdc7-
1/mcm5-bob1 cells to completely replicate DNA.
34
WT cdc7-1/mcm5-bob1 rae5 cdc7-1/mcm5-bob1 rad5
Figure 4.3 Rad5 is required for DNA synthesis by FACS analysis in the absence of Cdc7.
Asynchronous WT (YZy50) rad5∆ (YZy28) cdc7-1/mcm5-bob1 (1307) and cdc7-1/mcm5-bob1 rad5∆
(YZY30) and rev3∆ rad30∆ cdc7-1/mcm5-bob1 (YZy40) were arrested in G1 phase at 23˚C with α factor
for 3hrs and then transferred to 32°C to inactivate Cdc7 activity for 1hr before being released into S phase.
Cells were then released synchronously into YEPD medium containing 0.033% MMS at 32°C (Time=0).
Cell samples were taken for FACS analysis at indicated time points.
α factor arrested 3 hrs at 23°C 32°C for 1hr release from α factor at 32°C
in 0.033%MMS
35
Figure 4.4 Rad5 is required for DNA synthesis in the absence of Cdc7. Aliquots of the culture were
exposed to BrdU for 20-min pulses as indicated in the diagram. Cells were harvested at the end of the each
pulse for DNA isolation. Replication fork progression was monitored by chasing BrdU incorporation using
BrdU-IP-chip on Nimblegen 12x135K array slide.
ChrVI
36
Chapter 5: The mechanism for the synergistic defect between
Rad5 and DDK in response to MMS
We showed that the loss of DDK function is synergistic with deletion of RAD5 in
response to MMS for cell viability. The forks established from the early origins fired
normally in the absence of both DDK activity and Rad5 function but could not replicate
well indicated by the diminished BrdU signal along the time course in the BrdU-IP-chip
result. The synergistic effect causes the cells to remain arrested with unreplicated DNA
and unable to carry out S phase. However the mechanism of the synergistic defect
between Rad5 and Cdc7 is unknown.
5.1 CDC7 has synergistic defect with both the helicase and E
3
ligase function in
RAD5.
Studies of Rad5 sequence and structure have identified two functional domains: a RING
domain with E3 ubiquitin-ligase activity required for PCNA poly-ubiquitylation, and a
helicase domain with ATPase activity involved in regression of synthetic forked DNA
structures in vitro (Blastyak et al., 2007). To dissect which of the two activities is
required for the replication of damaged DNA in the absence of CDC7 activity, we
constructed strains with point mutations in the RAD5 gene that inactivate the E3
ubiquitin ligase activity (rad5-I916A) (Ulrich, 2003), or the ATPase activity rad5 GAA
(K538A/T539A) in addition to cdc7-1/mcm5-bob1. We saw that both the rad5-GAA and
rad5-I916A single mutants showed a similar degree of synergistic defect in completing
37
chromosome replication in combination with the loss of CDC7 activity in response to
MMS (Fig 5.1A). The BrdU-IP-chip results showed that either of the two single mutants
of Rad5 was similar to WT, but the BrdU signal started to decline after the second time
point 25-45mins in either the rad5E3
-
or rad5GAA combined with cdc7-1/mcm5-bob1,
which is similar to Rad5 deletion with cdc7-1/mcm5-bob1 double mutants (Fig 5.1B).
This result indicates that although the two Rad5 activities may play independent roles in
DNA damage tolerance and replication completion, CDC7 is not specifically involved in
PCNA polyubiquitination or helicase-mediated fork regression in concert with RAD5. It
is consistent with the finding that the two activities function coordinately in resolving
stalled replication forks and promoting efficient X-DNA formation derived from sister
chromatin recombination at stalled forks through forming the sister chromatin junctions
(Minca et al., 2010). However, the MMS sensitivity assay indicates that rad5 GAA
helicase single mutant was more sensitive to MMS than rad5 I916A E3 ligase single
mutant (Fig 5.2 B), which indicates that the helicase activity of Rad5 is more essential for
cells to maintain viability. Each of the single mutants of Rad5 is much more viable than
Rad5 deletion which means the two function domains supplement each other to maintain
the cell viability in response to MMS. However, the synergistic defect between cdc7-
1/mcm5-bob1 and Rad5 E3 ligase mutant is stronger than the synergistic defect between
cdc7-1/mcm5-bob1 and Rad5 helicase mutant (Fig 5.2 A), which indicates that Rad5 E3
ubiquitin ligase function is more critical when Cdc7 activity is lost.
B
38
Figure 5.1 CDC7 has synergistic defect with both the helicase and E
3
ligase function in RAD5 (A)
Asynchronous WT (YZy50) rad5E3- (YZy42) rad5GAA (YZy41) and cdc7-1/mcm5-bob1 rad5E3- (YZy44)
and cdc7-1/mcm5-bob1 rad5GAA (YZy46) were arrested in G1 phase at 23˚C with α factor for 3hrs and
shifted to 32˚C for 1hr then cells were released synchronously into YEPD medium containing 0.033%
MMS at 32°C (Time=0). Then Cell samples were taken for FACS analysis at indicated time points. (B)
Aliquots of the culture were exposed to BrdU for 20-min pulses as indicated in the diagram. Cells were
harvested at the end of the each pulse for DNA isolation. Replication fork progression was monitored by
chasing BrdU incorporation using BrdU-IP-chip on Nimblegen 12x135K array slide.
A WT Rad5 I916A Rad5 GAA cdc7-1/mcm5-bob1 cdc7-1/mcm5-bob1
Rad5 I916A Rad5 GAA
cdc7-1/mcm5-bob1
Rad5 I916A (E
3
-) Rad5 GAA (helicase-) min
B
39
Figure 5.2 Genetic interaction between Cdc7 and Rad5 GAA (helicase-) and Rad5I916A (E3 ligase-)
The different strains were spotted in 10-fold dilution series starting from OD 0.5 onto YEPD plates
containing the indicated concentrations of MMS and YEPD for control, as described in Materials and
Methods. Each plate series was incubated at 32°C for 2 days.
5.2 Elimination of TLS and Rad5 pathwaysshowed no synergistic defects in
replication fork progression in response to MMS
DDK has been shown to play a role in DNA damage tolerance pathway through
promoting TLS DNA damage bypass pathway (Pessoa-Brandao et al., 2004). If the
synergistic effect of replication defect in MMS between cdc7-1 and rad5∆ were because
of the abolishment of both the TLS DNA damage tolerance pathway and Rad5 dependent
template switch DNA damage tolerance pathway, the combination of the blockage in the
TLS pathway by deletion of the translesion DNA polymerases and the deletion of Rad5
would phenocopy synergistic defect in cdc7-1/mcm5-bob1 rad5∆. So we looked at the
fork progression in the triple mutant rev3∆ rad30∆ rad5∆. It showed that in the absence
of either Rad5 or TLS polymerases, the fork moved at similar rate as WT. In the rev3∆
40
rad30∆ rad5∆ strain, the fork also moved at similar rate as WT in MMS (Fig 5.3). That
indicates that the fork progression rate is not dependent on TLS pathway and Rad5
mediated template switch DDT pathway, since deletion of both pathways does not have
an effect on the fork progression rate. This result also indicates that both error free Rad5-
mediated template switch and error prone TLS pathways are not essential for fork
progression in MMS, which is consistent with the finding that the RAD6 DNA damage
tolerance pathway can be uncoupled from the replication fork and is functional beyond S
phase. (Daigaku et al., 2010, Karras et al., 2010). But it doesn’t exclude the possibility
that there is a difference in terms of the integrity of the newly synthesized DNA behind
the fork, presumably more ssDNA gaps behind the forks in the mutants. After all, the
triple mutant which has both error prone and error free bypass pathways knocked out was
very sensitive to MMS, which is almost comparable to cdc7-1/mcm5-bob1 rad5∆ mutant
(Fig 4.1). Nevertheless, the fork rate is indistinguishable compared to WT. Thus, the fork
rate of the cells is not necessarily an indication for sensitivity to MMS.
41
Figure 5.3 TLS∆ and Rad5∆ show no synergistic defects in response to MMS. (A) Asynchronous WT
(YZy50) rad5∆ (YZy28) rev3∆ rad30∆ (JPy4) and rev3∆ rad30∆ rad5∆ (YZY53) were arrested in G1
phase at 23˚C with α factor for 4hrs and then cells were released synchronously into YEPD medium
containing 0.033% MMS at 23°C (Time=0). Cell samples were taken for FACS analysis at indicated time
points. (B) Aliquots of the culture were exposed to BrdU for 20-min pulses as indicated in the diagram.
Cells were harvested at the end of the each pulse for DNA isolation. Replication fork progression was
monitored by tracing BrdU incorporation using BrdU-IP-chip on Nimblegen 12x135K array slide.
A
B
42
5.3 cdc7-1/mcm5-bob1 rad5∆ double mutant cannot recover after MMS exposure
The synergistic defect in replication in MMS in cdc7-1/bob-1 rad5∆ is not simply
because of the additive defect in cdc7-1 mediated TLS and the defect in Rad5-mediated
lesion bypass, but due to some other function of cdc7. Cdc7 mutant has less checkpoint
activation as shown in Fig 3.1 and Fig 3.2. Similar to other group’s finding (Karras et al.,
2010), we observed robust checkpoint activation by examining Rad53 phosphorylation in
the rad5∆ cells in constant MMS exposure in S phase (Fig 5.4A). This activation is
compromised in cdc7-1/mcm5-bob1 rad5∆ cells (Fig 5.4B), so it is possible that the
synergistic defect is related to compromised checkpoint activation in cdc7-1/mcm5-bob1.
If replication forks collapse in the cdc7-1/mcm5-bob1 rad5∆ cells in MMS, these cells
will fail to recover from MMS exposure as observed with checkpoint mutants e.g.:
rad53∆. But if the forks stall stably in the cdc7-1/mcm5-bob1 rad5∆ cells, replication
would be expected to resume during recovery from MMS exposure. To determine
whether the fork had collapsed or stalled stably in the Cdc7 and Rad5 double mutant, WT,
cdc7-1/mcm5-bob1, rad5∆, and cdc7-1/mcm5-bob1 rad5∆ were arrested in G1-phase,
using α-factor, for 3 hours at 23°C and then 32°C for an hour. These cells were released
into 0.033% MMS for one hour at 32°C. MMS was quenched and washed out and were
resuspended in YEPD for recovery at 32°C. As shown in Fig 5.5, the cdc7-1/mcm5-bob1
rad5∆ double mutant cells did not recover after exposure to MMS for 1hr suggesting that
the forks collapse irreversibly and cannot resume replication in the absence of Rad5 and
Cdc7 activity. This phenotype might be the result of the compromised checkpoint
43
activation in the double mutant as shown in Fig 5.4, since checkpoint is highly activated
in RAD5 deletion strain. The question is whether rad5∆ is synergistic with any other
checkpoint mutant.
A
0
1 WT 2 cdc7-1/mcm5-bob1 3 rad5∆ 4 cdc7-1/mcm5-bob1 rad5∆
Figure 5.4 Rad53 activation in rad5∆ and in cdc7-1/mcm5-bob1 rad5∆ double mutant. Asynchronous
WT (YZy50) rad5∆ (YZy28) cdc7-1/mcm5-bob1 (1307) and cdc7-1/mcm5-bob1 rad5∆ (YZY30) and
rev3∆ rad30∆ cdc7-1/mcm5-bob1 (YZy40) were arrested in G1 phase at 23˚C with α factor for 3hrs and
then transferred to 32°C to inactivate Cdc7 activity for 1hr before being released into S phase. Then cells
were released synchronously into YEPD medium containing 0.033% MMS at 32°C (Time=0). TCA prep
was collected at indicated timepoints after release into MMS for detecting Rad53 phosphorylation through
western blot.
0 0 30 30 60 60 90 90 120 120 150 150 180 180 mins
1 WT 2 rad5∆
0 1 2 3 4 1 2 3 4 1 2 3 4
1 2 1 2 1 2 1 2 1 2 1 2 1 2
60 mins 120mins 180mins
B
44
α-factor arrest 3hrs at 23C -> 32C for 1hr -> MMS 0.033% for 1hr -> recover in YEPD for 2hrs
Figure 5.5 cdc7-1/mcm5-bob1 rad5∆ double mutant could not recover after 1 hr 0.033% MMS
exposure. Asynchronous WT (YZy50) rad5∆ (YZy28) cdc7-1/mcm5-bob1 (1307) and cdc7-1/mcm5-bob1
rad5∆ (YZY30) and rev3∆ rad30∆ cdc7-1/mcm5-bob1 (YZy40) were arrested in G1 phase at 23˚C with α
factor for 3hrs and then transferred to 32°C to inactivate Cdc7 activity for 1hr before being released into S
phase. Then cells were released synchronously into YEPD medium containing 0.033% MMS at 32°C
(Time=0). 0.5% sodium thiosulfate was added to quench the MMS before resuspending the cells in YEPD.
Cell samples were taken for FACS analysis at indicated time points.
5.4 rad5∆ is not synergistic with mec1-100 to 0.033%MMS
To test whether mec1-100 is also synergistic with Rad5 deletion, Rad5 was deleted in
mec1-100 strain. Using mec1-100, a phenotypically similar strain to the cdc7-1 in terms
of checkpoint activation but not maintenance of fork stability (Cobb et al., 2005), we
wanted to test whether the synergistic defect caused by the rad5Δ in the cdc7-1/mcm5-
bob1 was a result of a deficiency in checkpoint activation of the Cdc7 mutant. As shown
in Fig 5.6, the Rad5 deletion is not synergistic with mec1-100 and mec1-100 is almost the
same sensitivity as WT on MMS plate.
W
T
cdc7-1/mcm5-bob1 cdc7-1/mcm5-bob1 rad5 ∆ rad5∆
140
120
100
80
60
40
20
60
(0)
40
20
0
0.033%
MMS
32°C
YEPD
WT
45
Figure 5.6 rad5∆ did not show synergistic defect with mec1-100. YZy50 (WT) YZy28 (rad5∆) YZy52
(mec1-100) and YZy59 (mec1-100 rad5∆) were spotted with 10μl in 10-fold dilution series starting from
OD 0.5 onto YEPD plates containing the indicated concentrations of MMS and YEPD for control, as
described in Materials and Methods. Each plate series was incubated at 32°C for 2 days.
Also as shown in Fig 5.7, we found that mec1-100 rad5∆ cells, unlike rad5∆ cdc7-
1/mcm5-bob1 cells, which showed arrested with G1 DNA content in response to MMS
were able to progress through S phase achieve G2 DNA content by FACS in the presence
of MMS. This suggested that there was no synergistic relationship between mec1-100 and
rad5Δ, and the cdc7-1/mcm-bob1 rad5Δ inability to complete replication was not due to
the compromised checkpoint activation.
46
180
160
140
120
100
80
60
40
20
0
Figure 5.7 Rad5 deletion doesn’t show synergistic defect with mec1-100 to 0.033% MMS. YZy50
(WT) and YZy59 (mec1-100 rad5∆) cells were arrested in G1 with α factor for 4 hours at 23°C and
released into S phase in YEPD with 0.033% constant MMS. Time points for FACS analysis were taken
every 20 minutes for 3 hours.
WT
mec1-100 rad5∆
47
Chapter 6: Detecting ssDNA gaps behind the replication forks
in Cdc7 mutant
The replication forks go fast in MMS in the cdc7-1/mcm5-bob1 mutant. However, it is
not clear whether the DNA synthesis is continuous. To detect the possible existence of
ssDNA gaps behind the fork, RFA1, which encodes the largest subunit of the ssDNA
binding protein RPA, was MYC-tagged in the WT and cdc7-1/mcm5-bob1 strains.
Figure 6.1 RPA-ChIP-Chip in WT and cdc7-1/mcm5-bob1 JPy8 (WT) and JPy9 (cdc7-1/mcm5-bob1 )
were arrested in Asynchronous in G1 phase at 23˚C with α factor for 3h and then transferred to 32°C for
1hr before release into S phase. Then cells were released synchronously into YEPD medium containing
0.033% MMS at 32°C (Time=0). Cells were collected for RPA ChIP.
RPA binding was detected along chromosome 6 both in WT and cdc7-1/mcm5-bob1 at
similar signal intensity (Fig 6.1). Since the cdc7-1/mcm5-bob1 strain had faster fork
movement in MMS, RPA binding also migrated further along the chromosome 6. This
48
indicates that the DNA synthesis occurred through bona-fide replication fork activity
with DNA unwinding at the replication fork. However, due to the resolution limitation of
the RPA-ChIP-chip method, we cannot conclude that more ssDNA gaps were left behind
the fork in the cdc7-1/mcm5-bob1 cells. DNA combing would be a better way to try and
definitely give us information on that.
To detect the chromosomal DNA integrity, PFGE was used to compare the kinetics for
the completion of DNA synthesis in the WT and cdc7-1/mcm5-bob1 strains after G1
arrest and release into 0.033% MMS for 1hr. As shown in Fig 6.2B, in cdc7-1/mcm5-
bob1, the intensity of the chromosomal DNA bands were reduced compared to WT. This
suggests that cdc7-1/mcm5-bob1 cells did not complete chromosomal replication despite
the rapid fork progression. The completeness for the replication of the 16 chromosomes’
DNA was delayed either by the sloppy DNA replication done in the absence of Cdc7
activity or by the deficiency of late or dominant origins’ firing in the cdc7-1/mcm5-bob1
strain. This result is also supported by the FACS analysis in Fig 6.2A, indicating that
cells were arrested in G2/M phase for a long time before entering the next cell cycle.
49
A
Figure 6.2 cdc7-1/mcm5-bob1 is defective in recovery after MMS exposure by PFGE. A.
Asynchronous WT (YZy50) and cdc7-1/mcm5-bob1 (1307) were arrested in G1 phase at 23˚C with α factor
for 3hrs and then transferred to 32°C to inactivate Cdc7 activity for 1hr before being released into S phase.
Then cells were released synchronously into YEPD medium containing 0.033% MMS at 32°C (Time=0)
for 1hr. 0.5% sodium thiosulfate was added to quench the MMS before releasing the cells into YEPD. Cell
samples were taken for FACS analysis at indicated time points. B. Samples were taken at indicated time
point for PFGE analysis as in material and methods.
20hrs
6hrs
5hrs
280’
260’
240’
200’
180’
160’
140’
120’
100’
80’
60’
40’
60’ (0’)
40’
20’
0’
MMS
YEPD
WT
T
cdc7-1/mcm5-bob1
B
50
Chapter 7: ORC depletion deregulates replication fork
progression in MMS
ORC is a six subunit protein complex that binds to origins and is requried for origin firing
(Bell et al., 2002). orc1-161 is a temperature-senstive allele of Orc1which results in tight
arrest with unreplicated DNA content when incubated at the restrictive temperateure of
37˚C, due to the defect in pre-RC assembly and origin firing (Gibsonl et al., 2006).
However, when orc1-161 was grown at the semi-permissive temperature of 32˚C, origin
firing was similar to cdc7-1/mcm5-bob1. To test the possiblity that reduced origin firing
leads to lower checkpoint activation that results in faster fork movement, WT and orc1-
161 were arrested in α factor at 23˚C for 3 hrs and shifted to semi-permissive
temperature 32˚C before release into 0.033% MMS at 32˚C for 3hrs. FACS analysis for
DNA content shows that orc1-161 has a much slower S phase progression due to the
compromised origin firing (Fig 7.1A). Similar to cdc7-1/mcm5-bob1 mutant, orc1-161
also showed faster fork movement in MMS, suggesting a relationship between the
number of origin firing and fork progression rate. As fewer forks are established, there
should be less checkpoint activationand less competition for the replication proteins and
nucleotides for DNA synthesis. Whether there is also less checkpoint activated in MMS
in orc1-161 strain is still under investigation.
51
Figure 7.1 orc1-161 also showed accelerated fork rate in MMS. Asynchronous T2y41 (WT) and T2y42
(orc1-161) were arrested in G1 phase at 23˚C with α factor for 3hrs and then transferred to 32°C to
inactivate Cdc7 activity for 1hr before being released into S phase. Then cells were released synchronously
into YEPD medium containing 0.033% MMS at 32°C (Time=0). Then Cell samples were taken for FACS
analysis at indicated time points. (B) Aliquots of the culture were exposed to BrdU for 20-min pulses as
indicated in the diagram. Cells were harvested at the end of the each pulse for DNA isolation. Replication
fork progression was monitored by chasing BrdU incorporation using BrdU-IP-chip on Nimblegen
12x135K array slide.
A
B
52
Chapter 8: DDK regulation of MCM2 in response to MMS
In Saccharomyces cerevisiae, members of the Mcm2-7 family undergo cell cycle-specific
phosphorylation. The phosphorylation of Mcm2-7 complex by Cdc7-Dbf4 (DDK) kinase
complex has been shown to activate its helicase activity and initiate the unwinding of
DNA at the origins (Labib, 2010). And there are several Mcm2-7 subunits mutants that
can bypass the essentiality of DDK for the initiation step of origin firing (Sclafani et al.,
2002, Sheu et al., 2010). But more and more evidence has shown that the helicase activity
of Mcm2-7 helicase complex is also under the control of the checkpoint under replication
stress (Ishimi et al., 2004, Yoo et al., 2004). It has been shown that phosphorylation of
MCM2 at S164 and S170 which are DDK phosphorylation sites are not essential for
viability (Bruck et al., 2009, Stead et al., 2011). However, a strain with mutations of
Mcm2 that abolish phosphorylation of those two residues (mcm2
AA
) is sensitive to MMS
and caffeine, and accumulates more RPA foci than WT in response to MMS. The
accumulation of RPA suggests the presence of ssDNA behind the fork. In contrast,
phosphomimetic mutations on those two residues S164E and S170E (mcm2
EE
) suppress
the MMS and caffeine sensitivity caused by deficiency of DDK function. More
importantly, DDK-dependent modification of Mcm2 at S164 and S170 inhibits DNA
unwinding by Mcm2-7 in vitro (Stead et al., 2011). The defect in DNA unwinding by
mcm2EE results from enhanced DNA binding.
53
ChrIII ChrVI mins
ARS304 ARS306 ARS603.5 ARS605 ARS606 ARS607
Figure 8.1 The mcm2
AA
mutant showed normal replication fork progression in MMS. Asynchronous
WT (YZy50) mcm2
AA
(YZy64) were arrested in G1 phase at 23˚C with α factor for 4hrs and then cells were
released synchronously into YEPD medium containing 0.033% MMS at 23°C (Time=0). Aliquots of the
culture were exposed to BrdU for 20-min pulses as indicated in the diagram. There is a 5 mins overlapping
between each time point to allow the cells to take up BrdU between the time points. Cells were harvested at
the end of the each pulse for DNA isolation. Replication fork progression was monitored by tracing BrdU
incorporation using BrdU-IP-chip using Nimblegen array.
We have found that the fork failed to slow down in response to MMS in the absence of
Cdc7 activity. We hypothesized that loss of Cdc7 function to regulate helicase activity
through phosphorylation of Mcm2 causes the faster fork movement of CDC7 mutant cells.
This idea is also consistent with the fact that the phosphorylation of Mcm2 and the
phosphomimic mcm2
EE
mutant both reduce the helicase activity in vitro (Stead et al.,
2011). To track the fork progression in MMS in mcm2
AA
cells in vivo using BrdU-IP-chip
55-75
10-30
25-45
40-60
54
assay, we introduced the mcm2
AA
allele into our strain background to test whether it
would phenocopy the abnormal fork rate we observed in Cdc7 mutants. However, the
fork rate in mcm2
AA
cells is no different than WT along both chromosomes 3 and 6 in Fig
8.1. This result indicates that defective phosphorylation of Mcm2 by DDK is not
sufficient for the observed deregulation of fork progression in CDC7 mutant cells.
55
Chapter 9: Material and Methods
9.1 Plasmid and strain constructions
All strains are derived from W303 background. Strains are described in Table 1. Gene
knockouts were constructed by PCR-based methods (Guldener et al., 1996, Longtine et
al., 1998). To construct cdc7as3 strain, the 1.6-kb HindIII EcoRI fragment of cdc7as3
containing the kinase-inactivating mutations L120A, V181A was isolated from
pMW515-cdc7as3 (L120A, V181A) (Ulrich, 2003) and sub cloned into EcoRI–HindIII-
digested pRS306. The cloned plasmid pRS306-cdc7as3 was linearized with EcoRI and
integrated at its native genome locus through pop-in pop out. The mutant was screened
by sequencing. To construct Rad5 helicase mutant strain and Rad5 E3 ubiqutin ligase
mutant strain, the plasmid YIp211-pRad5-rad5-GAA (helicase mutant)(Chen et al., 2005)
and YIp211-pRad5-rad5-I916A (E3 ligase mutant) (Ulrich, 2003) were digested with
HpaI and integrated into the genome through pop in and pop out method. To construct the
mec1-100 strain, the fragment containing the mec1-100 allele was isolated from the
plasmid pML 258.51 (Paciotti et al., 2001) by SacI and SpeI and subcloned into pRS406.
The cloned plasmid pRS406-mec1-100 was cut by BstEII and intergrated into the native
genome locus through Pop-in and pop-out. To construct the dbf4-4a strain, the plasmid
1819 pRS303-dbf4-4a (Zegerman et al., 2010)containing the dbf4-4a allele was cut by
ClaI and XbaI and cloned into pRS406. The cloned plasmid was cut by SwaI and
intergrated into the genome through pop-in and pop out. To construct the mcm2
AA
strain,
56
plasmid pMD367 (Stead et al., 2011) was digested by BsrGI and transformed to WT
strain and selected for the mcm2AA mutant after FOA plating through pop-in and pop out.
9.2 DNA content analysis
Prepare 1.5ml eppendorf tubes containing 1ml ice-cold H
2
O and sit them on ice. Harvest
0.5 mL cells (for each analysis 5 x 10
6
to 1 x 10
7
cells are required) into the tube
containing ice-cold H
2
O. Spin down cells briefly at full speed in micro centrifuge (only 2
seconds after reaching maximum speed), remove supernatant by the aspirator. Add in 300
µL ice-cold H
2
O and resuspend the pellet completely by vortex. Add 700µl of 95%
EtOH to fix cells. Incubate at 4° C for at least 2 hours (up to a few days is fine). Spin
down cells as above and resuspend the cell pellet in 1mL of 50 mM NaCitrate (pH = 7.4).
Spin down cells as above and resuspend in 0.5 ml of 50 mM NaCitrate containing 0.20
mg/ml (final concentration) DNAse-free RNase A. Incubate cells for 2~3 hours at 50° C
water bath with occasional inversion of the tubes. Brief centrifuge after the incubation
and add 12.5 µL of 20mg/ml Proteinase K, mix and continue the incubation for 1~2 hours
at 50°C. Add 0.5 mL of 50 mM NaCitrate containing 1 M Sytox Green (from 5mM
5000X stock) into the tube. Transfer to FACS tubes, protect from light and incubate 30
minutes at room temp or overnight at 4°C. Analyze on FACScan machine.
9.3 BrdU-IP-chip
The BrdU incorporation cells were grown in the presence of 800ug/mL BrdU (sigma).
25ml culture of 0.8OD cells were collected into ice-code 50ml conical tube with 250ml
57
10% Sodium Azide. The cell pellet was washed by ice-cold 1xTBS and snap frozen by
ethanol with dry ice. Then DNA was extracted by glass bead beating, sheared by
sonication to average size of ~500bps, and isolated by phenol/chloroform extraction and
ethanol precipitation. DNA was treated with RNAaseA and proteinase K and further
purified on Qiaquick PCR purification spin-column (Qiagen). About 1μg DNA was
combined with 20ug of sheared salmon sperm single stranded DNA in 1XPBS buffer and
was denatured at 98˚C for 10 mins and snap cool on ice for 10 mins and incubated with
anti-BrdU antibody ( invitrogen 1:500) at 4˚C for at least 2hrs to overnight followed with
protein G Dynabeads from Invitrogen. Half of the immunopercipitated DNA was
amplified by WGA kit from sigma (O'Geen et al., 2006). The reference “Total DNA”
sample was isolated from G1 arrested culture. And 10ng was used to for the WGA
amplification. 1μg of amplified Ip DNA was labeled with Cy5 and 1μg Total DNA was
labeled with Cy3 using klenow following the protocol from Nimblegene. 1μg Cy5
labeled immune-precipitated DNA and 1 μg Cy3 labeled Total reference DNA were
combined and dried down using speed-vacuum. The hybridization was performed
following the user guide from Nimblegen. Microarray analysis was performed by using
Matlab using the script generated especially for highly enriched IP-chip dataset (Knott et
al., 2009).
9.4 Viability assay
Cells were growing in corresponding DOB medium overnight to mid exponential phase
at about 0.5 OD. Then cells were spun down and resuspended into YEPD at the same OD
58
(exactly OD0.5). 10μl of culture were spotted on the YEPD plate and YEPD plates
containing different concentrations of MMS as indicated. For each strain, four 1:10 serial
dilutions of the original culture were spotted on the plate. Plates were grown at indicated
temperatures for 2 days at 32˚C and 3 days at 23˚C.
9.5 Pulse Field Gel Electrophoresis
10ml of OD 0.8 cells were collected into pre-chilled conical tube containing 100ul 1%
NaN
3
(final 0.01% NaN
3
). The pellet was washed with LET buffer (0.05M EDTA and
0.01M Tris Cl) twice. The pellet was resuspended in LET buffer to make the cell
concentration to 1.5 X 10
9
/ml. Then 100ul of cells were combined with 100ul 1.2% LMP
agarose (dissolved in 0.125M EDTA) embedded with Zymolyase (0.1mg/ml) into the
mold (Bio-Rad) to make gel plug. After the gel plug solidified, the cell plugs containing
Zymolyase were incubated in 0.5M EDTA with 7.5% β-mercaptoethanol at 37˚C for
overnight. Next day, the gel plugs were washed with LET buffer once and then
transferred into the solution containing 1% sarkosyl 0.5M EDTA pH9.5 containing
proteinase K (1mg/ml) and digested overnight at 50˚C. Then the cell plugs were washed
three times (15mins each time) with LET buffer and three times with 0.5XTBE each time
10mins. Then the plugs were run at a Bio-Rad PFGE apparatus at 24 hrs. The gel were
stained with EtBr and visualized under UV light.
59
9.6 Western blot
Analysis of Rad53 Protein extracts were prepared by trichloroacetic acid precipitation
(TCA) as described previously (Foiani et al., 1994). The protein extracts were separated
by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), using 7.8%
polyacrylamide (77:1) gels. Goat Rad53 polyclone antibody (santa cruz YC-19) was used
at 1:1,000 and secondary antibody bovine anti-goat IgG-HRP (sc-2350) at 1:10,000 in
conjunction with Super Signal west Femto (Pierce) for quantitative chemiluminescent
detection of protein with a ChemiDoc XRS 170-8070 (Bio-Rad) and Quantity One
Analysis software (Bio-Rad).
60
Chapter 10: Discussion
10.1 Replication fork progression is regulated in response to DNA damage
It has been shown that checkpoint controls the late origin firing, however, it was
concluded that the fork elongation rate in MMS is checkpoint independent using a
density hybrid shift approach (Tercero et al., 2001). In Schizosaccharomyces pombe, the
hsk1-89 recovered from HU block and released into MMS had a faster S phase compared
to WT, indicating Hsk1
Cdc7
is required for the stability of the arrested replication forks
(Matsumoto et al., 2005). hsk1
ts
cells were also unable to properly delay S-phase
progression in the presence of DNA alkylating agent, such as MMS (Matsumoto et al.,
2005). Szyjka and colleagues showed that replication forks are slow to restart when the
deactivation of Rad53 is compromised in pph3∆ (Szyjka et al., 2008). This actively
regulated slowing of the fork in response to DNA damage, or elongation checkpoint, may
facilitate DNA repair by delaying replication of the damaged template and provide time
for repair.
Here, we provide further evidence that replication fork rate is a regulated process by
showing that in the absence of Cdc7 activity forks failed to slow in response to DNA
damage. Defective Cdc7 function resulted in reduced origin firing and consequently
reduced checkpoint activation and fewer forks established to compete for limiting
replication proteins and metabolites. This phenotype is shared by orc1-161 which is
involved in pre-RC assembly for origin firing. The rapid fork progression on MMS-
61
damaged DNA demonstrates that the MMS-induced DNA lesions do not physically
impede the fork and indicates the slowing of replication fork is an active process.
Furthermore, the results suggest that replication fork slowing by MMS damage on the
template DNA is related to the status of origin firing. Reduced origin firing generates
fewer replication forks and hence, less checkpoint activation and less competition for
limiting replication fork factors, resulting in. faster fork movement. Thus, the faster fork
movement depends on two conditions: less origin firing than normal, and less checkpoint
activation. Less checkpoint activation alone did not lead to faster fork movement, since
none of the checkpoint mutants, including mec1-100 and rad53∆, show faster fork
movement in MMS. Possibly the material or some replication fork components for the
DNA synthesis are limiting in the cell, since late origin along with early origin could all
be fired in checkpoint mutant due to the de-suppression of the late origin firing. It would
be interesting to test whether we could observe the faster fork movement in mec1-100
strain if we could limit the origin firing. We also could test whether we still could see the
faster fork movement if we introduce the SLD3-m25 dbf4-m25, which will bypass the
suppression of the late origin by the checkpoint, to cdc7-1/mcm5-bob1 mutant to allow all
the origin fire in the early S phase in MMS and then inactivate Cdc7 activity.
In addition, more evidence has shown that homologous recombination (HR) components
actually play a role in fork rate. In hamster cell lines defective in HR either by over-
expression of RAD51 dominate-negative form, or by defect in the RAD51 paralogue
XRCC2 or the breast tumor suppressor BRCA2. They exhibited a similar reduction in the
62
rate of replication-fork progression and an elevated fork density. (Daboussi et al., 2008).
In the absence of Rad52 and Rad18, replication fork progression is impeded as analyzed
by density-shift assay (Vazquez et al., 2008). Moreover, in unperturbed S phase, sgs1∆
has been shown to have a faster fork movement (Versini et al., 2003), which is not
dependent on Rad53 but because of the de-suppression of illegitimate recombination
occurring at stalled replication forks. This also indicates that the fork movement rate is
related to the ongoing fork actions at the fork where it encounters damage on the
templates. Recently, it has been shown that DDK phosphorylates checkpoint clamp
component Rad9 and promotes its release from damaged chromatin to facilitate
homologous recombination (Furuya et al., 2010). It would be interesting to test whether
CDC7 mutants have any interaction with recombination proteins in terms of the DNA
synthesis at the fork. At least in S. pombe, hsk1 cells suffer increased rates of mitotic
recombination and require recombination proteins for survival.
Recently it was found that dNTP pools affect fork progression rate. The fork progression
could be accelerated ~3-fold increase in the presence of MMS by over-expression of
RNR, an enzyme that regulates a rate-limiting step in dNTP biosynthesis (Poli et al.,
2011). This is consistent with our finding that less forks established lead to faster fork
movement in cdc7-1/mcm5-bob1 and orc1-161.
Finally, in HeLa cells, Cdc7 depletion in HU- and etoposide-treated cells affects the
phosphorylation status of Mcm2 and Mcm4 proteins. (Tenca et al., 2007).
63
More directly, the phosphorylation of Mcm2 by DDK slows down the helicase activity in
vitro by increasing the DNA binding activity (Stead et al., 2011). It is tempting to
hypothesize that the modification of Mcm2 by DDK slows down the helicase activity.
However, the finding that the mcm2AA mutant did not exhibit faster fork movement
through damaged DNA indicates that the abrogation of DDK phosphorylation on Mcm2
is not sufficient for the faster fork movement we observed in cdc7 mutant to occur. It
might be the limiting replication factors that prevent more rapid fork movement in
mcm2
AA
mutant.
10.2 Loss of Cdc7 function creates a critical dependence on Rad5 to complete DNA
replication
Cdc7’s critical role in replication initiation has been extensively studied and much more
appreciated than its other functions. However, more and more evidence indicates that
Cdc7 is required after replication initiation (Masai et al., 1995, Takeda et al., 1999, Dolan
et al., 2010). Our results also showed that cdc7-1/mcm5-bob1 was sensitive to chronic
MMS exposure above 0.001% (Fig 4.1). cdc7-1/mcm5-bob1 lost viability after acute
exposure to 0.033% MMS during S phase after G1 arrest release. This indicates that Cdc7
is required for DNA damage response after early S phase.
64
The requirement of Cdc7 function in response to DNA damage could come from several
aspects. First of all, Cdc7 might be involved in replication fork stability. In fission yeast,
hsk1
cdc7
(hsk1-1312) shows synthetic lethal with loss of Rqh1, which is RecQ helicase in
fission yeast to help rescue stalled forks (Snaith et al., 2000). Also Hsk1
Cdc7
/dfp1
Dbf4
physically interacts with the Swi1
Tof1
/ Swi3
Csm3
replication fork pausing complex in
fission yeast (Matsumoto et al., 2005). It is reasonable to hypothesize that the sensitivity
of cdc7 to MMS is due to the collapse of the forks in MMS as seen in checkpoint mutants
eg. rad53∆ or mec1∆. However, our results indicate that the stability of the forks in cdc7
mutant in response to MMS was not affected based on the FACS analysis which showed
that cells still could recover and go through S phase after one hour of MMS or HU
treatment. This means the forks were stable and can resume the replication once the
damage was gone, otherwise the fork would be irreversibly collapsed and could not
resume the function (Tercero et al., 2001). Replication fork collapses in mutants lacking
the Rad53/Cds1 checkpoint kinase in response to HU treatment and cannot resume
replication when HU is removed to recover (Lopes et al., 2001). In contrast, in cdc7-
1/mcm5-bob1 mutant replication forks actually move at a faster rate compared to WT.
This indicates that Cdc7 is not required for fork stability and the replication fork does not
collapse.
The strong BrdU signal in BrdU-IP-chip experiment in addition to RPA chromatin
immunoprecipitation signal along the chromosome where the forks went through all
65
indicate that the DNA synthesis in Cdc7 mutant is carried out by bona fide replication
fork, or at least is coupled to a particular kind of fork activity happening at the replication
forks, which requires both Rad5 helicase activity for fork regression and Rad5 ubiquitin
ligase activity for poly-ubiquitylation of PCNA, since the DNA synthesis in cdc7 mutant
in response to MMS is totally dependent on Rad5 functions. The slight slower fork rate in
rad5∆ at 32˚C also indicates that Rad5 functions directly on the fork. This is consistent
with the finding that both of the two activities of Rad5 are required for sister chromatid
recombination to bypass DNA damage at stalled replication forks in response to
Adozelesin, which also causes stalling of the fork (Minca et al., 2010). Since the
completion of DNA synthesis in MMS in cdc7 mutant is dependent on Rad5, it indicates
that Cdc7 normally inhibits the Rad5 mediated fork restart or channels the stalled
replication fork away from the Rad5 medicated fork restart pathway which could happen
faster than any other fork restart pathways eg. TLS. Or in the absence of Cdc7, the
replication forks which encountering DNA damage on the templates create more
abnormal structures which need Rad5 activity to resolve. It would be interesting to test
whether the X-structure on 2-D gel analysis which indicates the sister chromatin
recombination structure in response to MMS will have any increase in the cdc7-1/bob1
mutant. An increase of the X-structure means that cells would have to take the
homologous recombination pathway to bypass the lesion in the absence of Cdc7 activity.
Our experiment showed that deletion of TLS error prone polymerase did not change the
fork progression rate and did not affect the faster fork movement in cdc7-1/mcm5-bob1
suggested that TLS pathway could be uncoupled from the fork activity and it is not
66
responsible for the DNA synthesis in cdc7-1/mcm5-bob1. This is consistent with the
finding that RAD6 DNA damage tolerance pathway can operate uncoupled from the
replication fork and is functional in G2/M phase. (Daigaku et al., 2010, Karras et al.,
2010). TLS is error prone and is required for DNA damage induced mutagenesis. In cdc7
mutants, stalled forks might restart using Rad5-mediated fork regression restart pathway
which is error free, thus cdc7 mutants are generally hypo-mutagenic. Consistent with our
findings, it has been found recently that hsk1
cdc7
-1312 is synthetic lethal with rhp51∆
Rad51
and that there is increase in recombination centers indicated by Rad52 foci in hsk1
cdc7
-
1312 even under permissive conditions. This suggests that hsk1-1312 causes intrinsic
damage that requires the recombination apparatus for repair, even at permissive
temperature (Dolan et al., 2010).
Fewer origins fire in the cdc7-1/mcm5-bob1 due to the incomplete bypass of the Cdc7
function in activating MCMs complex by the mcm5-bob1 allele. Thus, the MMS
sensitivity could be the consequence of not enough origin firing. Less origin firing
generates fewer forks to sense the damage, so there is less checkpoint activation (Tercero
et al., 2003). However, the checkpoint activation seems not to be the reason for the
sensitivity since there is no obvious sensitivity seen in mec1-100 cell, which is defective
in activating the checkpoint but maintains the fork stability as Cdc7 does. It would be
interesting to see whether firing more origins in Cdc7 will rescue its sensitivity to MMS
using mutants that de-suppress the late origins inhibited by the checkpoint. It would be
interesting to test whether the phosphorylation of Mcm2 by DDK would trigger the
67
change of the fork structure by the change of DNA binding activity in order to facilitate
one way of restarting the fork rather than another way. It would be also interesting to test
whether by introducing the mcm2EE to cdc7-1/mcm5-bob1 rad5∆ would rescue the
sensitivity to MMS in the cdc7-1/mcm5-bob1 rad5∆. Alternatively, we could address this
question by testing whether Rad5 has a synergistic defect with mcm2AA.
10.3 Cdc7 functions in checkpoint activation in response to DNA damage
In Schizosaccharomyces pombe, Hsk1
Cdc7
undergoes Cds1
Rad53
-dependent
phosphorylation in response to HU, and it is a direct substrate of purified Cds1
Rad53
in
vitro (Snaith et al., 2000). Cds1 is poorly activated in hsk1 mutants after HU treatment,
indicating that there may be a feedback loop linking these two kinases (Snaith et al., 2000,
Takeda et al., 2001). Genetic and physical interactions between Cdc7 and Rad53 have
also been described in budding yeast (Dohrmann et al., 1999). Dbf4 also undergoes
Rad53/Cds1 dependent phosphorylation after HU treatment (Takeda et al., 1999,
Weinreich et al., 1999) . This phosphorylation of Dbf4 reduces the Cdc7/Dbf4 kinase
activity and inhibits late origin firing (Weinreich et al., 1999). Furthermore, there is a
Rad53-dependent removal of Dbf4 from chromatin following HU treatment (Pasero et al.,
1999). Recently, it has been shown that combining Sld3 mutant and Dbf4 mutant that
cannot be phosphorylated by Rad53 can bypass the inhibition of late origin firing by the
checkpoint (Lopez-Mosqueda et al., 2010, Zegerman et al., 2010). It is intriguing to
hypothesize that Rad53 dependent phosphorylation of Dbf4 dissociates Cdc7/Dbf4
complex from the late origins to inhibit firing in response to DNA damage in S phase
68
while redirecting the complex to the replication fork to regulate the fork progression or
replication fork restart when the fork is stalled by the lesion on the template. However,
we do not see any difference in the dbf4-4A cells (Zegerman et al., 2010) in terms of the
replication fork progression rate in MMS. It indicates that the regulation of DDK by
Rad53 in S phase is not related with fork rate. It also implies that DDK is still active after
phosphorylation of Dbf4 by Rad53; otherwise we should see the faster fork as we saw in
Cdc7 mutants cells. This is consistent with the finding that complex formation, chromatin
association, and kinase activity of DDK are not inhibited during the DNA-damage-
induced S-phase checkpoint response in Xenopus egg extracts and mammalian cells
(Tenca et al., 2007, Tsuji et al., 2008). Cdc7 is an active kinase in human cancer cells
undergoing replication stress (Tenca et al., 2007). Also more origins fire in dbf4-4A
which might be the reason that we did not see faster forks.
The compromised checkpoint activation might be because that Cdc7 is directly involved
in Rad53 phosphorylation. However, direct evidence is lacking. It is more likely that less
ssDNA is presented in cdc7 mutant cells which in turn have less checkpoint activation
either from less forks that are established from limited origin firing or less uncoupling of
the helicase and polymerase happening on the fork.
Checkpoint activation is compromised in cdc7-1/mcm5-bob1. This is consistent with the
findings that that deletion of the N-terminal inhibitory domain of Mcm4 can bypass the
requirement for DDK in initiation of replication in unperturbed S phase, but it does not
69
bypass its requirement for intra-S-phase checkpoint response in the presence of
hydroxyurea (Sheu et al., 2010). It also has been shown that DDK plays a role in the full
activation of rad53 (Ogi et al., 2008).
Checkpoint activation and replication fork recovery is a dynamic process, since Rad53
could be phosphorylated by the kinases and consistently dephosphorylated by the
phosphatases. Pph3-Psy2 is a phosphatase complex required for Rad53
dephosphorylation and replication fork restart during recovery from DNA damage
(O'Neill et al., 2007, Szyjka et al., 2008). We observed that the checkpoint activation
could be increased by deletion of PPH3, which indicates that Cdc7 is upstream of Pph3
in checkpoint activation pathway. It would be interesting to test whether the checkpoint
would still be reduced if we allow all the origins firing in the beginning of S phase in
MMS by introducing the SLD3-m25 and dbf4-m25 then inactivate Cdc7 activity.
70
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Zegerman, P. and Diffley, J. F. X. (2010). Checkpoint-dependent inhibition of DNA
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478.
77
Appendix A: Cdc7 is required for fork stalling in response to
HU
α factor arrested 3 hrs at 23°C---->release at 23°C into 0.2M HU for 1hr----> raise Tm to 32°C for 2hrs
=
Figure A.1 Cdc7 is needed for maintaining checkpoint activation in response to HU. JPy8 (WT) and
JPy9 (cdc7-1/mcm5-bob1 ) were arrested in G1 phase at 23˚C with α factor for 4hrs. Then cells were
released synchronously from G1 arrest into YEPD medium containing 0.033% MMS at 23°C (Time=0).
Then Tm was shifted to 32°C to inactivate Cdc7 activity after an hour at 23˚C. Cell samples were collected
for TCA preps at indicated time points for looking at Rad53 activation by western blot.
mins
Figure A.2 Cdc7 is required for proper fork stalling in 0.2M HU. JPy8 (WT) and JPy9 (cdc7-1/mcm5-
bob1) were arrested in G1 phase at 23˚C with α factor for 4hrs. Then cells were released synchronously
from G1 arrest into YEPD medium containing 0.033% MMS at 23°C (Time=0). Then Tm was shifted to
32°C to inactivate Cdc7 activity after an hour at 23˚C. Aliquots of the culture were exposed to BrdU for 1hr
pulses as indicated in the diagram. Cells were harvested at the end of the each pulse for DNA isolation.
replication fork progression was traced by chasing BrdU incorporation using BrdU-IP-chip on Nimblegen
12x135K array slide.
WT
WT
cdc7-1/mcm5-bob1
0-60 23˚C
60-120 32˚C
120-180 32˚C
Cdc7-1/mcm5-bob1
78
Figure A.3 RPA-ChIP-Chip in WT and cdc7-1/mcm5-bob1 in response to 0.2M HU JPy8 (WT) and
JPy9 (cdc7-1/mcm5-bob1) were arrested in G1 phase at 23˚C with α factor for 4hrs. Then cells were
released synchronously from G1 arrest into YEPD medium containing 0.033% MMS at 23°C (Time=0).
Then Tm was shifted to 32°C to inactivate Cdc7 activity after an hour at 23˚C. Cells were collected at
indicated time points for RPA ChIP.
WT
60 23˚C
120 32˚C
180 32˚C
Cdc7-1/mcm5-
bob1
mins
79
Appendix B: Post-replication Repair Pathway and replication
fork progression rate in MMS
A
Figure B.1 A. rad18∆ siz1∆ does not affect fork rate in response to 0.033% MMS Asynchronous WT
(YZy50) JPy5 (rad18 ∆ siz1∆) were arrested in G1 phase at 23˚C with α factor for 4hrs and then cells were
released synchronously into YEPD medium containing 0.033% MMS at 23°C (Time=0). Then Cell
samples were taken for FACS analysis at indicated time points. B. Aliquots of the culture were exposed to
BrdU for 25-min pulses as indicated in the diagram. Cells were harvested at the end of the each pulse for
DNA isolation. Replication fork progression was traced by chasing BrdU incorporation using BrdU-IP-chip
on Nimblegen 12x135K array slide.
B.
80
Appendix C: PFGE analysis for chromosome replication in
constant MMS treatment in WT and cdc7-1/mcm5-bob1
WT cdc7-1/mcm5-bob1
Figure C.1 PFGE analysis for chromosome replication in constant MMS treatment in WT and cdc7-
1/mcm5-bob1 A. Asynchronous WT (YZy50) and cdc7-1/mcm5-bob1 (1307) were arrested in G1 phase at
23˚C with α factor for 3hrs and then transferred to 32°C to inactivate Cdc7 activity for 1hr before being
released into S phase. Then cells were released synchronously into YEPD medium containing 0.033%
MMS at 32°C (Time=0) for 7hrs. Cell samples were taken for FACS analysis at indicated time points. B.
Samples were taken at indicated time point for PFGE analysis as in material and methods.
81
Appendix D: The synergist between Rad5 deletion and cdc7as3
Figure D.1 The synergist between Rad5 deletion and cdc7as3A. Asynchronous cdc7as3 (YZy19) and
cdc7as3 rad5∆ (JYy7) were arrested in G1 phase at 23˚C with α factor for 4 hours. Then cells were divided
in half and released synchronously into YEPD medium containing 0.033% MMS either in the presence or
absence of 25μM PP1. Then DNA content by FACS analysis samples were collected at time points
indicated. B. Aliquots of the culture were exposed to BrdU for 25-min pulses as indicated in the
diagramThere is a 5 mins overlapping between each time point to allow the cells to take up BrdU between
the time points. Cells were harvested at the end of the each pulse for DNA isolation. Replication fork
progression was monitored by tracing BrdU incorporation using BrdU-IP-chip using Nimblegen 12x135K
array slide.
82
Appendix E: Pph3 deletion can not rescue the sensitivity of
cdc7-1/mcm5-bob1 rad5∆ to MMS
Figure E.1 Deletion of Pph3 does not rescue the sensitivity of cdc7-1/mcm5-bob1 to MMS YZy50 (WT)
RSy1307 (cdc7-1/mcm5bob1) and YZy34 (pph3∆) and YZy35 (cdc7-1/mcm5-bob1 rad5∆) were spotted
with 10μl in 10-fold dilution series starting from OD 0.5 onto YEPD plates containing the indicated
concentrations of MMS and YEPD for control, as described in Materials and Methods. Each plate series
was incubated at 32°C for 2 days.
Figure E.2 pph3∆ could not rescue the synergistic defect of rad5 and cdc7-1/mcm5-bob1. YZy30
(cdc7-1/mcm5-bob1 rad5∆) and YZy35 (cdc7-1/mcm5-bob1 rad5∆ pph3∆) were arrested in G1 phase at
23˚C with α factor for 3hrs and shifted to 32˚C for 1hr then cells were released synchronously into YEPD
medium containing 0.033% MMS at 32°C (Time=0). Then Cell samples were taken for FACS analysis at
indicated time points
cdc7-1/mcm5-bob1
rad5∆ pph3∆
200
180
160
140
120
100
80
60
40
20
0
cdc7-1/mcm5-bob1
rad5∆
83
Appendix F: Fork progression and checkpoint activation after
cdc7as3 inactivation at latter S Phase
Figure F.1 Fork progression and checkpoint activation after inactivation of cdc7as3 activity at latter
S Phase A. Asynchronous cdc7as3 (YZy10) were arrested in G1 phase at 23˚C with α factor for 4 hours.
Then cells were divided in half and released synchronously into YEPD medium containing 0.2M HU for
1hr. Then cells were centrifuged down and washed once with YEPD then released into YEPD in the
presence or absence of 25μM PP1 at 23˚C. Then MMS was added at the concentration of 0.033% after 30
mins (Time=0). Then DNA content by FACS analysis samples were collected at time points indicated. B.
Then cells were collected at the indicated time point for western blot analysis with anti-Rad53
antibody.“HU 1hr” is one of the two samples at that time point was loaded to save the lanes. C. Aliquots of
the culture were exposed to BrdU for 15-min pulses as indicated in the diagram. When cells were in MMS,
15 mins time pulses were taken. There is a 5 mins overlapping between each time point to allow the cells to
take up BrdU between the time points. Cells were harvested at the end of the each pulse for DNA isolation.
Replication fork progression was monitored by tracing BrdU incorporation using BrdU-IP-chip using
Nimblegen 12x135K array slide.
84
HU 1hr
YEPD 30m
MMS 0-15
MMS10-25
MMS20-35
MMS40-55
MMS60-75
cdc7as3
cdc7as3+PP1
α factor arrest 4hrs release into HU 1h
YEPD without PP1
Add MMS0.033%
YEPD with PP1
30mins
A
B
C
85
Appendix G: List of strains
Table G Strains used in this study
Strain Name Genotype
YZy8 ade2-1 ura3-1 his3-11,15 trp1-1 BrdU-Inc::LEU2 can1-100 ARS608::His3 ARS609::Trp1 RAD5+ bar1::HisG CDC7WT
YZy10 ade2-1 ura3-1 his3-11,15 trp1-1 BrdU-Inc::LEU2 can1-100 ARS608::His3 ARS609::Trp1 RAD5+ bar1::HisG cdc7as3
YZy18 ade2-1 ura3-1 his3-11,15 trp1-1 BrdU-Inc::LEU2 can1-100 ARS608::His3 ARS609::Trp1 RAD5+ bar1::HisG CDC7WT ARS305::BrdUInC(URA3)
YZy19 ade2-1 ura3-1 his3-11,15 trp1-1 BrdU-Inc::LEU2 can1-100 ARS608::His3 ARS609::Trp1 RAD5+ bar1::HisG cdc7as3 ARS305::BrdUInC(URA3)
YZy50 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1)
RSy1298 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1
RSy1307 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1
YZy35 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 pph3::KanMX
YZy55 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) dbf4-4A
Cvy65 ade2-1 ura3-1 his3-11,15 trp1-1 p405-BrDU(leu2) can1-100 ARS608::HIS3 ARS609::TRP1 RAD5+ bar1::HisG mrc1-AQ
SSy212 ade2-1 ura3-1 his3-11,15 trp1-1 p405-BrdU(leu2) can1-100 ars608∆::HIS3 ars609∆::TRP1 RAD5+ bar1::hisG
YZy52 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) mec1-100
YZy28 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad5::KanMX
JPy3 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad30Δ rev3::KanMX
JPy4 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad30Δ rev3Δ
YZy40 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 rad30Δ::KanMX
rev3Δ
YZy30 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 rad5::KanMX
YZy53 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad30Δ rev3Δ rad5::kanMX
T2y41 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 orc1::hisG leu2::ORC1 (LEU2) ars305::BrdU-Inc (KanMX)
T2y42 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 bar1::hisG ars608::HIS3 ars609::TRP1 orc1::hisG leu2::orc1-161 (LEU2) ars305::BrdU-Inc (KanMX)
YZy64 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) mcm2AA
JPy5 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad18:: KanMX siz1:: ura3
JPy8 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) RFA1-13Myc::KanMX
JPy9 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 RFA1-
13Myc::KanMX
JYy6 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 rad5D::KanMX
pph3D
YZy59 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) mec1-100 Rad5::KanMX
YZy46 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 rad5GAA
YZy44 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5 bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) cdc7-1 mcm5-bob1 rad5I916A
YZy41 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad5GAA
YZy42 ade2-1 ura3-1 his3-11,15 trp1-1 leu2-3,112 can1-100 RAD5+ bar1::hisG ars608::HIS3 ars609::TRP1 ars305::BrdU-Inc (TRP1) rad5I916A
Abstract (if available)
Abstract
Cdc7-Dbf4 is an essential protein kinase complex required for every single origin firing. As a target of the intra-S checkpoint, Cdc7 kinase activity has also been implicated in the response to replication fork stress, with a role in translesion DNA synthesis (TLS). We have examined the role of Cdc7 in the regulation of replication forks, particularly in response to MMS, which normally stalls replication forks and inhibits late origin firing. We find that replication forks proceed as fast as with no damage along an MMS-damaged template both in cdc7as3 and cdc7-1/mcm5-bob1 cells. However the DNA synthesis in cdc7-1/mcm5-bob1 in MMS is defective, indicated by the slower recovery after MMS by PFGE, suggesting the replication is incomplete. These deregulated forks did not rely on TLS pathway but are dependent on both helicase and E3 ligase function of Rad5 for continued fork progression along MMS-damaged DNA, demonstrating a role for Rad5 at the replication fork. Phosphorylation of MCM2 by DDK was not sufficient for slowing down Mcm2-7 helicase activity in vivo. Temperature-sensitive mutant orc1-161, which is defective in pre-replication complex (pre-RC) assembly, phenocopies the defects in origin firing and faster fork progression in MMS in Cdc7 mutants, suggesting that decreased origin firing is the common source for the deregulated fork progression. We attribute the effect of Cdc7 depletion on replication fork progression in MMS to the reduced origin-firing, which leads to less established forks and in turn the deficient checkpoint activation in MMS. The mec1-100 cells, which initiate many origins but also have compromised checkpoint activation, fail to phenocopy the faster fork movement in MMS, suggesting that the number of active replication forks also influences fork rate, perhaps due to competition for limiting factors for DNA replication. These findings provide new insights into how Cdc7, besides its essential role in replication initiation, could imply a function in the regulation of replication fork progression in response to DNA damage possibly through checkpoint activation and replication fork restart in MMS.
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Creator
Zhong, Yuan
(author)
Core Title
The role of Cdc7 in replication fork progression in response to DNA damage
School
College of Letters, Arts and Sciences
Degree
Doctor of Philosophy
Degree Program
Molecular Biology
Publication Date
05/01/2012
Defense Date
03/08/2012
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University of Southern California
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Aparicio, Oscar M. (
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), Forsburg, Susan (
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