Identification of TICRR, a novel checkpoint and
replication regulator
MASSACHUSETTS INSTITJTE
OF TECHNOLOGY
by
MAY 25 2011
Nelly Marie Cruz
LIBRARIES
B.S. University of Puerto Rico
Mayaguez, PR 2005
ARCHIVES
SUBMITTED TO THE DEPARTMENT OF BIOLOGY
IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTORATE OF PHILOSOPHY
AT THE
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
JUNE 2011
C 2011 Massachusetts Institute of Technology
All rights reserved
Signature of Author....
Nelly M. Cruz
Department of Biology
May 9, 2011
Certified by...........
Jacqueline A. Lees
Professor of Biology
Thesis Supervisor
Accepted by......
........................................
Robert T. Sauer
Professor of Biology
Chair, Biology Graduate Committee
Identification of TICRR, a novel checkpoint and
replication regulator
by
Nelly M. Cruz
Abstract
The eukaryotic cell cycle refers to a sequence of events by which a cell
duplicates its genomic DNA and divides into two daughter cells. Deregulation of the cell
cycle can cause aberrant cell proliferation, as well as genomic and chromosomal
instability, events that contribute to the development of cancer. Along with the machinery
that promotes cell cycle progression, cells have evolved surveillance mechanisms, or
checkpoints, that protect the cells from DNA lesions. Understanding the molecular
mechanisms by which checkpoints act is of clinical relevance, since mutations in
checkpoint components are often associated with human developmental disorders and
cause a predisposition to cancer.
A collection of 336 zebrafish (Danio rerio) lines, each carrying a recessive
embryonic lethal mutation caused by a retroviral insertion, was screened for novel genes
required for the ionizing radiation-induced G2/M checkpoint. Among the mutant lines that
displayed a defect in checkpoint activation, two carry insertions in a novel gene that we
have named ticrr (for TopBP1 interacting, checkpoint and replication regulator). The loss
of ticrr impairs DNA replication and disrupts the S/M checkpoint, leading to premature
mitotic entry of cells with partially replicated genomes and mitotic catastrophe.
Therefore, Ticrr is a novel cell cycle regulator essential for genomic integrity with roles in
replication as well as in the S/M and G2/M checkpoints.
We have identified the human ortholog of Ticrr and showed that both the human
and zebrafish Ticrr proteins associate with TopBP1, a protein with known roles in
checkpoints and replication. We hypothesized that TICRR is required for pre-initiation
complex assembly during replication initiation, in an analogous manner to the TopBP1
yeast ortholog Dpbl 1 and the yeast protein Sld3. Consistent with this model, we show
that ticrr-deficiency disrupts chromatin association of pre-initiation complex, but not prereplication complex, components in the zebrafish.
The work described in this thesis demonstrates the utility of zebrafish for
performing genetic screens for cell cycle regulators. We have used human cell lines to
complement our studies in the zebrafish and showed that taking advantages of the
strengths that each system offers represents a powerful strategy to elucidate gene
function.
Thesis Supervisor: Jacqueline A. Lees
Title: Professor of Biology
Acknowledgments
I owe my deepest gratitude to my advisor Jackie Lees, for her guidance,
continuous support, and encouragement, which have been invaluable to my growth as a
scientist over the years. I would also like to thank the members of my thesis committee,
Frank Solomon, Angelika Amon and Steve Bell; as well as Nancy Hopkins for their
insightful comments and contributions to my research.
I am grateful to my friends and colleagues in the Koch Institute, specially past
and present members of the Hopkins and Lees lab, for sharing reagents and their
knowledge with me and making the Koch Institute a great place to work. I would like to
particularly thank Chris Sansam, who helped me greatly throughout my graduate career
and Crystal Lee, who has been an excellent classmate, benchmate and friend. I would
also like to show my gratitude to Simona, Keren, Mindy, Tiziana and Amy, for great
companionship and all the positive energy they bring to the lab.
I am indebted to my family and friends for their unconditional love and support.
The encouragement of many friends has been indispensable and I would like to
particularly acknowledge Grisel, Gisselle, Ingrid, Ricky and Francis for always being
there for me. I would also like to mention my grandparents Nelida, Jose M., Merida and
Guillermo, who have been a true inspiration for me. I am very grateful to my brother Jose
G., my sister Cristina, my boyfriend Jose Luis and my parents Jose E. and Ana for
innumerable things, especially for always believing in me and supporting all my
endeavors. They have been a driving force for me through my graduate studies and to
them I dedicate this thesis.
Table of Contents
A bstra ct ............................................................................................................................
2
Acknowledgments ......................................................................................................
3
CHAPTER 1: Introduction ...........................................................................................
9
O ve rv iew ..........................................................................................................................
10
The eukaryotic cell cycle .............................................................................................
11
Cyclin-dependent kinases and the regulation of the cell cycle .................... 11
Regulation of replication initiation in eukaryotes ..........................................................
15
Pre-replication complex (pre-RC) assembly.............................................................
17
Pre-initiation complex (pre-IC) assembly.................................................................
20
Mechanism for prevention of re-replication
26
.............................
Cell cycle checkpoints ................................................................................................
28
ATM and ATR: initiators of the signaling cascade in response to DNA damage.........31
The G1/S checkpoint ..............................................................................................
34
DNA-damage induced G2/M arrest .............................................................................
35
Checkpoints during S-phase ..................................................................................
36
The zebrafish as a model system for the study of cell cycle regulation....................... 39
R e fe re nc e s ......................................................................................................................
41
CHAPTER II: A zebrafish screen for novel DNA damage response genes identifies
TICRR..............................................................................................................................51
Results.............................................................................................................................
A genetic screen for zebrafish mutants with a defective IR response
55
........... 55
Insertional mutations in zebrafish that affect normal mitotic progression................60
Disruption of de novo purine nucleotide synthesis in zebrafish abrogates IR-induced
cell cycle arrest............................................................................................................62
An insertional mutation in a novel gene that abrogates IR-induced cell cycle arrest in
th e z e b ra fish ................................................................................................................
63
Disc ussio n .......................................................................................................................
What role do the identified genes play in normal cell cycle progression? ...............
What role do the identified genes play in the DNA damage response? .........
. ....... . .
71
71
72
Are the identified genes required for normal cell cycle progression and checkpoints in
...... .............. .............. ... ........ . . 74
other vertebrates? .....................................................
Materials and methods ................................................................................................
Zebrafish maintenance
G2/M checkpoint assays
.......................................
75
..... 75
.......................................... 75
Genotyping and expression analysis of zebrafish lines...........................................
75
In Vitro Kinase Assay ..............................................................................................
76
FACS Analysis.............................................................................................................
77
Acknow ledgem ents ....................................................................................................
77
References ......................................................................................................................
77
CHAPTER III: TICRR associates with TopBP1 and is required for replication
initiation..........................................................................................................................80
S um m a ry .........................................................................................................................
81
Intro duction ......................................................................................................................
82
Re sults .............................................................................................................................
85
Disruption of ticrr impairs S-phase progression and causes premature chromatin
condensation ...............................................................................................................
85
TICRR binds TopBP1
BPI............................................
..................................................
90
Ticrr-deficiency inhibits pre-IC form ation .................................................................
98
Disc us sio n .....................................................................................................................
100
TICRR in DNA Replication ........................................................................................
100
TICRR in the SIM and G2/M Checkpoints................................................................. 102
Materials and methods ..................................................................................................
Human cell culture and chromatin fractionation .
Immunoprecipitation .......
..............
.........
105
................107
....................... 107
Zebra fish chrom atin preparation................................................................................108
Antibodies..................................................................................................................108
Acknow ledgem ents .......................................................................................................
108
R efe re nce s ....................................................................................................................
109
CHA PTER IV: Discussion ...........................................................................................
112
Key conclusions.............................................................................................................113
A screen in zebrafish identifies Ticrr....................................113
Ticrr is required for DNA replication and functions in a similar manner to Sld3 ............ 114
The role of CDK activity in regulating TICRR function in DNA replication.....................117
Other proteins with functional similarity to TICRR.........................................................118
Possible roles of TICRR in the maintenance of genome stability..................................119
R e fe re nc e s ....................................................................................................................
12 2
APPENDIX A: Characterization of the functional significance of the TICRRTopBP1 interaction......................................................................................................125
Re sults a nd Discussion .................................................................................................
126
The TopBP1 interaction domain lies within the C-terminal half of TICRR ................. 126
Two putative CDK phosphorylation sites are highly conserved in TICRR ................. 127
A TICRR mutant that localizes to PCNA and RPA foci .............................................
Mate rials and methods ..................................................................................................
133
13 8
Human cell culture and transfections ........................................................................
138
Immunoprecipitation .. ..........................................................................
139
Immunofluorescence
139
............................................
A n tib o dies ..................................................................................................................
14 0
A cknow le dg eme nts .......................................................................................................
14 0
R e fe re nce s ....................................................................................................................
14 0
Figures and Tables
CHAPTER I: Introduction ...........................................................................................
9
Figure 1. Overview of the CDK-Cyclin complexes and the periods of the cell cycle at
w hich they are active................................................................................................
. . 13
Figure 2. Regulation of Rb and E2F activities in mid-late G1............
...... 14
Figure 3. Sequential assembly of pre-RC proteins to origins leads to loading of the
MCM2 -7 helicase .............................................................................................................
20
Figure 4. Recruitment of additional proteins to origins leads to formation of the pre-IC
and subsequent activation of the MCM2-7 helicase.............................
...................... 25
Figure 5. General scheme of the signal transduction pathways that are activated in
.... 30
....................
response to DNA damage or stalled replication forks.
Figure 6. A Simplified scheme of DNA damage-induced cell cycle arrest and apoptosis.
.........................................................................................................................................
36
CHAPTER II: A zebrafish screen for novel DNA damage response genes identifies
T ICRR..............................................................................................................................51
Figure 1. Zebrafish can be used to screen for IR-induced G2 checkpoint response
ge ne s...............................................................................................................................
56
Figure 2. A zebrafish screen for novel DNA damage response and cell cycle regulators.
.........................................................................................................................................
58
Figure 3. hi1487 homozygous mutants show an aberrant accumulation of pH3 positive
cells in the absence of exogenous DNA damage........................................................62
Figure 4. Mitosis after irradiation phenotype in hi923 mutants.
.............
..... 64
Figure 5. hil 573 and hi3202A zebrafish embryos show a Mitosis After Irradiation (MAI)
phe noty pe........................................................................................................................
65
Figure 6. The checkpoint defect results from disruption of a novel gene, ticrr. ..........
67
Figure 7. Cdc2 kinase activity remains high in ticrr mutants after IR treatment. ......... 68
Figure 8. Cells in ticrr homozygous mutants continue to enter mitosis after IR treatment.
.........................................................................................................................................
69
Table 1. Insertional mutants from the Hopkins zebrafish collection that showed an
increase in pH3 staining after IR treatment. . ..................
.......
......... 61
Table 2. Conservation between D. rerio Ticrr and orthologs identified by psiBLAST......70
CHAPTER III: TICRR associates with TopBP1 and is required for replication
initiation..........................................................................................................................80
Figure 1. Ticrr is required for normal DNA replication. ............................
Figure 2. Ticrr is required for S/M checkpoint function.
..................
86
.....
88
Figure 3. A DNA replication defect but not premature mitotic entry occurs in 24 hpf ticrr
muta nts ............................................................................................................................
90
Figure 4. Human TICRR is a chromatin-associated protein....
................ 93
Figure 5. Human TICRR interacts with TopBP1..........................................................94
Figure 6. Human TopBP1 interacts with zebrafish TopBP1.
...............
Figure 7. TICRR interacts with the first two BRCT domains in TopBP1
.....
95
............ 96
Figure 8. TICRR can bind TopBP1 in a CDK independent manner..........
.... 98
Figure 9. Ticrr is essential for the chromatin association of the pre-IC component Psfl.99
CHA PTER IV: Discussion ...........................................................................................
112
APPENDIX A: Characterizing the functional significance of the TICRR-TopBP1
interactio n ....................................................................................................................
125
Figure 1. Residues 976-1261 of TICRR protein are required and sufficient for TopBP1
bindin g...........................................................................................................................
127
Figure 2. Sequence alignment of conserved CDK consensus motifs in the TICRR
pro te in ............................................................................................................................
129
Figure 3. Mutation of Thr 969 and Ser 1001 in the TICRR protein causes a reduction in
T o pBP1 bin din g .............................................................................................................
13 0
Figure 4. Loss of TICRR in human cells causes a replication defect. ........................... 131
Figure 5. Ticrr is required for S/M checkpoint function in human ce is.....
..... 132
Figure 6. TICRR knockdown induces cellular senescence in U2OS cells.....................133
Figure 7. GFP-TICRR'
565-19
1 0 localizes
Figure 8. GFP-TICRR'
565~1
91 0foci
to foci within the nucleus..........
do not colocalizes with 5H2AX. ..............
Figure 9. GFP-TICRR 1 565-1910 foci colocalizes with RPA and PCNA.
...... 134
135
.............. 137
Figure 10. Expression of GFP-TICRR 1565-1910 impairs S-phase progression..................138
CHAPTER 1: Introduction
Overview
The cell cycle refers to a sequence of events by which a cell duplicates its
hereditary material and divides into two daughter cells. This process is the basis for
growth and development of all living organisms. The cell cycle is divided into four
phases: G1, S, G2 and M phase. G1 refers to a gap during which the cell prepares for
DNA synthesis occurring during S phase. The replication of chromosomal DNA is
followed by a gap called G2 during which the cell prepares for mitosis, or M phase.
During M phase the replicated chromosomes are segregated and cells divide. There is a
decisive moment in G1, referred to as the restriction point in mammalian cells and start
in yeast, in which the cell commits to enter the cell cycle. Under specific circumstances,
cells in G1 can enter a resting state called GO before committing to DNA replication.
Cells in GO account for the majority of the non-proliferating cells in the human body
(Nurse, 2000).
Deregulation of the cell cycle can cause aberrant cell proliferation, as well as
genomic and chromosomal instability, events that contribute to the development of
cancer (Malumbres and Barbacid, 2009). Therefore, cell cycle events are under precise
temporal control. Along with the machinery that promotes cell cycle progression, cells
have surveillance mechanisms, or checkpoints, that ensure the correct ordering of
events and trigger cell cycle arrest in response to DNA damage or errors. The work
described in this thesis has led to the identification of TICRR, (for TopBP1 interacting,
checkpoint and replication regulator), a novel protein essential for normal cell cycle
progression. In the following introduction, I will review the regulatory mechanisms of the
eukaryotic cell cycle with a particular focus on replication initiation, a process in which
TICRR plays an important role. I will then discuss the checkpoints that ensure the fidelity
of cell division and protect the cell from the deleterious effects of DNA damage. Finally, I
will discuss the advantages of using zebrafish as a model organism for the identification
of novel cell cycle and checkpoint regulators.
The eukaryotic cell cycle
Cyclin-dependent kinases and the regulation of the cell cycle
Cyclin dependent kinases (CDKs), best described as the drivers of eukaryotic
cell cycle progression, are proline-directed serine/threonine protein kinases that
phosphorylate substrates appropriate for the phase of cell cycle that the kinase
regulates. In yeast, cell cycle progression is controlled by a single CDK, called Cdc28 in
Saccarhomyces crevisiae and Cdc2 in Schizosaccharomyces pombe (Forsburg and
Nurse, 1991).
This mechanism is more complicated in vertebrates, with four CDKs
(CDK1, 2, 4 and 6) playing a role in regulating the cell cycle (Satyanarayana and Kaldis,
2009). CDKs are regulated in several ways to ensure that their activity is limited to
specific stages. Cyclins, the CDK binding partners, comprise a diverse family of proteins
first identified in sea urchin eggs and later found to be conserved from yeast to humans
(Evans et al., 1983; Lew and S, 1992). Their levels oscillate throughout the cell cycle,
accumulating at a specific stage where they bind and promote CDK activity. Like yeast,
mammalian cells express multiple cyclins.
In the current model for the mammalian cell cycle, each phase is driven by
specific CDK-cyclin combinations (Figure 1)(Malumbres and Barbacid, 2009). Mitogenic
signals induce expression of the D-type cyclins (D1, D2 and D3) that preferentially bind
and activate CDK4 and CDK6 during G1 (Sherr, 1995). Key substrates of the G1 CDKcyclin complexes are the pocket proteins, pRb, p107 and p130, which repress E2F
activity by directly binding to them (Figure 2). The E2F family of transcription factors
activate genes encoding proteins involved in cell cycle regulation and DNA synthesis,
including the late-G1 cyclin E and the S-phase cyclin A and CDK2. Phosphorylation of
the pocket proteins at multiple sites prevents their association with E2Fs, thereby
allowing the transcription of genes required for S entry (Figure 2). This is one of the
principal events for passage through the restriction point (Trimarchi and Lees, 2002).
In turn, E-type cyclins bind and activate CDK2, which further phosphorylate the
pocket proteins, leading to their complete inactivation. CDK2-cyclin E drives the G1/S
transition (Tsai et al., 1993). CDK2 is subsequently bound by cyclin A during late stages
of DNA replication to drive the transition to G2. Finally, CDK1 is thought to be activated
by cyclin A at the end of G2 to facilitate the onset of mitosis. Following nuclear envelope
breakdown, cyclin A is degraded, facilitating formation of the CDK1-cyclin B complex
responsible for driving cells through mitosis.
Regulatory mechanisms of CDK activity
As stated above, the binding of a cyclin subunit is required for CDK activity. In
this section, I will provide a brief description of the additional mechanisms of CDK
regulation. All these regulatory mechanisms act in concert at the right time to promote
the transition from one phase of the cell cycle to the other.
Phosphorylation
Full activation of the CDK-cyclin complex usually requires phosphorylation of a
conserved threonine residue located on the activation loop of the kinase domain (Thr161 in CDK1, Thr-60 in CDK2, and Thr-172 in CDK4). These phosphorylations induce
conformational changes and enhance the binding of cyclins (Jeffrey et al., 1995). On the
other hand, CDK1 activity can be inhibited by phosphorylation of residues within the
ATP-binding pocket of the CDK (Thr-14 and Tyr-15 in CDK1) by Weel and Myt1
kinases. Dephosphorylation at these sites by the Cdc25 family of phosphatases is
necessary for activation of CDK1 and further progression through the cell cycle (Lew and
Kornbluth, 1996) (Figure 1).
Cdc25A
CDK4/6
cyclin D1/2/3
CDK2
Cyclin E
T
Cdc25A/B/C
CDK2
Cyclin A
S
sil
CDK1
Cyclin A
G2
CDK1
Cyclin B
MI
Restriction
Point
Figure 1. Overview of the CDK-Cyclin complexes and the periods of the cell cycle
at which they are active.
Mammalian cells use several CDKs and cyclins to regulate passage through the cell cycle. The
activity of mammalian CDK-cyclin complexes is induced by Cdc25 phosphatases, which catalyze
the removal of inhibitory phosphorylations on the CDK subunit. Adapted from Trimarchi and
Lees, 2002.
CDK inhibitors
There are two families of CDK inhibitors that have been defined on the basis of
their primary sequence and biochemical properties (Figure 2). The INK4 family includes
p15, p16, p18, and p19, which specifically inactivate the G1 CDKs (CDK4 and CDK6).
These form stable complexes with the CDK before cyclin binding, preventing association
of cyclin D. The Cip/Kip family includes p21, p27 and p57, and they inhibit a broader
spectrum of CDK-cyclin complexes, including CDK2-cyclinE, CDK2-cyclin A, CDK1cyclin A and CDK1-cyclin B. They modulate CDK activity by interacting with both
subunits of the CDK-cyclin complex. p21 also inhibits DNA synthesis by binding to and
inhibiting PCNA, a protein that acts as a processivity factor for DNA polymerase (Waga
S 1994). These inhibitors are regulated by a variety of mechanisms; for example, the
expression of p21 is under the transcriptional control of the p53 tumor suppressor (Sherr
and Roberts, 1995).
INK 4
CycD CDK4
CIP/KIP
(v3E
CVA CDK2
CDK2
G1
S
Figure 2. Regulation of Rb and E2F activities in mid-late G1.
The pocket protein Rb represses E2F activity by directly binding to E2F. CDK4/6-Cyclin D
complexes phosphorylate Rb, releasing some E2F, which stimulates transcription of cyclin E and
CDK2. The CDK2- Cyclin E complex further phosphorylates Rb, resulting in rapid increase of
E2F and increased CDK2 activity drives passage through the restriction point. Two types of CDK
inhibitors contribute to cell cycle control in mammals, the INK4 and CIP/KIP family of proteins,
which act by binding to CDK and inhibiting its activity. Adapted from Trimarchi and Lees, 2002.
Intracellular localization
The intracellular localization of different cell cycle-regulating proteins also
contributes to correct cell cycle progression. Cyclin B is actively exported from the
nucleus until the beginning of prophase. The 14-3-3 group of proteins regulate the
intracellular trafficking of various proteins. For instance, the CDK activating phosphatase
Cdc25 is kept in the cytoplasm during interphase through interaction with 14-3-3 proteins
(Wilker and Yaffe, 2004).
Role of protein degradation in cell cycle control
The activity of CDKs is also controlled by the ubiquitin-mediated proteolysis of
key regulators such as cyclins and CDK inhibitors. Two E3 ubiquitin ligases, the SkplCULl-F-box (SCF) complex and the anaphase-promoting complex/cyclosome (APC/C)
mediate the specific ubiquitination of these regulators (Nakayama and Nakayama,
2006). Two specific events are very well characterized; specifically, the SCF regulates
entry into S-phase by targeting CDK inhibitors and G1 cyclins for degradation (Ang and
Wade Harper, 2005). The APC/C is active from mid-mitosis to the end of G1 and
facilitates exit from mitosis by targeting the mitotic cyclins, A and B, for degradation. The
APC/C also plays a major role in controlling the onset of anaphase and is the key target
of the spindle assembly checkpoint (Wasch and Engelbert, 2005).
Regulation of replication initiation in eukaryotes
DNA replication is the process by which cells duplicate their genome prior to cell
division. To duplicate their genomes in a timely manner, eukaryotes have many
replication start sites, or origins of replication. Origins in the budding yeast S. cerevisiae
contain an essential 11-bp consensus sequence and a non-conserved region that is also
important for origin activity (Marahrens and Stillman, 1992). In contrast, no evident
consensus sequence has been found in origins of other eukaryotes and it is thought that
other factors may facilitate the selection of specific segments in the genome, such as
DNA topology, chromatin structure, gene transcription and nucleosome formation (Masai
et al.). Despite the apparent lack of conservation for origin sequences, the proteins
involved in origin selection are conserved among eukaryotes.
DNA replication initiation is tightly controlled so that each origin is fired only once
and re-replication does not occur. Most of our understanding on the mechanism
underlying replication initiation comes from studies in yeast, a powerful model for
studying molecular genetics and biochemical function. In vitro and in vivo studies in
metazoans have revealed that the key players and events required for replication
initiation are highly conserved among eukaryotes. It has become clear that replication
initiation is a very dynamic and coordinated process involving recruitment of various
protein complexes to origins. The process is divided into two steps, characterized by the
formation of two major complexes, the pre-replication complex (preRC) and the preinitiation complex (prelC). To ensure that DNA replication is restricted to a single round
per cell cycle, these steps are temporally regulated and they take place during different
stages (Diffley et al., 1994). Assembly of pre-RC can only occur during late M and early
G1, when CDK activity is low (Dahmann et al., 1995). When cells enter S phase, the
activity of CDK and Cdc7-Dbf4 (DDK), promote the recruitment of additional proteins to
the pre-RC and the subsequent activation of the replicative helicase MCM2-7 (Labib,
2010). Additionally, CDK plays an important role in inhibiting re-assembly of pre-RC in
origins, an important mechanism to prevent re-replication (discussed in following
sections)(Arias and Walter, 2007).
Below, I will describe the proteins required for replication initiation in eukaryotes
and mechanisms by which these proteins are recruited to the origins and DNA
replication is initiated. I will also describe the mechanisms by which the cell regulates
this process to ensure that the entire genome is duplicated only once. I will focus on
studies in S. cerevisiae, since they provide the framework for the current model of
replication initiation. I will also review relevant findings in metazoans that expand our
understanding of this process in more complex organisms.
Pre-replication complex (pre-RC) assembly
The first step in the initiation of DNA replication is the selection of sites that will
act as origins of replication. This is mediated by the sequential assembly of ORC, Cdc6,
Cdtl and MCM2-7 proteins onto origins to generate a pre-RC (Figure 3) (Bell and Dutta,
2002). Pre-RC formation directs the loading of the Mcm2-7 helicase onto origin DNA, a
process also known as origin licensing. The loaded MCM2-7 helicase remains inactive
until S-phase entry when two kinases, CDK and DDK, promote its activation (discussed
in the following section).
Recent in vitro studies in budding yeast, combined with in vivo genetics and
biochemical approaches, have revealed details on the mechanism by which the pre-RC
is formed. A key feature is the ATP regulation of various steps. The origin recognition
complex (ORC) is the first pre-RC component to associate with origins and is essential
for the recruitment of other factors (Figure 3B)(Bell, 2002). It consists of six different
subunits, Orcl-6, first identified in budding yeast for its ability to bind origins in an ATP
dependent manner (Bell and Stillman, 1992). Orc1, Orc4 and Orc5 are members of the
AAA+ family of ATPases, while Orc2 and Orc3 are distantly related to this class of
proteins (Erzberger and Berger, 2006). However, only the ATP binding and hydrolysis by
the largest ORC subunit, Orc1, has been shown to be required for ORC function in preRC assembly (Bowers et al., 2004; Klemm et al., 1997). Specifically, Orc1 binding to
ATP promotes association of the complex with origin DNA (Klemm et al., 1997), while
hydrolysis occurs after the initial loading of the MCM2-7 helicase (Randell et al., 2006).
Studies in Drosophila and in Xenopus in vitro system (using recombinant human ORC)
indicate that the requirement of Orc1 ATPase activity for replication is conserved in
metazoans (Chesnokov et al., 2001; Giordano-Coltart et al., 2005).
Like Orc1, Cdc6 is a member of the AAA+ family of ATPases whose ATP binding
and hydrolysis is necessary for pre-RC assembly (Perkins and Diffley, 1998; Randell et
al., 2006; Weinreich et al., 1999). The current model for pre-RC assembly proposes that
ORC recruits ATP-bound Cdc6 to origins, and this complex promotes the recruitment of
a pre-formed Cdtl-Mcm2-7 complex to origins (Figure 3C) (Randell et al., 2006;
Tsakraklides and Bell, 2010). Cdtl exists in a complex with Mcm2-7 (Tanaka and Diffley,
2002) and these two components associate to DNA with similar kinetics at equimolar
ratios (Randell et al., 2006), suggesting that they are recruited to the origins as a
complex. This is most likely mediated through Cdtl binding to Orc6 (Chen et al., 2007).
The next step is the hydrolysis of ATP by Cdc6, which promotes the dissociation
of both Cdc6 and Cdtl from the complex, and triggers tight binding of the MCM2-7
complex with DNA (Figure 3D)(Randell et al., 2006; Tsakraklides and Bell, 2010). Two
recent studies, using an in vitro loading system with purified budding yeast proteins,
show that the inactive helicase is loaded onto double stranded DNA as a head-to-head
double hexamer (Evrin et al., 2009; Remus et al., 2009). The common view is that the
helicase is active as a single hexamer. Indeed, a study in the Xenopus egg extract in
vitro system provides evidence supporting a model in which the two Mcm2-7 complexes
uncouple upon activation and travel away from one another, and therefore function
autonomously (Yardimci et al., 2010). Currently, it is not clear how these two complexes
dissociate and move away from origins to form two replication forks.
The steps described above appear to be repeated multiple times, yielding preRCs containing multiple MCM2-7 complexes (Bowers et al., 2004; Randell et al., 2006).
ORC ATP hydrolysis occurs after Cdc6 ATP hydrolysis and is required for multiple
rounds of MCM2-7 loading (Figure 3E, F)(Randell et al., 2006). The function of the extra
Mcm2-7 complexes at origins is not well understood, but studies in human cells suggest
that they rescue collapsed replication forks upon replication stress (Ge et al., 2007;
Ibarra et al., 2008).
Although previous data indicate that the budding yeast ORC associates with
origins throughout the cell cycle, a recent study involving in vitro reconstitution of preRCs shows that it is released after Mcm2-7 loading and ATP hydrolysis (Tsakraklides
and Bell, 2010). It remains to be seen if release of ORC upon Mcm2-7 loading also
occurs in vivo. The fact that Cdc6, Cdtl and possibly ORC are released from the origins
once the helicase is loaded is consistent with in vitro studies in Xenopus egg extracts
indicating that ORC, Cdc6 and Cdtl are not required to maintain Mcm2-7 association
with the origin after loading is complete (Hua and Newport, 1998; Jares and Blow, 2000;
Rowles et al., 1999). Although it is not known whether the described mechanism
operates in the same fashion in other organisms, the sequential association of ORC,
Cdc6, Cdtl and Mcm2-7 at origins appears to be conserved in other eukaryotes,
including S. pombe and Xenopus.
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Cdc6
CF
Mcm2-7-Cdtl
Cdtl
9
Cdc6
ATP
E
Figure 3. Sequential assembly of pre-RC proteins to origins leads to loading of the
MCM2-7 helicase.
(A, B) ATP-bound ORC binds to origin DNA; red star represents ATP. (C, D) Association of
Cdtl and MCM2-7, which are recruited to origins as a complex, follows the association of ATPbound Cdc6. (E) ATP hydrolysis by Cdc6 leads to the loading of MCM2-7 complex and release
of Cdc6 and Cdtl. (F) ATP-hydrolysis by ORC allows re-initiation of this cycle for further
loading of MCM2-7 helicase complexes onto origins (Adapted from Masai H et al. 2010)
Pre-initiationcomplex (pre-IC) assembly
Initiation of replication requires the recruitment of a number of factors to the
existing pre-RC in a CDK and DDK dependent manner (Figure 4). The large multiprotein
complex that is formed at this step is referred to as the pre-initiation complex (pre-IC).
The main purpose of the pre-IC is to activate the Mcm2-7 helicase. Recent studies
provide insights into how this may occur. GINS and Cdc45 act as cofactors of the
helicase, and these proteins form a complex at the onset of replication initiation. This
complex was first co-purified in Drosophila, but has also been co-purified from S.
cerevisiae and there is evidence for its existence in humans and Xenopus (Aparicio et
al., 2009; Gambus et al., 2006; Moyer et al., 2006; Pacek et al., 2006). Importantly, the
binding of Cdc45 and GINS has been shown to activate Mcm2-7 helicase activity in vitro
(Ilves et al., 2010). The structure of the Cdc45-Mcm2-7-GINS complex has been recently
solved using single-particle electron microscopy and provides insightful details into how
Cdc45 and GINS may promote helicase activation (Costa et al., 2011). This study
proposes that the Mcm2-7 complex loaded onto origins during pre-RC formation exist in
a ring-like open configuration, with a gap between Mcm2 and Mcm5 subunits. This
structure alternates between two conformations and Cdc45 and GINS binding stabilizes
one of the structures by bridging the gap. This is followed by ATP binding, which
promotes ring closure around DNA.
Formation of the Cdc45-Mcm2-7-GINS
complex and subsequent helicase
activation requires CDK and DDK activity. In the following section, I will describe our
current knowledge on how CDK and DDK regulate DNA replication initiation. I will focus
on studies that have been performed in the budding yeast, where currently this process
is best understood.
The role of DDK
The S phase kinase DDK is required for recruiting Cdc45 and other pre-IC
components to origins (Sclafani et al., 2002; Zou and Stillman, 2000). Studies in a
number of organisms indicate that MCM2-7 is the major target of DDK phosphorylation
(Labib, 2010; Masai and Arai, 2002). All budding yeast Mcm2-7 subunits, with the
exception of Mcm5, are phosphorylated by DDK in vitro. The major targets are the amino
terminal tails of Mcm2, Mcm4 and Mcm6 (Labib, 2010). The exact mechanism by which
DDK acts in replication initiation has been unclear, however, new details are starting to
emerge. An interesting study reveals that DDK phosphorylation of Mcm4 alleviates an
inhibitory activity of its N-terminal region (Sheu and Stillman, 2010). Removing the Nterminal domain of Mcm4 allows DDK-independent initiation of DNA replication,
suggesting that Mcm4 is the minimum target of DDK for replication initiation. However, in
the absence of DDK these cells grow slowly and the Cdc45-Mcm2-7 complex is
defective, suggesting that DDK has additional roles during replication (Sheu and
Stillman, 2010). Further characterization of the functional significance of other DDK
targets is needed in order to gain a complete picture of the role of this essential kinase in
replication.
The role of CDK
The protein Dpbl 1 has a dual role in replication initiation and in DNA damage
checkpoints. To further understand Dpbl 1's function, Kamimura et al. identified factors
interacting with Dpbl 1 by isolating synthetically lethal mutants in S. cerevisiae
(Kamimura et al., 1998). The pre-IC components SId2 and Sid3, the major CDK targets
during replication initiation, were identified in this screen. Sld2 is phosphorylated by CDK
in at least 7 sites, which triggers Sid2 binding to Dpbl 1 and this interaction is essential
for replication initiation (Masumoto et al., 2002; Tak et al., 2006). Dpbl1 contains four
BRCT domains, which often function as motifs for the binding of phosphopeptides
(Glover et al., 2004). Sld2 interaction was mapped to the C-terminal pair of the four
BRCT domains in Dpbl1 (Tak et al., 2006).
In budding yeast, SId3 forms a complex with Cdc45 and their recruitment to
origins is mutually dependent (Figure 4A, B) (Kamimura et al., 2001). SId3 is
phosphorylated by CDK at three sites, and this promotes its association with the Nterminal pair of BRCT domains in Dpbl 1 (Tanaka et al., 2007; Zegerman and Diffley,
2007). Like Sid2, this interaction is essential for replication. Furthermore, the role of CDK
activity in replication can be bypassed by using phospho-mimetic mutations in Sid2
together with a fusion of SId3 to Dpbl 1, suggesting that SId2 and SId3 are the minimal
CDK targets required for replication initiation (Zegerman and Diffley, 2007).
The mechanism by which these proteins activate DNA replication has not been
elucidated. By treating cells with a cross-linking reagent it was found that GINS, DNA
polymerase e, Dpbl 1 and phosphorylated Sld2 could all form a complex as cells enter S
phase, dependent on CDK activity but not DDK activity (Muramatsu et al., 2010). This
complex has been called the preloading complex. Based on these and previous findings,
a model has been proposed in which Sld3-Cdc45 complex is recruited to origins and
helps to recruit the pre-loading complex through direct interaction of SId3 with Dpbl1 in
a CDK dependent manner (Figure 4C). GINS binds Mcm2-7 and, together with Cdc45,
activate the helicase. Sid2, Sld3 and DpB1 1 are released from origins through an
unknown mechanism while the other factors become part of the replisome (Figure 4D).
Conservation in metazoans
It seems likely that the mechanism by which DDK promotes activation of the
Mcm2-7 helicase and initiation of replication will be fundamentally similar between yeast
and other eukaryotes. There is evidence indicating that the N-termini of Mcm2 and
Mcm4 are phosphorylated by DDK in human cells, consistent with studies of their yeast
counterparts (Masai et al., 2000; Masai et al., 2006; Montagnoli et al., 2006).
Additionally, DDK and CDK activity is required for formation of the Cdc45-Mcm2-7-GINS
helicase complex in human cells (Im et al., 2009). However, the major CDK targets
during replication remain elusive in higher eukaryotes. This may be in part because, until
recently, orthologs of SId2 and Sld3 had not been identified in metazoans. The primary
sequence of SId3, DpB1 1 and Sld2 have diverged greatly during evolution even between
budding yeast species, making the search for orthologs much more difficult than for
many other replication factors. Orthologs of Dpb11 have been identified in metazoans
(TopBP1/Mus101/Cut5), all containing BRCT domains although the number of repeats
varies between species (Garcia et al., 2005). Importantly, all orthologs are required for
the initiation of DNA replication in the organisms examined (Garcia et al., 2005).
An ortholog of Sld2 has been recently identified in higher eukaryotes, the
RecQL4 DNA helicase (also called RecQ4) that is mutated in the human genetic disease
Rothmund-Thomson syndrome, which predisposes patients to cancer (Sangrithi et al.,
2005). Mutations causing human Rothmund-Thomson syndrome often cause truncations
that leave the N-terminal portion of the protein intact. Strikingly, this part of the protein
shows homology with yeast Sld2, although it is relatively limited (Sangrithi et al., 2005).
Depletion of RecQL4 in Xenopus egg extracts blocks chromosome replication and this
can be rescued by the N-terminus portion carrying homology with Sld2 (Matsuno et al.,
2006). Moreover, the Xenopus RecQL4 N-terminus binds Cut5/TopBP1 (ortholog of
Dpbl 1), suggesting that RecQL4 has a related role to yeast SId2 during the initiation of
replication (Matsuno et al., 2006).
Subsequent studies revealed that there are clear differences between RecQL4
and Sld2, as well as human and Xenopus RecQL4. Although Xenopus RecQL4 is
phosphorylated by CDK in vitro, the binding to Cut5/TopBP1 can occur even after
dephosphorylation of RecQL4, suggesting that, unlike SId2's interaction with Dpbl 1,
phosphorylation is not absolutely required for the interaction. RecQL4 also seems to be
acting at a later step, the recruitment of DNA polymerase a. However, similarly to yeast
SId2, human RecQL4 is required for assembly of the Cdc45-Mcm2-7-GINS complex and
is not required for elongation (Im et al., 2009; Xu et al., 2009). The reasons for the
differences between Xenopus and human RecQL4 are unclear, but these studies
suggest that the precise mechanism of action of RecQL4 in replication initiation has
diverged among different organisms.
An ortholog of Sid3 remains elusive. Inthis thesis I will describe the identification
of a metazoan protein that has the core characteristics of SId3. It associates with
TopBP1 and is essential for initiation of chromosome replication, specifically for pre-IC
formation. We propose that this protein, TICRR, is the metazoan ortholog of the Sid3
protein.
A
DDK
SId3-Cdc45
CDK@
Dpbll, Sld2, Po E,I GINS
Dpbll, Sld2, Sld3
C
D
Figure 4. Recruitment of additional proteins to origins leads to formation of the
pre-IC and subsequent activation of the MCM2-7 helicase.
(A) Sld3 and Cdc45 associate with the pre-RC in a mutually dependent manner. (B) A complex
consisting of Dpb 11, Sld2, DNA polymerase c (pol e) and GINS is assembled in a CDK
dependent manner and subsequently recruited to origins through a CDK-dependent interaction
between Sld3 and Dpbl1. (D) Dpbll, Sld2 and Sld3 are released through an unknown
mechanism while Cdc45 and GINS form a stable complex with MCM2-7, activating the helicase.
(Adapted from Araki, H 2010)
Mechanism for prevention of re-replication
Eukaryotes have evolved multiple mechanisms to ensure that replication is
initiated from each origin only once during each cell cycle (Arias and Walter, 2007;
Diffley, 2004). The temporal separation of events (i.e. MCM loading during G1 phase
and activation during S phase) is key for preventing re-replication. Thus, replication
initiation is tightly coordinated with cell cycle progression, whose major feature is the
oscillation of CDK activity. Pre-RC formation can only occur during G1, when CDK
activity is low. The ubiquitin ligase APC/C plays a major role in ensuring low CDK activity
during G1 and promoting pre-RC formation (Wasch and Engelbert, 2005). The APC/C is
active from mid mitosis to the end of GI and mediates proteolysis of mitotic cyclins and
the accumulation of CKIs throughout G1 phase, suppressing CDK activity. In addition, it
promotes degradation of the metazoan-specific inhibitor of Cdtl, Geminin (McGarry and
Kirschner, 1998). APC/C also regulates DDK activity by targeting the kinase regulatoy
subunit, Dbf4, for proteasomal degradation during G1 (Masai and Arai, 2002). Therefore,
the APC/C is responsible for inactivation of CDK and DDK as cells enter G1 and setting
up the right environment for pre-RC formation.
All additional mechanisms that prevent re-replication inhibit the first step of
replication initiation, pre-RC assembly. Once MCM2-7 is loaded, ORC, Cdc6 and Cdtl
are not required for initiating replication indicating that their primary role is to deliver
MCM2-7 to origins (Duncker et al., 1999; Hua and Newport, 1998; Rowles et al., 1999).
This feature allows for the inactivation of these pre-RC components outside of G1
phase, without compromising initiation of replication from already licensed origins.
In the budding yeast, all mechanisms known to date to ensure that DNA is
replicated only once per cell cycle depend on CDK activity. CDK phosphorylates all preRC components and these modifications have distinct consequences on each target.
CDK phosphorylates two ORC subunits, Orc2 and Orc6 (Nguyen et al., 2001). In
addition, direct binding of the S phase cyclin, Clb5, to an RXL motif in Orc6 also
contributes to preventing rereplication (Wilmes et al., 2004). Recently, it was shown that
CDK phosphorylation of Orc2 and Orc6 and possibly the interaction of Orc6 and Clb5,
block Cdtl binding to Orc6, preventing MCM2-7 helicase loading (Chen and Bell, 2011).
Cdc6 is inhibited by CDK in three ways: phosphorylation of Swi5 (Cdc6 transcriptional
activator) prevents Cdc6 nuclear import, Cdc6 phosphorylation promotes its proteolysis,
and direct association with mitotic CDK inhibits its association with origins (Mimura et al.,
2004; Moll et al., 1991). CDK also promotes the nuclear export of Mcm2-7, leading to its
exclusion from the nucleus in G2 and M phases (Nguyen et al., 2000).
Other factors are involved in regulating the inhibition of pre-RC formation in
metazoans and CDK plays a less important role. Cdtl seems to be the major target for
inhibition. Cdtl destruction by Cul4-Ddbl-Cdt2 ubiquitin ligase during S phase is a highly
conserved mechanism (Arias and Walter, 2007). Another well-conserved mechanism
among metazoans is the inhibition of Cdtl by Geminin, a protein that is cell cycle
regulated. Geminin was originally identified in Xenopus in a screen for APC/C substrates
(McGarry and Kirschner, 1998). It is destroyed during mid-mitosis to G1 and reaccumulates during late G1 when APC is inactivated. Geminin binds Cdtl, inhibiting
MCM2-7 binding to Cdtl and subsequent loading of the MCM2-7 complex onto origins
(Fujita, 2006). In mammals, Cdtl is targeted by the SCF E3 ubiquitin ligase for
proteolysis in a CDK dependent manner (Fujita, 2006).
Although the mechanisms by which ORC is inhibited are fundamentally
conserved, there are notable differences from organism to organism (DePamphilis,
2005). In general, chromatin binding is regulated and one or more ORC subunit
dissociate from chromatin after pre-RC assembly. Like in yeast, CDK phosphorylation
plays an important role in regulating ORC activity. It has been shown that mitotic Cdk1
phosphorylates one or more ORC subunits, and this blocks pre-RC assembly until
mitosis is complete. Finally, in flies and mammals, Orc1 is targeted for proteolysis. Cdc6
and Mcm2-7 do not seem to be major targets of negative regulation in metazoans (Arias
and Walter, 2007).
Cell cycle checkpoints
The integrity of genomic DNA can be extensively altered by errors occurring
during DNA replication and by reacting with molecules in normal cellular environments,
such as reactive oxygen and nitrogen species. In addition, cells are constantly exposed
to DNA damage caused by environmental sources such as UV and ionizing radiation as
wells as a myriad of chemical agents. (Jackson and Bartek, 2009). The repair of this
damage is essential for the survival and proper function of all eukaryotic cells.
Eukaryotic cells have developed several mechanisms to counteract the
potentially deleterious effects of DNA damage, which are collectively known as the DNA
damage response. Inthe following section I will focus on one mechanism, the cell cycle
checkpoints. Checkpoints comprise a complex network of pathways that cause arrest or
delay of cell cycle progression after DNA damage, allowing additional time for repair to
take place before the cell enters the next phase of the cell cycle (Lukas et al., 2004).
When damage is irreparable, those signaling pathways can permanently prevent
proliferation of damaged cells by mediating apoptosis or senescence. These control
mechanisms are crucial for the maintenance of genomic integrity because they minimize
the risk of DNA lesions to be converted into inheritable mutations. Defects in the DNA
damage checkpoints lead to accumulation of mutations and chromosomal aberrations,
which in turn increase the probability of developing genetic disorders and diseases
including cancer (Jackson and Bartek, 2009).
Checkpoints can be described as signal transduction systems consisting of
sensors that detect DNA lesions, mediators that amplify the signal and transducer and
effector kinases that transmit the signal to downstream targets (Figure 5) (Harper and
Elledge, 2007). DNA damage-induced checkpoints have been identified at the G1/S and
G2/M boundaries as well as during S phase of the cell cycle. Many DNA damage
sensors and signal transducers are integral components of all three checkpoints. ATM
and ATR are two protein kinases of the phosphatidylinositol 3-kinase (P13K)-like family
that are critical for the initiation of the signaling cascade in response to DNA damage
(Figure 5). ATM and ATR have similar structures; they both contain FAT and FATC
domains and a catalytic P13K domain. In addition, they preferentially phosphorylate
serine or threonine residues followed by glutamine. There are also clear differences
between these kinases; ATM is activated primarily by double strand breaks, while ATR
responds to a broader range of DNA lesions and stalled replication forks (Shiloh, 2003).
Transgenic mice lacking ATM are viable, while loss of ATR causes early embryonic
lethality, suggesting that ATR plays essential roles during normal cell cycle regulation
(Brown and Baltimore, 2000; de Klein et al., 2000; Elson et al., 1996).
A group of proteins, including Claspin, 53BP1, Mdcl, and BRCA1, mediate the
transmission of checkpoint signaling to two major downstream targets, Chk1 and Chk2
kinases. Activation of Chk1 is ATR-dependent, while Chk2 is an important target of ATM
(Harper and Elledge, 2007). The final targets of the ATR/Chk1 and ATM/Chk2 pathways
depend on the cell cycle phase at which damage is sensed and include components of
the cell cycle, apoptotic, and DNA repair machinery (Figure 5). Importantly, there is
considerable redundancy amongst pathways comprising the eukaryotic DNA damage
checkpoints and a simple linear pathway targeting a single substrate can rarely be
identified. The growing number of cell cycle and DNA repair regulators that are modified
in response to checkpoint activation underscores the complexity of this mechanism and
how numerous proteins work in concert to successfully execute the checkpoint response
(Matsuoka et al., 2007). In the following sections, I will provide an overview of the
mammalian DNA damage checkpoints by outlining the general principles of the key
signaling pathways orchestrated by ATR and ATM. In addition, I will review how the final
response is determined depending on the cell cycle phase at which damage is detected
by these signaling pathways.
Genotoxic
stress
/)00000000000000OO\
Sensor proteins
Transducers
ATR and ATM
Effectors
Chk1 and Chk2
Cell cycle
arrest
Senescence
DNA repair
Apoptosis
Figure 5. General scheme of the signal transduction pathways that are activated in
response to DNA damage or stalled replication forks.
DNA damage or stalled replication forks are sensed by a group of proteins, leading to the
activation of the ATM and ATR kinases which in turn activate the Chk1 and Chk2 kinases. ATM
and ATR function in combination with Chk1 and Chk2 to phosphorylate a number of targets
leading to a variety of cellular outcomes. (Adapted from Kastan MB and Bartek J 2004).
A TM and A TR: initiators of the signaling cascade in response to DNA
damage
ATM activation
ATM was identified as the product of the gene mutated in the autosomal
recessive disorder Ataxia telangiectasia, characterized by cerebellar degeneration,
immunodeficiency, chromosomal instability and radiation sensitivity (Savitsky K 1995). In
mammalian cells, ATM has critical roles in cellular responses to double strand breaks
(DSBs), which can arise from exposure to ionizing radiation and genotoxic chemicals, as
well as from cellular processes such as DNA replication, meiosis and V(D)J
recombination (Sun et al., 1989; Ward, 1988). Early studies on ATM indicated that its
kinase activity is induced very rapidly in response to DSBs, suggesting it acts at an early
stage of the signal transduction cascade (Banin et al., 1998; Canman et al., 1998).
Experimental evidence indicates that without cellular stress, ATM forms dimers
that are catalytic inactive. Very rapidly after DSBs have occurred, one ATM molecule
phosphorylates residue Serine 1981 on an interacting ATM molecule, leading to
disruption of the complex and release of active ATM monomers (Bakkenist and Kastan,
2003). Two additional autophosphorylation sites, S367 and S1893, are also important for
ATM activation in response to DSBs (Kozlov et al., 2006). Interestingly, this mechanism
of ATM activation appears to be less important for the activity of mouse ATM, where
mutation of the S1987 site (equivalent to S1981 in the human protein) does not affect
ATM activity (Pellegrini et al., 2006).
It is not clear what signal triggers the
autophosphorylation of ATM, but there is evidence suggesting that such signal could be
the result of changes in higher-order chromatin structure caused by DSBs (Bakkenist
and Kastan, 2003; Kim et al., 2009; Kruhlak et al., 2006). In addition, several proteins
are required for ATM activity (as discussed below) and may play a direct role in inducing
autophosphorylation and subsequent activation of ATM.
Much attention has been focused on understanding the early events that take
place between DSB formation and ATM activation. One of the hallmarks of the response
to DSBs is the rapid accumulation of several checkpoint signaling and repair proteins at
the location of the lesion to form distinct foci. The recruitment of proteins to DSBs is
thought to facilitate repair of the lesion and amplification of the checkpoint signal. In
support of this concept, localization of checkpoint signaling components to chromatin is
sufficient to trigger activation of the checkpoint in the absence of DNA damage
(Soutoglou and Misteli, 2008). Imaging technology in live mammalian cells has
elucidated the temporal order in which the initial proteins are recruited to DSBs (Lukas et
al., 2005). Mrell, Rad50 and Nbs1, components of the MRN complex, are the first
proteins to bind to the sites of DSB formation (Lisby et al., 2004; Lou et al., 2006; Lukas
et al., 2004) and thus have been implicated in the initial detection of DSBs and
transduction of the DNA damage signal. Their binding is independent of all proteins that
have been tested, including ATM. Consistent with the notion that MRN acts as a sensor
of DSBs, it was shown that the MRN complex mediates recruitment of ATM to sites of
DSBs and stimulates ATM kinase activity (Lee and Paull, 2004). The recruitment of ATM
occurs through a protein-protein interaction between the MRN component Nbsl and
ATM (Lee and Paull, 2004; You et al., 2005).
Another key event in the response to DSBs is the phosphorylation of the H2A
histone variant H2AX by ATM (Burma et al., 2001). H2AX is phosporylated very rapidly
after DSB formation, and form foci at the sites of DSBs that can be detected within 1 min
after exposure to ionizing radiation (IR) (Rogakou et al., 1999). Phosphorylation of H2AX
provides a high-affinity binding site for the mediator protein MDC1, which in turn
orchestrates the recruitment of downstream factors to the sites of DSBs (Stucki et al.,
2005). 53BP1, a protein proposed to function as a co-activator of ATM, is one of the
factors
recruited
to
foci.
Depletion
of 53BP1
by
siRNA
decreases ATM
autophosphorylation in response to IR, as well as the phosphorylation of ATM substrates
(DiTullio et al., 2002; Wang et al., 2002). The mechanism by which 53BP1 contributes to
ATM activity is unclear.
Numerous ATM substrates have been identified with functions in a wide range of
processes (Kastan and Lim, 2000; Matsuoka et al., 2007). Chk2 is an effector kinases
rapidly phosphorylated by ATM in response to DNA damage (Ahn et al., 2000; Matsuoka
et al., 1998; Matsuoka et al., 2000). Chk2 is not immobilized at sites of DNA damage
after IR treatment, suggesting that it acts as a signal distributor (Lukas et al., 2003).
Functionally, Chk2 can activate both apoptosis and cell cycle arrest pathways.
ATR activation
ATR is activated in response to a number of lesions, including stalled replication
forks, UV-induced dimmers, and double strand breaks. Activation of ATR requires its
association with ATRIP, TopBP1, the Rad17-RFC2-5 complex and the Rad9-Radl-Husl
(9-1-1) complex (Flynn and Zou). ATR forms a stable complex with ATRIP, which
regulates the localization of ATR to sites of replication stress and DNA damage and is
essential for ATR signaling (Cortez et al., 2001). Single stranded DNA is the primary
trigger for ATR activation. When single stranded DNA is generated, it is coated by
replication protein A (RPA) (Wold and Kelly, 1988). ATR is recruited to ssDNA via its
stable partner ATRIP, which binds to RPA-coated ssDNA (Namiki and Zou, 2006; Zou
and Elledge, 2003). The Rad17-RFC2-5 complex is independently recruited to ssDNA
(Zou et al., 2002). The presence of a dsDNA-ssDNA junction stimulates this complex to
load a second complex to the sites of DNA damage, the Rad9-Radl-Husl (9-1-1)
complex (Zou et al., 2003). Recruitment of the 9-1-1 complex by Rad17 enables
substrate selection by ATR.
TopBP1 is a BRCT-domain containing protein that functions in replication
initiation. TopBp1 is also recruited to the sites of DNA damage, where it directly
stimulates ATR kinase activity. A region sufficient for ATR activation, called the ATR
activation domain (AAD), has been identified between BRCT domains VI and VII of the
TopBP1 protein (Kumagai et al., 2006). Studies in human cells and Xenopus egg
extracts showed that TopBP1 interacts constitutively with Rad9 and identified a
phosphorylation site in the C-terminus of Rad9 that is required for the interaction (Lee et
al., 2007; Makiniemi et al., 2001; St Onge et al., 2003). The region of TopBP1 that binds
to Rad9 has been mapped to the first two BRCT domains (Lee et al., 2007). The
functional significance of this interaction appears to be to recruit TopBP1 to sites of
damage, where ATR has been independently recruited, to induce activation of ATR (Lee
and Dunphy, 2010).
Chk1 is the major downstream effector kinase of ATR, which propagates and
amplifies the checkpoint signal (Guo et al., 2000; Liu et al., 2000; Zhao and PiwnicaWorms, 2001). A major role of Chk1 is to inhibit the cell cycle phosphatases Cdc25A,
Cdc25B and Cdc25C (Sanchez et al., 1997; Sorensen et al., 2003; Zhao et al., 2002)
thereby preventing them from promoting progression through S phase or entry into
mitosis (Figure 6).
The G1/S checkpoint
CDK2, which is essential for entry into S-phase, is the key target of the G1/S
DNA damage checkpoint. The rapid induction of the checkpoint is mediated by the
inhibition of Cdc25A, a phosphatase that activates CDK2. Chk1/Chk2 phosphorylation of
Cdc25A promotes its degradation through the ubiquitin-proteasome system in the
presence of DNA damage (Donzelli and Draetta, 2003). A p53-dependent pathway can
also arrest cell cycle progression at the G1/S transition. p53 is phosphorylated by
ATR/ATM and Chk1/Chk2 in response to DNA damage, causing its dissociation from
Mdm2, an E3 ubiquitin ligase that targets p53 for degradation (Figure 6) (Wahl and Carr,
2001). p53 upregulates the transcription of proapoptotic and cell cycle inhibiting genes
such as the CDK inhibitor p21, which inhibits CDK2 (Figure 6). The response caused by
this pathway takes longer to occur because it requires the transcription and
accumulation of newly synthesized proteins (Bartek and Lukas, 2001). Another
mechanism to arrest the cell cycle at the G1/S transition is the DNA damage-induced
degradation of CDT1, a licensing factor essential for the initiation of replication. It has
been shown that CDT1 also undergoes ubiquitination and proteolysis in response to
DNA damage (Higa et al., 2003; Hu et al., 2004).
DNA-damage induced G2/M arrest
The G2/M checkpoint can be divided into two phases known to control mitotic
entry upon DNA damage. The early phase is ATM dependent, transient, and is reflected
by an abrupt reduction in mitotic index. This early G2/M checkpoint is activated within
one hour after damage and causes the arrest of cells that were in G2 at the time of
damage. The arrest results from phosphorylation of Cdc25C, a phosphatase required for
activation of CDK1, the mitotic CDK. Cdc25C phosphorylation leads to its sequestration
from the nucleus by 14-3-3. The late phase is ATM independent and reflected by an
accumulation of cells in G2 (Xu et al., 2002). It causes the arrest of cells that had been in
earlier phases of the cell cycle at the time of damage, but had continued progressing to
the G2 phase. The p53 pathway is known to contribute to the late phase of the G2/M
checkpoint through transcriptional activation of p21 and 14-3-30, which results in
increased inactivation and cytoplasmic sequestration of CDK1 (Figure 6) (Bunz et al.,
1998; Hermeking et al., 1997).
DNA Damage/Replication Stress
Figure 6. A Simplified scheme of
DNA damage-induced cell cycle
arrest and apoptosis.
ATM/ATR
Chk1/Chk2
I
MDM2
N
CDC25 14-3-3
p53
CDK activity is inhibited by DNA
damage checkpoints in several ways
including
inhibition
of
Cdc25
phosphatases and activation of p21. p5 3
controls the transcription of genes
involved in cell death and cell cycle
arrest. (Adapted from Kastan MB and
Bartek J 2004).
p21
CDKs
Apoptosis
Cell cycle progression
Checkpoints during S-phase
The genome is particularly susceptible to mutation during S phase, where
replication errors or replication stress can result in the stalling of replication forks,
replication fork collapse and subsequent DNA damage that is potentially deleterious to
the cell. Thus, monitoring DNA replication is essential for maintaining the integrity of the
genome. Two checkpoints have been described during S phase, the intra-S phase and
the S/M checkpoint (Bartek et al., 2004). The intra-S checkpoint prevents origin firing in
the presence of DNA damage or stalled replication forks, leading to a slowing in S-phase
progression. In addition, it is important for stabilizing stalled forks, allowing proper
resumption of DNA synthesis once the damage is repaired. The S/M checkpoint
monitors DNA replication and prevents mitotic entry until the genome is fully replicated.
These two responses converge at the same effector protein kinases and a clear
distinction is often difficult to make.
The intra-S phase checkpoint
ATR plays a particularly important role in S phase checkpoints in response to
DNA damage. Collapsed or stalled replication forks in S phase activate ATR, leading to
phosphorylation of Chk1 and the subsequent inhibition of Cdc25A phosphatase activity.
This prevents initiation of new replication origins, causing a reversible delay in cell cycle
progression (Bartek and Lukas, 2001). There is a second branch of the intra-S phase
checkpoint involving SMC1, NBS1 and BRCA1 and FANCD2. However, the specific
roles of these proteins remain unclear.
The S/M checkpoint
The S/M checkpoint is a surveillance mechanism that prevents initiation of
mitosis until the genome is fully and accurately replicated. In the absence of a functional
S/M checkpoint, cells enter mitosis prematurely and chromosomal fragmentation is
prone to occur. Mammalian cell fusion studies showing that fusion of a S-phase cell with
a G2-phase cell inhibits mitosis demonstrated the existence of such a checkpoint (Rao
and Johnson, 1970). Mitosis in the absence of DNA synthesis is observed when
mammalian cells are treated with caffeine, an inhibitor of the ATM and ATR kinases
(Brinkley et al., 1988; Schlegel and Pardee, 1986). These studies suggested that
regulators maintain the normal order of cell cycle events by detecting the completion of
DNA synthesis. Accumulating evidence suggests that the ATR-Chk1 pathway may fulfill
this function.
A role for ATR in normal cell cycle progression was first implied by the fact that
ATR and other components of the ATR signaling pathway, such as Chk1, are essential
for mammalian development and viability (Brown and Baltimore, 2000; Takai et al.,
2000). Mouse blastocysts lacking ATR accumulate chromosome breaks, a phenotype
attributed to loss of the S/M checkpoint (Brown and Baltimore, 2000). Furthermore,
several components of the ATR pathway associate with replicating chromatin during Sphase in the absence of exogenous DNA damage (Dart et al., 2004; Hekmat-Nejad et
al., 2000). Activation of CDK1-cyclin B is the key event required for mitosis to initiate.
Chk1 has been shown to localize to interphase centrosomes but not mitotic centrosomes
during unperturbed cell cycle progression, preventing premature activation of CDK1cyclin B by negatively regulating Cdc25B (Kramer et al., 2004).
There is also evidence for a role of Chk1 during an unperturbed S-phase.
Specifically, it is required for maintaining replication fork stability and regulates activation
of late origin firing by phosphorylating Cdc25A in the absence of DNA damage.
(Sorensen et al., 2003; Zhao et al., 2002). Depletion of Chk1, Rad9, Hus1, ATR or
Claspin in human cells leads to an accumulation of Cdc25 in the absence of external
DNA damage (Sorensen et al., 2004) suggesting that these proteins are involved in the
regulation of Cdc25A during replication. Furthermore, it was shown that inhibition of ATR
and Chk1 increases origin firing and loading of the replication factor Cdc45 in
undamaged cells (Syljuasen et al., 2005). Together, these studies provide evidence
supporting a model in which the ATR-Chk1 pathway is active at a low level during
normal S-phase due to intrinsic lesions arising from DNA replication and activation of
checkpoints then further activates the pathway. In addition, the ATR-Chk1 pathway
maintains DNA replication through structurally unfavorable stretches of DNA, termed
fragile sites in mammalian cells (Casper et al., 2002; Cha and Kleckner, 2002).
The zebrafish as a model system for the study of cell
cycle regulation
In the early 1970's, Dr. George Strisinger identified the zebrafish as a model
system for the study of vertebrate biology (Grunwald and Eisen, 2002). For the following
decades, zebrafish studies focused on developmental biology largely due to the
advantages that this organism offers. Zebrafish have a short generation time, around 34 months, and large numbers can be housed in a small space at relatively low cost. In
addition, fertilization of oocytes occurs externally, permitting easy manipulation of the
embryos. There are two particular characteristics that make the zebrafish a very
attractive model for developmental studies: the embryos (100-200 per clutch) are
transparent, and their development is very rapid. In fact, cleavage divisions, gastrulation,
morphogenesis and organogenesis all occur within 24 hours (Driever et al., 1996;
Kimmel et al., 1995). Therefore, scientists could follow in real time the behavior of single
cells or group of cells in the embryo at different stages of development.
Soon it became evident that the zebrafish represented a powerful tool for
studying genes that regulate specific developmental processes and large-scale screens
were developed. In particular, two groups used the methylating agent ethylnitrosourea
(ENU)
as a mutagen and isolated mutants with morphological developmental
phenotypes (Driever et al., 1996; Haffter et al., 1996). The advantage of this process is
that many mutants can be isolated in a relatively fast manner, but the cloning of the
affected genes is quite laborious. High throughput gene and EST mapping projects
revealed extensive conserved syntety between the zebrafish and human genomes and
assisted in the identification of mutated genes of interest by positional cloning (Barbazuk
et al., 2000). Nowadays, chemical mutagenesis is widely used for performing screens
and isolating mutants with phenotypes of interest.
The Hopkins lab developed an alternative approach for large-scale screens using
a retrovirus as an insertional mutagen. Although this method is less efficient, its
advantage relies on providing a feasible way of identifying the mutated gene due to the
insertion of a tag at the site of the lesion. In this screen, 525 embryonic lethal mutant
zebrafish were isolated, that represent lesions in 390 different cell essential genes. It
was estimated that this screen unveiled approximately 25% of all genes required for
embryonic and early larva development (Amsterdam et al., 2004). These mutants can be
classified into two categories based on their gross morphological phenotypes; one class
of mutants have developmentally specific phenotypes, whereas the other class display
non-specific phenotypes such as extensive cell death or general growth defects. The
latter class of mutants includes housekeeping genes such as genes involved in
transcription, RNA processing and translation. Mutants for known cell cycle regulators,
such as polo-like kinase, Cyclin B1 and Aurora B kinase, also fall into this category
(Amsterdam et al., 2004).
Importantly, 20% of the genes were novel or poorly characterized at the time
they were cloned (Amsterdam et al., 2004). We predicted that among these novel genes
were regulators of the cell cycle. Our lab designed an assay for screening the zebrafish
insertional mutant library for a defect in the ionizing radiation induced checkpoint
response in order to identify novel components required for the DNA damage response.
In the following chapters, I will describe how such a screen was performed, leading to
the discovery of a novel protein, Ticrr (for TopBP1 -interacting, checkpoint and replication
regulator). I will also describe how the analysis of the defects in ticrr zebrafish mutants
gave insight into the function of this protein, which was later complemented with the
characterization of the human ortholog.
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CHAPTER II: A zebrafish screen for novel DNA
damage response genes identifies TICRR
Nelly M. Cruz, Cristopher L. Sansam and Jacqueline A. Lees
Experimental contributions:
C.L.S. designed the screen and conducted the experiments in Figure 2, 6, 7 and 8. N.C.
conducted the large scale zebrafish screen.
Summary
Eukaryotes have evolved numerous checkpoint pathways to protect genomic
integrity during normal cell division and in response to DNA damage. Mutations in DNA
damage checkpoint components often lead to genetic disorders and cancer in humans.
We have screened a collection of zebrafish insertional mutants to identify new
components of these checkpoints. Specifically, we assayed activation of the IR-induced
G2/M checkpoint by examining the mitotic index of zebrafish embryos after IR treatment.
We have identified 12 zebrafish mutant lines that showed persistence of mitotic cells
after IR treatment. 5 of these also had aberrant mitotic index without IR treatment,
suggesting that the disrupted genes in these zebrafish lines play a role in cell cycle
progression. One of the mutant zebrafish that had a defect in checkpoint activation
carries an insertion in a previously uncharacterized gene that we have named ticrr. The
ticrr zebrafish gene encodes a 220 kDa protein with no identifiable domains that could
suggest its biochemical function. We have found orthologs of the Ticrr protein in other
species, from primitive metazoans to humans. Characterization of Ticrr will further our
understanding of the process of cell cycle regulation and conservation of genome
integrity in response to DNA damage.
Introduction
Upon sensing DNA damage, eukaryotic cells activate a number of signal
transduction pathways collectively known as the DNA damage response. Activation of
these pathways leads to cell cycle arrest, DNA repair and, when the damage is
irreparable, initiation of apoptosis. Three molecularly distinct DNA damage checkpoints
induce cell cycle arrest at the G1/S and G2/M boundaries, or a delay in S phase
progression, in the presence of DNA damage (Lukas et al., 2004). Many DNA damage
sensors and signal transducers are shared by all three checkpoints. ATR and ATM are
two protein kinases of the phosphatidylinositol-3 (PI-3) kinase-like family that mediate
the initiation of checkpoints in response to DNA damage. ATM responds to double
strand breaks (Lee and Paull, 2004), whereas ATR responds more broadly to DNA
damage and replication interference (Shiotani and Zou, 2009). ATR and ATM kinases
phosphorylate a number of targets that propagate and amplify the signal, including the
effector kinases Chk1 and Chk2.
Checkpoints trigger cell cycle arrest by regulating CDK activity. Specific CDKcyclin combinations regulate cell cycle progression at different stages of the cell cycle,
and they are inhibited after DNA damage by various mechanisms. A key mechanism is
the inhibition of the Cdc25 phophatases (CDKs activators) (Donzelli and Draetta, 2003).
Additionally, checkpoints activate DNA repair proteins and cause stabilization of the
transcription factor p53, which upregulates the transcription of pro-apoptotic and cell
cycle inhibiting genes such as BAX and the CDK inhibitor p21 (Giono and Manfredi,
2006).
The importance of the DNA damage response in human pathology is well
established. Defects in the DNA damage checkpoints lead to accumulation of mutations
and chromosomal aberrations, which in turn increase the probability of developing
genetic diseases. In fact, mutations in various genes encoding DNA damage checkpoint
proteins are associated with a variety of human disorders and these are often
characterized by an increased predisposition to cancer (Jackson and Bartek, 2009).
Thus, considerable attention has been given to the dissection of components of the DNA
damage response and their function. Despite great advances that have been made in
recent years, it has become clear that the DNA damage response is a complex network
of interacting signaling pathways with many key players, some of which have not been
identified (Harper and Elledge, 2007). The zebrafish is as an excellent model in which to
conduct genetic screens for vertebrate cell cycle and checkpoint regulators. This is
primarily due to its small size and fecundity but also because maternally contributed
mRNAs allow embryos to survive to developmental stages at which defects in cellessential genes can be assayed.
Through a pilot genetic screen, our lab has validated our ability to identify novel
cell cycle regulators using zebrafish (Sansam et al., 2006). In this prior study, mitotic
index was assayed through whole mount staining of zebrafish embryos for
phosphorylated (Ser 10) histone H3 (pH3). We established that the number of pH3positive cells decreases rapidly when zebrafish embryos are exposed to 15 Gy of
ionizing radiation, showing that the G2/M checkpoint is intact in these embryos. We used
this assay to perform a large-scale screen of the Hopkins' collection for genes required
for the IR-induced G2/M checkpoint. In addition, the pH3 assay facilitated the detection
of mutants that have a defect in mitotic progression. Here, we summarize the screen
results and highlight candidates of interest for both phenotypes. Of particular interest are
the hi1573 and hi3202A mutants, which had robust accumulation of pH3 positive cells
after IR treatment. Each of these lines carries a mutation in an uncharacterized gene
that we have named ticrr (the origin of this name will be discussed in Chapter 3). We
show that the insertions are linked to the observed phenotype and the accumulation of
54
pH3 positive cells is caused by a checkpoint defect and not a mitotic defect. This study
further confirms the potential of zebrafish for identification of novel checkpoints and cell
cycle regulators.
Results
A genetic screen for zebrafish mutants with a defective IR response
We have designed and validated a screen for the discovery of novel DNA
damage response genes in zebrafish (Sansam et al., 2006). Using an antibody against
serine-10 of phosphorylated Histone H3, a marker of cells in mitosis (Crosio et al.,
2002), we established that zebrafish embyos activate a G2 checkpoint in response to
ionizing radiation (IR). Whole mount staining of wild-type zebrafish embryos at 32 hours
post-fertilization (hpf) with anti-pH3 revealed that approximately 3% of the cells are in
mitosis at this developmental stage (Sansam et al., 2006). Within one hour of exposure
to ionizing radiation (IR), the anti-pH3 positive cells are largely absent in wild-type
embryos, indicating that there is a robust DNA damage-induced cell cycle arrest in
zebrafish embryonic cells (Figure 1A). Based on cell cycle kinetics experiments, we
established that G2 lasts more than one hour at this developmental stage (Sansam et
al., 2006). To asses the transit of cells from S phase into G2/M, embryos were pulsed
with BrdU and chased for varying times after the BrdU pulse, then fixed and stained for
both BrdU and pH3. The rate at which BrdU-positive cells entered mitosis reflects the
duration of S and G2. Using this assay, it was determined that the average duration of
G2 in wild-type embryos is between 1 and 2 h (Sansam et al., 2006). Therefore, the
decrease in mitotic index one hour after IR exposure represents a G2 checkpoint and
not a G1 or S phase checkpoint.
In human cells, a rapidly activated checkpoint requires ATM kinase. To
determine if the zebrafish G2/M checkpoint is also dependent on ATMIATR, embryos
were treated with caffeine, an ATM/ATR inhibitor, before irradiating and staining them
with anti-pH3 antibody. The caffeine-treated embryos maintained a similar level of pH3positive cells to the unirradiated controls, indicating that caffeine prevented activation of
the checkpoint (Figure 1A) (Sansam et al., 2006). This indicates that ATM/ATR kinases
are required for activating this checkpoint, and more importantly, we can identify
embryos with an abrogated checkpoint by examining pH3 staining after exposure to IR.
b
Figure 1. Zebrafish can be used to screen for IR-induced G2 checkpoint response
genes.
(A) Mitotic cells were labeled in 32hpf embryos using an anti-pH3 antibody in untreated (-IR),
IR-treated (+IR) and caffeine pretreated/IR-treated (+IR; +Caffeine) wild-type zebrafish embryos.
(B) Analysis of pH3 showed a low mitotic index in wild-type embryos but not dtl mutant
clutchmates I h after IR exposure, which is indicative of a G2 checkpoint defect. Adapted from
Sansam, C.L, 2006.
The Hopkins laboratory generated a zebrafish collection of approximately 500
heterozygote lines carrying insertional mutations in over 300 different genes required for
embryonic development (Amsterdam et al., 2004).The sensitivity of the pH3 assay was
verified by staining known checkpoint and cell cycle mutants in the Hopkins collection,
and a pilot screen was performed on several mutants, including some with insertions in
novel genes. We were able to identify mutants that have an increase number of mitotic
cells after irradiation, compared to the non-irradiated embryos. We observed the mitosis
after irradiation (MAI) phenotype in fish carrying mutations in SMC1 and MCM7, proteins
known to be required in DNA damage checkpoints in mammalian cells, demonstrating
that the screen can be used to find mutants that fail to arrest after DNA damage. In
addition, zebrafish carrying an insertional mutation in the gene denticleless (dtl) showed
a defect in the DNA damage checkpoint (Figure 1B). In a prior study, our lab established
both the biological properties and mechanism of action of the human ortholog of the Dtl
protein. This analysis showed that Dtl is an essential component of the early G2/M
checkpoint and is required for normal cell cycle control, primarily to prevent re-replication
(Sansam et al., 2006).
We have now applied this screen to a large collection of zebrafish mutants that
carry stable viral insertions within 336 different genes. These lines are fully viable as
heterozygotes but the homozygous mutants display developmental defects 24-96 hours
post-fertilization (hpf) that are typically lethal (Amsterdam et al., 2004). For our cell cycle
screen, we intercrossed the heterozygous mutants, treated 50 or more of the resulting
embryos at 32 hpf with ionizing radiation (IR), and assayed pH3 staining 1 hour later
(Figure 2A). This timepoint in development was chosen for the screen for two reasons.
First, the number of pH3 positive cells in wild-type embryos is optimal during this time,
allowing easy detection of an increase or decrease in staining. Secondly, maternal
transcripts should be largely absent by this time, allowing the emergence of phenotypes
that otherwise could have been masked. Lines were considered to have altered mitotic
index if approximately one quarter of the embryos showed altered pH3 staining relative
to the rest of the clutch. For those lines that displayed an aberrant number of pH3
positive cells, the staining was repeated at least two more independent times in order to
confirm the phenotype. PCR genotyping of the embryos was used to confirm that the
phenotype was linked to the mutant insert.
A
Normal Response
Mitosis Phenglyoe
MAI Phenotyoe
±18
-IR
j32 hpf
15 Gy IR
-IR
-IR
15 hour
anti-pH3 stain
Figure 2. A zebrafish screen for novel DNA damage response and cell cycle
regulators.
(A) Scheme of checkpoint screen. Heterozygous carriers were intercrossed to yield clutches of
embryos that were expected to consist of 25% homozygous mutants. The clutches were irradiated
or left untreated and fixed one hour later, followed by staining with a pH3 antibody. (B) Possible
outcomes of the checkpoint assay. The pH3 staining of mutant zebrafish embryos can uncover
both a mitosis phenotype and a mitosis after irradiation (MAI) phenotype.
Using this approach, we identified 12 mutant lines that showed a high mitotic
index compared to their wild-type and heterozygous clutchmates after IR exposure.
These mutants can be classified into two classes (Figure 2B). The first class of mutants
also displayed a relatively high mitotic index without irradiation, which could be indicative
of a delay or arrest in mitosis and suggests that the affected gene is involved in mitotic
progression. Alternatively, it could indicate a higher rate of proliferation. In the second
. ...
....
. ...
class of mutants, the non-irradiated embryos did not display a difference in their mitotic
index. These are considered to be checkpoint activation failure mutants. The screen
results are summarized in Table 1.
Table 1. Insertional mutants from the Hopkins zebrafish collection that showed an
increase in pH3 staining after IR treatment.
Zebrafih
Gene
line
Gene
mutated
function
Casein
kinase
pH3 staining
pH3 staining
Increase
Increase
factor Increase
Increase
untreated
IR-treated
hi1002
1, alpha 1
Signal transduction
hi1487
Site-1 protease
Transcription
Regulator
hi2578
nopl 0
Ribosome biogenesis
Increase
Increase
hi2820
clasp1
Regulator of microtubule
Increase
dynamics
Increase
hi3512B
bc2
Unknown
Increase
Increase
hi923
v-ATPase
SDF/54 kD
Vacuolar proton pump
No difference
Increase
hi933
pbx4
Transcription factor
No difference
Increase
hi1573
hi3202A
5730590G19like
Unknown
No difference
Increase
hi1858A
CWC25-like
Unknown
No difference
Increase
hi2688
paics
de novo purine synthesis
No difference
Increase
hi3526B
gart
de novo purine synthesis
No difference
Increase
hi3685
hi3685
pG 1
uRNA
subunit G
Transcription
No difference
Increase
Insertionalmutations in zebrafish that affect normal mitotic progression
Many mutants in the collection have non-specific developmental phenotypes,
such as extensive cell death, small eyes and head, and deformed bodies (Amsterdam,
There are two classes of genes that could cause those phenotpyes when
2003).
mutated: housekeeping genes and genes involved in cell proliferation. Given that at the
time of the screen approximately 20% of the mutated genes were novel or poorly
described, we reasoned that novel cell cycle genes could also be identified in our
screen, specifically genes that are required for normal mitotic progression. Indeed, in our
pilot screen we observed that we could identify mutants for known mitotic proteins, such
as polo-like kinase and the kinesin-related motor protein Eg5, because they not only had
a high mitotic index after IR treatment, but also in the absence of exogenous DNA
damage.
By also scoring the mitotic index in unirradiated embryos, we identified several
zebrafish lines in which the homozygous mutants had higher pH3 staining compared to
wild-type and heterozygous clutchmates (Figure 3, data not shown). hi2820 mutants,
which carry an insertion in the gene encoding the evolutionary conserved protein
Clasp1, fall into this category. Even though the zebrafish Clasp1 protein has not been
characterized, studies with its human (CLASP1), Xenopus (Xorbit/CLASP), Drosophila
(MAST/Orbit), and C. elegans (CLS-2/ R106.7) homologs revealed that they all play key
roles during mitotic progression (Galjart, 2005). Thus, it is not surprising that disruption
of the zebrafish Clasp1 protein causes a defect in mitotic progression. The human
Clasp1 protein has a stabilizing function at the kinetochore, which is essential for the
bipolar alignment of chromosomes on the mitotic spindle (Maffini et al., 2009; Maiato et
al., 2003; Pereira et al., 2006).
hi1002 homozygous mutants also had a aberrantly higher number of pH3
positive cells (data not shown). These zebrafish are mutant for Casein kinase 1 isoform
alpha (CKla), a multifunctional Ser/Thr kinase. Depletion of the yeast homolog, HRR25,
causes defects in a number of processes including chromosomal segregation
(DeMaggio et al., 1992). Mammalian CKla localizes to mitotic spindles, suggesting that
its role in chromosomal segregation may be conserved in higher eukaryotes (Brockman
et al., 1992; Nousiainen et al., 2006). Moreover, one of the mammalian CKIa interacting
proteins is the regulator of chromosome condensation 1 (RCC1), which plays a role in
nuclear transport and mitotic spindle formation (Dubois et al., 2002). CKla has also
been found to act as a negative regulator of the Wnt and Hedgehog signaling pathways
(Price, 2006). Thus, the accumulation of pH3 positive cells in the ckla zebrafish mutants
could be caused by a block in mitosis as a result of a defect in chromosome segregation,
or by aberrant proliferation due to misregulation of the Wnt and Hedgehog pathways.
Additional experiments are necessary to distinguish between these two possibilities.
Lastly, hi3512B (mutant for Bc2), hi1487 (mutant for site-1 protease, Figure 3)
and hi2578 (mutant for Nop1O) also displayed a mitotic phenotype. A role in mitosis has
not been described for these proteins to this day. Interestingly, two independent studies
found that human BC2 localizes to the midbody of dividing cells (Morita et al., 2007) and
to the mitotic spindle (Nousiainen et al., 2006), which is highly suggestive of a role for
BC2 during mitosis.
(1)1
CL
>t
+IR
Figure 3. hi1487 homozygous mutants show an aberrant accumulation of pH3
positive cells in the absence of exogenous DNA damage.
Mitotic cells were labeled in 32hpf embryos using an anti-pH3 antibody in untreated (-IR), IRtreated (+IR) hil487 wild-type and mutant clutchmates. Untreated mutant embryos show an
increase in pH3 positive cells, indicative of a defect in mitosis prgression.
Disruption of de novo purine nucleotide synthesis in zebrafish abrogates
IR-induced cell cycle arrest
Several mutants had a higher pH3 staining only after IR treatment, suggesting
these mutants fail to undergo an IR-induced G2 arrest (Table 1). A striking observation is
that two of these mutants, hi3526B and hi2688, carry insertions in genes that encode for
enzymes involved in de novo purine synthesis. Specifically, hi3526B carries a mutation
in the gart gene, while hi2688 carries a mutation in the paics gene. These genes encode
two multifunctional enzymes that catalyze several steps of inosine monophosphate
(IMP) synthesis. The synthesis of purine nucleotides is important for many processes:
nucleotides are constituents of the genetic material, cofactors in enzymatic reactions,
and the major energy source that drives most cellular reactions.
Ribonucleotide depletion and/ or slowing replication forks activates a p53
mediated checkpoint response independently of direct DNA damage, which leads to G1
and G2 arrest (Linke et al., 1996). Interestingly, this response seems to be abrogated by
inhibition of de novo purine synthesis pathway. Bronder and Moran showed that
inhibition of GART activity in human cell lines leads to nuclear accumulation of p53;
however, the transcription of the downstream targets p21, mdm2, and bax, was impaired
(Bronder and Moran, 2003). Consequently, cells progress through G1 and G2, and
accumulate in S phase due to decreased ribonucleotide pools that impair DNA
synthesis. This effect is specific for inhibition of de novo purine synthesis, since inhibition
of de novo pyrimidine synthesis does not impair the G1 and G2 arrest (Bronder and
Moran, 2003). Our checkpoint assay specifically surveys the early G2 checkpoint, which
is thought to be p53 independent (Taylor and Stark, 2001). It is unknown if inhibition of
de novo purine synthesis will also have an effect on p53-independent checkpoint
activation in response to exogenous DNA damage. However, it is intriguing that two lines
in our collection carrying mutations in genes required for pyrimidine nucleotide synthesis,
hi3510 and hi2694, were not scored as having a checkpoint defect in our screen (data
not shown). hi3510 is mutated for thymidylate synthase, whereas hi2694 carries a
mutation in the trifunctional protein CAD, which contains the three first enzyme activities
of de novo pyrimidine synthesis. This suggests that the mitosis after irradiation
phenotype observed in hi3526B and hi2688 is specifically caused by disruption of the de
novo purine synthesis and not by decreasing nucleotide pools in general.
An insertional mutation in a novel gene that abrogates IR-induced cell
cycle arrest in the zebrafish
It is important to note that most of the DNA damage response candidates
identified in this screen had a subtle phenotype; the mutants showed a slight, albeit
consistent, increase in the pH3 staining after IRtreatment (Figure 4). The reason for the
modest phenotype may be the existence of compensatory mechanisms (see
discussion). One of the mutants that scored as having a modest defect was hi3685,
mutated for RNA polymerase 11, subunit G. The role of RNA polymerase 11 in
transcription has been widely described. Recently, a similar screen for G2-M checkpoint
genes in Drosophila also identified several subunits of RNA polymerase 11as being
required for proper checkpoint response (Kondo and Perrimon, 2011). The role of this
protein in checkpoints could be indirect; perhaps it is required for the transcription of
other checkpoint proteins. The mutants that showed only a slight increase in pH3
staining were not further pursued in this study.
Figure 4. Mitosis after irradiation
phenotype in hi923 mutants.
0.
Analysis of pH3 staining showed a slight
increase in the mitotic index in IR-treated
+IR
hi923
homozygous
mutant
embryos
compared to wild-type clutchmates.
Notably, hi1573 homozygotes showed a robust increase in mitotic index,
compared to their wild-type and heterozygous clutchmates, after IR exposure (Figure 5).
In contrast, the mitotic index without radiation in the hi1573 mutants was
indistinguishable from that of the wild-type embryos (data not shown). This mutant was
not only identified as a potential candidate in the larger-scale screen of the mutant
collection, but was also scored as having a mitosis after irradiation phenotype in the
initial pilot screen. Because its phenotype was robust, we decided to focus our attention
on the further characterization of hi1573 mutant embryos.
wild-type clutchmate
mutant clutchmates
Figure 5. hi1573 and hi3202A zebrafish embryos show a Mitosis After Irradiation
(MAI) phenotype.
Whole mount immunostaining of pH3 showed that IR treatment induces the appropriate block to
mitotic entry in 36 hpf wild-type embryos, while hil573 and hi3202A homozygous mutant
clutchmates retained a high mitotic index.
We mapped the hi1573 viral insertion just upstream of an uncharacterized open
reading frame (5730590G19-like) on chromosome 25 (Figure 6A). A second line in the
collection, hi302OA, contains a distinct mutant allele of 5730590G19-like with a viral
insertion in the first predicted coding exon (Figure 6A). Notably, hi3202A homozygotes
have a normal mitotic index in the absence of irradiation (data not shown) and display
the MAI phenotype (Figure 5B) just like the hi1573 mutants. The hi1573 and hi3202A
homozygotes also have identical developmental phenotypes: at 36 hpf these lines
develop a dark head that is characteristic of widespread apoptosis in the central nervous
system (Amsterdam et al., 2004). We confirmed that cells in these mutants are
apoptotic, with increased acridine orange and anti-cleaved caspase-3 staining appearing
at 26 hpf (data not shown). Typically, the retroviral insertions in the Hopkins mutant
collection cause a decrease in mRNA expression that results in recessive, loss-offunction phenotypes. We performed quantitative RT-PCR to measure the expression of
5730590G19-like at 36 hpf in wild-type and mutant embryos and found that both the
hi1573 and hi3202A insertions cause a strong reduction in mRNA levels of this gene
(Figures 6B-C). Taken together, these data indicate that disruption of the novel gene
5730590G19-like causes embryonic lethality and the mitosis after irradiation phenotype.
We have named this gene ticrr for reasons that will be described in the following chapter.
Hereafter, I will refer to this gene as ticrr.
66
m
m
I
UUIU
Y~Tuuyiyi
,uu~
hi1573
hi202A
Exon 1
Exon 2
Wild-type Littermates Mutants
~C
08 -t
ta 0.6
.2 0.4
Z0.2
X0
wt
mut
Figure 6. The checkpoint defect results from disruption of a novel gene, ticrr.
(A) Schematic of the zebrafish ticrr gene denoting the position of viral insertions in the hil573
and hi3202A mutant lines. (B-C) Quantification of total mRNA by RT-PCR showed that the
levels of ticrr mRNA were greatly reduced in both the (B) hil573 and (C)hi3202A homozygous
mutants at 36 hpf.
A critical step in regulating entry into mitosis is the activation of Cdc2 kinase by
Cdc25 phosphatases. In the presence of DNA damage, Cdc25 phosphatases are
inhibited, thereby preventing Cdc2 kinase activation. We sought to determine whether
the increased mitotic index in the ticrr mutants after DNA damage was associated with
high Cdc2 kinase activity. Thus, we performed an in vitro assay using Cdc2-cyclin B1
complexes immunoprecipitated from lysates of 40 hpf embryos that had been irradiated
or left untreated. Embryos were identified as either mutant or wild-type based on the
presence or absence of the characteristic ticrr developmental phenotype. IRexposure of
wild-type embryos resulted in a 14-fold decrease in the phosphorylation of a synthetic
peptide substrate for Cdc2 (Figure 7). This is consistent with the decrease in mitotic
index and the activation of the G2/M checkpoint in response to IR. In contrast, the
activity of Cyclin B1-Cdc2 remained high after DNA damage in the ticrr mutants (Figure
7), consistent with the persistence of mitotic cells.
cu
6c 0.-6
-IR
CD
CN
1
M*+IR
0.4 -
S0.2
CU
Wt
CdC2 0
mut
&Q4
Figure 7. Cdc2 kinase activity remains high
in ticrr mutants after IR treatment.
Cyclin B1-Cdc2 was immunoprecipitated from 40
hpf unirradiated (-IR) and irradiated (+IR) pools of
wild-type (+/+ and +/-) and mutant (-/-) hil573
embryos. The total levels of Cdc2 were determined
by western blotting (representative experiment
shown). Assessment of cyclin B1-Cdc2 kinase
activity (mean ± SD; n=3 biological replicates)
showed that the hil573 mutants retain high activity
after IR treatment.
We reasoned that the mitosis after irradiation phenotype and the high Cdc2
kinase activity could arise in two possible ways: the mutation could abrogate the
checkpoint that prevents mitotic entry after DNA damage or it could reflect a failure to
efficiently exit mitosis. To distinguish between these two possibilities, we performed a
nocodazole trapping experiment using 36 hpf embryos. For this assay, nocodazole was
added immediately after exposure to IR and then the embryos were incubated for 2
hours before disaggregating and quantifying the percentage of pH3-positive cells with
4N DNA content by FACS. Since nocodazole causes early to mid-mitotic arrest, an
accumulation of mitotic cells following IR and nocodazole treatment would indicate that
cells continue to enter mitosis and thus a defect in the G2/M checkpoint. In the absence
of irradiation, the level of 4N, pH3-positive cells was increased in the nocodazoletreated, versus the untreated, embryos for both the wild-type (3.4-fold) and ticrr mutants
(2-fold; Figure 8). IR treatment greatly reduced the level of mitotic cells in the wild-type
embryos in both the absence and presence of nocodazole (Figure 8). This is consistent
with the existence of a robust G2/M checkpoint. In contrast, the level of 4N, pH3-positive
cells in the ticrr mutants was completely unaffected by IR treatment. Indeed, we still
observed the 2-fold increase in mitotic cells that results from nocodazole treatment.
Taken together, our data show that the G2/M checkpoint is abrogated in the ticrr mutants
and these cells continue to enter mitosis following DNA damage.
4-
-IR
3 -
+IR
Co
I1
noc.
+
-
wt
-
+
mut
Figure 8. Cells in ticrr homozygous mutants continue to enter mitosis after IR
treatment.
Unirradiated and irradiated 36 hpf wild-type (+/+ and +/-) and hil573 mutant (-/-) clutchmates
were maintained in the absence or presence of nocodazole for 2 hrs. The percentage of pH3positive cells was quantified by FACS (mean ± SD; n = 20,000 cells counted for each of three
biological replicates).
The zebrafish ticrr gene is predicted to encode a 1,824 residues protein
(NM_001003887.1) with an anticipated molecular mass of 202 kDa. Genes encoding
proteins with significant homology exist in other vertebrates, including humans, mouse
and Xenopus (Table 2). We also found putative Ticrr orthologs in other metazoans
including the primitive chordates Branchiostoma floridae and Ciona intestinalis, the
echinoderm Strongylocentrotus purpuratus, the arthropod Ixodes scapularis, the
nematode Caenorhabditis elegans, the cnidarian Nematostella vectensis, and the
placozoan Trichoplax adhaerens. The conservation across these proteins maps to
specific stretches, yielding high confidence in its significance, thoug the overall
conservation is poor (Table 2). We were unable to detect putative Ticrr orthologs in
Drosophila and there was no significant homology to any yeast protein. We have
uncovered additional roles for this protein, which will be described in the following
chapter.
Table 2. Conservation between D. rerio Ticrr and orthologs identified by psiBLAST
ClustalW
siBLAST
Species
0. rerio
.sapiens
M.musculus
G.gallus
.laevis
.floridae
C.intestinalis
S. purpuratus
.scapularis
C.elegans
N. vectensis
T. adhaerens
Accession #
Length
(amino acids)
NP_001003887.1*
1824
NP689472.3
NP 084111.1
XP413862.2
AAH73061.1
XP002215127.1
XP002120860.1
XP792617.1
XP002416131.1
NP491752.1
XP001628252.1
XP_002111061.1
1910
1889
1679 (1937**)
1394
1911
1441
2305 (1768*)
1495
919
1845
1069
HSP
Length
1126
1114
1113
1098
1147
1172
654
1111
472
1125
515
Results
%
%
Identity Positives
35
35
34
36
22
19
22
20
21
22
18
52
52
50
54
38
34
37
35
37
37
37
%
Gaps
Alignment
%
Identity
8
7
13
6
12
16
16
19
15
14
17
30
28
29
30
16
14
11
16
11
14
14
*Sequence used in Psi-BLAST was truncated at amino acid 1181 before a long stretch of low
complexity proline-rich sequence.
** Length of predicted XI and Sp proteins from manually refined gene models.
HSP=high scoring segment pairs.
Discussion
To maintain their genome integrity, cells have a complex network of pathways
that together compose the DNA damage response. Loss of components of the DNA
damage checkpoints often leads to genetic disorders and cancer in humans. We have
used the zebrafish to identify new components of these checkpoints. Along the way, we
have also identified genes that are potentially required for mitotic progression. The
uncovering of these genes raises two important questions: first, what are the
mechanisms by which these mutations lead to higher mitotic index in the zebrafish, and
second, are these genes involved in similar processes in other vertebrates?
What role do the identifiedgenes play in normal cell cycle progression?
We observed that a group of the mutants identified in the screen have an
aberrant number of pH3 positive cells in the absence of exogenous DNA damage.
Histone H3 is phosphorylated on serine 10 during late G2 and becomes
dephosphorylated during anaphase, in mitosis (Crosio et al., 2002). We speculate that
the accumulation of pH3 positive cells in these mutants could be a result of the inability
of those cells to progress through G2/M and exit mitosis. Entry into and progression
through mitosis requires precise coordination and many proteins are involved in the
process. Thus, this phenotype could be caused by disruption of many pathways that
regulate G2/M transition, chromosome segregation and mitotic exit.
The other class of mutants described in this study accumulates pH3 positive cells
only after IRtreatment. This phenotype has been shown to be characteristic of disruption
of the G2/M checkpoint (Sansam et al., 2006; this study). Many checkpoint genes are
also important for normal cell cycle progression. The mutants in the Hopkins collection
are embryonic lethal, so it is highly probable that the candidates we have identified also
have a role in normal cell cycle progression. Defects in normal cell cycle progression
could be uncovered by analyzing cell cycle kinetics in the mutant embryos. Further
characterization of the cell cycle phenotypes of the mutants identified will help uncover
the specific role of the mutated proteins in mitotic progression and/or cell cycle
regulation.
What role do the identifiedgenes play in the DNA damage response?
Two mutants in the collection with disrupted de novo purine synthesis had a
slight increase in the number of pH3 positive cells after IR treatment. It is plausible that
the impaired IR-induced checkpoint in these zebrafish mutants is caused solely due to
the reduction of dNTP pools and the mutated proteins do not have a bona fide
checkpoint role. It has been shown that inhibition of de novo purine synthesis causes
rapid depletion of GTP and ATP pools. The fact that disruption of the pyrimidine
synthesis pathway does not impair the checkpoint response suggests that this effect
may be a consequence of depletion of the ATP pools specifically. ATP is used as a
substrate by kinases to phosphorylate proteins, a modification that checkpoints highly
rely on for signal transmission (Huen and Chen, 2008). In accordance with this
hypothesis, p53 is stabilized in response to de novo purine inhibition but is not
phosphorylated or acetylated in a number of residues that are important for its function
as a transcriptional activator (Bronder and Moran, 2003). Another process important for
proper checkpoint function is the ubiquitylation/proteosomal degradation of certain key
players (Bassermann and Pagano; Huen and Chen, 2008), a process dependent on
ATP. It remains an open question if cells impaired for purine synthesis are unable to
arrest due to an indirect effect of the limited nucleotide pools, or whether these enzymes
play a direct role in checkpoint activation that has not been described before.
We have uncovered a number of candidates that have an IR-induced G2/M
checkpoint defect in zebrafish. The difference in pH3 positive cells for most of the
mutants identified was small, but highly reproducible. It is possible that the mutated
genes have a minor role or are indirectly involved in the checkpoint. In addition, they
could be important players of the G2/M checkpoint but other pathways compensate for
their depletion. Indeed, activation and maintenance of the G2 arrest requires targeting of
multiple regulatory processes involved in the normal progression of the cell from G2 into
M phase. Several checkpoint pathways act in parallel and converge at specific nodes.
An example of this is the regulation of the mitotic kinase Cdc2-CyclinB, which is inhibited
by several mechanisms in response to DNA damage (Stark and Taylor, 2006).
Recently, a similar screen for finding novel G2/M checkpoint components was
performed in Drosophila (Kondo and Perrimon). For this genome-wide screen,
knockdown of individual genes was achieved using RNA interference technology and
phosphohistone H3 staining was used to assay mitotic index in cells treated with the
anticancer drug doxorubicin, which induces double-stranded breaks. This study found
that knockdown of known checkpoint proteins, like ATR, only caused a modest
checkpoint defect. Furthermore, the authors were able to show that when ATR
knockdown is combined with knockdown of Nbs1, another checkpoint protein, the
checkpoint defect is stronger. This is an example of the synergy that exists for
checkpoint activation. Thus, the defects in checkpoint activation observed in some
zebrafish mutants in my screen could be weak due to compensation.
Nevertheless, our screen has been successful in identifying two mutants in the
same gene with a robust increase in mitotic index after IRtreatment, suggesting that this
protein is a major player in checkpoint activation. These mutations disrupt a novel gene
that we have named ticrr. We have confirmed that the aberrant accumulation of pH3
cells in the ticrr mutants is caused by a failure to activate the G2/M checkpoint in
response to IRtreatment.
In addition to the G2/M checkpoint, DNA damage response checkpoints have
been identified at the G1/S transition and during S phase of the cell cycle. After DNA
damage, these checkpoints block progression through the cell cycle until the damage is
repaired. However, they are mechanistically different and, while some components are
shared amongst the three checkpoints, others are checkpoint-specific. Thus, the
identified genes could be specifically required for the early G2/M checkpoint or they
could have a more general role. In addition, specific types of damage can induce certain
pathways of the DNA damage response. It may be informative to examine how these
mutant embryos respond to other types of damage, including damage caused by UV
radiation and chemotherapeutic agents like camptothecin and doxorubicin, which target
DNA topoisomerase I and II, respectively. A more extensive analysis of the mutant
zebrafish will be crucial for understanding the role of these proteins in DNA damage
checkpoints and cell cycle regulation.
Are the identified genes required for normal cell cycle progression and
checkpointsin other vertebrates?
An advantage of performing screens for DNA damage response genes in
zebrafish is that it is relatively straightforward to identify candidate vertebrate orthologs
based on protein sequence comparison. This screen has identified a novel gene, ticrr, as
a checkpoint regulator. We have identified orthologs of this gene in a number of
metazoans, including humans. A particularly useful system for studying cell cycle and
DNA damage regulators is human cells, where a wide array of tools is available. The
following chapter will describe how we have taken advantage of these tools to gain
insight into the molecular basis for Ticrr's role in cell cycle control. Further studies with
the orthologs on Ticrr will establish the conservation, and therefore importance, of Ticrr's
biological functions.
74
In summary, we have shown the power of the zebrafish to uncover genes
required for cell cycle progression and checkpoints. Future characterization of the
proteins identified in this screen will lead to a better understanding of the pathways that
underlie the DNA damage response and will significantly advance our ability to unravel
the complex processes maintaining the integrity of the genome.
Materials and methods
Zebrafish maintenance
Zebrafish were maintained as previously described (Amsterdam et al., 2004). For the
screen, heterozygous insertion carriers were intercrossed and at 24 hpf the embryos
were manually dechorionated and 1-phenyl-2-thiourea (PTU,
0.003%) added to
suppress pigmentation. At 32 hpf, 60 embryos from each clutch were subjected to G2/M
checkpoint analysis.
G2/M checkpointassays
To initiate a DNA damage response, zebrafish embryos were exposed to 15 Gy IR from
a 6Co source and analyzed for pH3 staining one hour later as previously described
(Sansam et al., 2006). To test whether caffeine could inhibit the zebrafish G2
checkpoint, 32-hpf embryos were placed in 2 mM caffeine and incubated for 30 min at
28.50C before the IR/pH3 assay. For the nocodazole trapping experiment, embryos were
exposed to IR, immediately placed in nocodazole (150 ng/mI + 1% DMSO), and
incubated at 28.5C for 2 hours before analysis of pH3/DNA content by FACS.
Genotyping and expression analysis of zebrafish lines
Primers for genotyping were as follows:
hil 573 - forward primer (F): AAGCAAGCTACATCTCAAAGCA;
hil 573 -reverse primer (R): CGGAAAACCCTGAAGTGTGAT;
hi3202A - (F) CAGATCCCCTGGTTATAAGTGTTGC;
hi3202A - (R) CAGTTATGGCCTCTCAGA ATGTCGT;
reverse primer for provirus: GCTAGCTTGCCAAACCTACAGGT.
Ticrr mRNA levels in hil 573 were measured by two-step real time RT-PCR with SYBR
Green I Dye (Applied Biosystems) using the primers F: CTGAACAGTTTGCAT
GGATGG and R: CTTTCGGCTGTATGTCCTGCT and normalized against zebrafish bactin (F: CATCAGCATGGCTTCTGCTCTGTATGG
and R: GACTTGTCAGTGTACA
GAGACACCCTG). Ticrr mRNA levels in hi3202A were analyzed by limiting dilution RTPCR using the Ticrr primers above and RPL35 as the reference mRNA (F:
GCTGCTTCCAAGCTC TCAAAAATCC and R: TGCCTTGACGGCGAACTTGCGAATG).
In Vitro Kinase Assay
50 embryos for each data point were treated with 15 Gy IR, and incubated for 30
minutes at 28.5*C. Embryos were then dechorionated, deyolked and homogenized in
400pl KLB (50mM Tris pH 7.4, 150mM NaCl, 0.1% Triton X-100, 4mM EDTA, 0.1% NP40, 50mM NaF, 0.2mM Na3VO 4, 1OuM Leupeptin, 5ug/ml Aprotonin, 1mM PMSF). The
homogenate was adjusted to 1 mg/ml prior to precipitation with anti-Cyclin B1. The
beads were washed twice with KLB, and KAB (50mM Tris pH 8.0, 10mM MgC 2 , 1mM
EGTA) and incubated for 15 minutes at 300C in 20pl of kinase assay mix [KAB with
40pM cold ATP; 100pM peptide (HATPPKKKRK);
(6000Ci/mmol)].
1mM DTT; 0.5pCi/ul y-32P-ATP
Substrate phosphorylation was quantified by filter binding and
scintillation counting.
FACS Analysis
Mutant embryos were identified by either developmental phenotype or PCR genotyping
of a fraction of individual, fixed embryos. Cells were disaggregated by triturating
embryos in 0.25% Trypsin/1 mM EDTA using a p200 and fixed in 70% ethanol at -200C
overnight. Suspensions of cells from 20 wild-type or mutant embryos were pooled and
prepared for pH3/propidium iodide FACS analysis as described (Pozarowski and
Darzynkiewicz, 2004). FACS analysis was conducted by FACScan (Becton-Dickinson).
DNA content was quantified by ModFit LT (Verity Software), and pH3 and BrdU was
quantified by FlowJo (Tree Star, Inc.).
Acknowledgements
I would like to thank Chris Sansam for use of his previously published data,
included in Figure 1. I would also like to thank Kate Anderson for maintenance of the
mutant lines of fish, and Tim Angelini and Sam Farrington for maintenance of the
zebrafish colony.
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CHAPTER III: TICRR associates with TopBP1 and
is required for replication initiation
Nelly M. Cruz, Cristopher L. Sansam, Paul S. Danielian, Adam
Amsterdam, Melissa L. Lau, Nancy Hopkins and Jacqueline A. Lees
The material presented in this chapter was adapted, with permission, from the following
publication:
*Sansam, C.L., *Cruz, N.M., Danielian, P.S., Amsterdam, A., Lau, M.L., Hopkins, N., and
Lees, J.A. (2010). A vertebrate gene, ticrr, is an essential checkpoint and replication
regulator. Genes Dev 24, 183-194.
*These authors contributed equally to this work.
Experimental contributions:
C.L.S. conducted the experiment showed in Figures 1, 2 and 3. P.D. aided with the
cloning of TopBP1 deletion mutants used in Figure 7. N. Cruz performed the
experiments for Figures 4, 5, 7, 8 and 9.
Summary
Eukaryotes have numerous checkpoint pathways to protect genome fidelity
during normal cell division and in response to DNA damage. Through a screen for G2/M
checkpoint regulators in zebrafish, we identified ticrr, a previously uncharacterized gene
that is required to prevent mitotic entry after treatment with ionizing radiation. ticrrdeficiency is embryonic lethal in the absence of exogenous DNA damage because it is
essential for normal cell cycle progression. Specifically, the loss of ticrr impairs DNA
replication and disrupts the S-M checkpoint, leading to premature mitotic entry of cells
with partially replicated genomes and mitotic catastrophe. Here, we identify the human
TICRR ortholog and show that it associates with TopBP1, a known checkpoint protein
and a core component of the DNA replication pre-initiation complex (pre-IC). The
TICRR-TopBP1 association is stable without chromatin and requires BRCT motifs
essential for TopBP1's replication and checkpoint functions. We hypothesized that
TICRR acts together with TopBP1 in pre-IC formation, in an analogous manner to the
yeast protein Sld3. Consistent with this model, we show that ticrr-deficiency disrupts
chromatin binding of pre-IC, but not pre- RC, components. Taken together, our data
show that TICRR acts in association with TopBP1 and plays a dual role in pre-IC
formation and checkpoint response.
Introduction
Eukaryotic cells possess numerous mechanisms to ensure genome fidelity. In
dividing cells, DNA replication is the primary potential source of errors. To ensure that
DNA replication occurs at the appropriate cell cycle stage and only one copy of the
genome is synthesized, this process is divided into two temporally distinct steps (Bell
and Dutta 2002; Sclafani and Holzen 2007). During G1, the replicative helicase complex,
Mcm2-7, is loaded onto origin DNA by the ORC, Cdc6 and Cdtl proteins in a process
called pre-replication complex (pre-RC) formation. However, the loaded helicase is
inactive, and is only activated upon entry into S phase during pre-initiation complex (preIC) formation. This process requires CDK and DDK (Dbf4-dependent kinase) activity and
involves recruitment of additional proteins to the Mcm2-7 complex, including TopBP1,
Cdc45 and the GINS complex (Hashimoto and Takisawa 2003; Kubota et al. 2003;
Aparicio et al. 2009). Once activated, the helicase and its associated proteins recruit the
remaining DNA synthesis machinery leading to the formation of a pair of bidirectional
replisomes.
In addition to the core replication machinery, multiple checkpoint pathways exist
to protect cells from DNA damage arising from replication errors and/or genotoxins
(Bartek et al. 2004; Harper and Elledge 2007). Two of these pathways function during
DNA replication: the intra-S phase checkpoint stabilizes existing replication forks while
inhibiting firing of late origins, and the S/M checkpoint prevents the cell from prematurely
entering mitosis before it has fully replicated the genome (Bartek et al. 2004). Abrogation
of the latter pathway allows cells to enter mitosis with incompletely replicated
chromosomes, leading to chromosome fragmentation, segregation defects and, often,
mitotic catastrophe (Canman 2001). In G2, a checkpoint blocks mitotic entry when DNA
is damaged. This pathway is rapidly activated when cells are exposed to ultraviolet or
ionizing radiation, which primarily causes bulky DNA adducts or double-stranded DNA
breaks, respectively. Mutations that prevent cells from appropriately responding to DNA
damage cause human developmental disorders, cancer and aging (Harper and Elledge
2007). Importantly, nearly all cancer cells have partially impaired checkpoints and thus
checkpoint pathway components have emerged as important targets for anti-cancer
drugs.
Considerable attention has focused on identifying the proteins that contribute to
these checkpoint pathways. This has established the PIKK kinases, ATM and ATR, as
playing a central role in DNA damage response (Harrison and Haber 2006). In
vertebrates, ATM responds primarily to double-stranded breaks, while ATR is more
versatile, responding to a wide range of damage or replication stress (Brown and
Baltimore 2003). ATR is a key player in the S/M checkpoint and is also required for the
radiation-induced G2/M checkpoint (Nghiem et al. 2001). Once activated, ATR
phosphorylates and
activates the
Chk1
kinase.
Importantly,
activated
Chk1
phosphorylates and inhibits the Cdc25 phosphatases, thereby preventing activation of
the Cyclin/CDK kinases and blocking cell cycle progression.
Numerous other proteins have been identified as sensors and/or mediators in
checkpoint signaling pathways (Harrison and Haber 2006). Human TopBP1 and its
orthologs Cut5 (Xenopus laevis), Mus101 (Drosophila melanogaster), Cut5/Rad4
(Saccharomyces pombe) and Dpbl1 (Saccharomyces cerevisiae), are particularly
intriguing because they have been shown to be critical not only for checkpoint activation
in response to DNA damage, but also DNA replication initiation in unperturbed cells
(Garcia et al. 2005). Studies in numerous organisms established TopBP1 as essential
for the initiation of DNA replication. Consistent with these observations, TopBP1
associates with Cdc45 and the recruitment of TopBP1, Cdc45 and GINS, which is
required for the pre-RC to pre-IC transition, appears to be interdependent (Van Hatten et
al. 2002; Kubota et al. 2003; Takayama et al. 2003; Schmidt et al. 2008). SId3, a protein
that is essential for pre-IC formation in yeast, also associates with Dpbl 1TopBP1
However, to date Sid3 orthologs have not been identified in higher eukaryotes.
Importantly, TopBP1 has also been identified as being essential for both the
intra-S and S/M checkpoints in numerous organisms (Garcia et al. 2005). Although it is
formally plausible that these checkpoint functions are an indirect consequence of
TopBP1's replication function, this is not the case: studies with conditional and
separation of function mutants show that TopBP1's checkpoint function can be clearly
separated from its action in pre-IC formation (McFarlane et al. 1997; Saka et al. 1997;
Hashimoto et al. 2006; Yan et al. 2006). Finally, TopBP1 is also required for activation of
the G2/M checkpoint in response to DNA damaging agents (Garcia et al. 2005). The
widespread roles of TopBP1 are consistent with the broad spectrum of phenotypes
resulting from TopPB1 deficiency. For example, Drosophila mus101TopBP1 mutants
display defects in chorion gene amplification, hypersensitivity to DNA damaging agents
and mitotic chromosome instability (Yamamoto et al. 2000).
The predominant feature of the TopBP1 protein is that it contains multiple BRCT
motif repeats (four in yeast and eight in humans) (Garcia et al. 2005). These domains
commonly mediate protein-protein interactions, and in certain cases pairs of BRCT
domains act as phosphopeptide binding motifs. Structure-function studies have revealed
a critical role for the N-terminal BRCT domains in TopBP1 function (Garcia et al. 2005).
The BRCT domains in this part of the protein are conserved from yeast to humans.
Accordingly, this N-terminal half is both necessary and sufficient for DNA replication
(Hashimoto et al. 2006). BRCT motifs I and 11 are required for binding to CDK
phosphorylated Sld3 and also for TopBP1's checkpoint function in vertebrates (Lee et al.
2007; Tanaka et al. 2007; Zegerman and Diffley 2007; Yan and Michael 2009).
Additional BRCT domains in the carboxy-terminal half of TopBP1 of higher eukaryotes
are also involved in the response to DNA damage and replication stress. Specifically,
BRCTs VII and VIII are important for ATR-dependent phosphorylation of Chk1 in
response to replication stress in Xenopus extracts (Yan et al. 2006; Yan and Michael
2009), and BRCT domain V is required for TopBP1 to form nuclear foci in response to
damage or stalled replication forks (Yamane et al. 2002).
We have screened a mutant zebrafish collection for identifying novel DNA
damage response regulators. The screen led to the identification of a novel gene, ticrr,
required to activate the ionizing radiation induced G2/M checkpoint. Upon further
characterization, we have identified additional roles of this gene: it is required for Sphase progression and the S/M checkpoint. This spectrum of defects is highly
reminiscent of those arising in TopBP1 mutants. Accordingly, we show that TICRR binds
to TopBP1 in vivo and is essential for pre-IC formation in a similar manner to TopBP1. In
recognition of its broad spectrum of functions, we have named this gene ticrr for
TopBP1-interacting, checkpoint and replication regulator.
Results
Disruption of ticrr impairs S-phase progression and causes premature
chromatin condensation
DNA damage checkpoint genes are often required for normal cell cycle control.
For example, TopBP1 is required for both checkpoint activation in response to DNA
damage and for the initiation of DNA replication in undamaged cells (Garcia et al. 2005).
Consistent with this precedent, our analysis of the hi1573 and hi3202A zebrafish
mutants carrying insertion in the ticrr gene revealed that ticrr is required for cell cycle
control in the absence of DNA damage. FACS analysis of DNA content revealed that 40
hpf ticrr mutant embryos had a higher percentage of cells with between 2N and 4N DNA
content than the wild-type control, suggesting an accumulation of cells in S-phase
MMMP '_
Mir
(Figure 1A). To further explore this possibility, we compared the ability of wild-type and
ticrr mutant 40 hpf embryos to incorporate the nucleotide analog BrdU (Figure 1B). The
wild-type embryos had a high level of cells that had between 2N and 4N DNA content
and were BrdU-positive. This is indicative of the high rate of cell proliferation at this early
stage of zebrafish development. In contrast, the 40 hpf ticrr mutant embryos
incorporated very little BrdU even though a significant fraction of the cells had between
2N and 4N DNA content. This shows that the ticrr-deficient cells are impaired for DNA
replication.
A
B
wt
mut
G2/M
100.
S
75
U
wt
25
LGII
DNA Content (PI)
mut
BrdU+
D
25
20
T
15
10
DNA Content (PI)
Figure 1. Ticrr is required for normal DNA replication.
(A) Cell cycle profile from a representative pool of 40 hpf hil573 wild-type (+/+ and +/-) and
mutant (-/-) zebrafish embryos and quantification of cells with G1, S or G2/M DNA content
(mean ±SD, n=3 biological replicates) showed an increase in S-phase cells in the ticrr mutants.
(B) BrdU and PI FACS analysis of cells from pools of BrdU pulse labeled, wild-type and ticrr
mutant embryos. Quantification of the BrdU+ population (mean +SD, n=3 biological replicates)
showed a dramatic reduction in replicating DNA in ticrr mutants.
In addition to this replication defect, the 40 hpf ticrr mutants displayed defects in
mitotic progression. First, quantification of the percentage of cells in each phase of
mitosis by visual inspection of chromatin morphology of pH3-positive cells showed that
86
the distribution of these populations differed between wild-type and ticrr mutant zebrafish
(Figure 2A). Specifically, the ticrr mutants had a much higher percentage of cells that
appeared to be in prometaphase, as judged by the presence of condensed chromatin
that is not aligned on the metaphase plate. Moreover, they had a reduced percentage of
anaphase cells and, within this population, anaphase bridges were prevalent (Figure
2A). To explore this defect further, we analyzed chromosome spreads from colchicinearrested cells (Figure 2B). This revealed a low incidence of cells with highly fragmented,
condensed chromosomes in the ticrr mutant, but never wild-type, zebrafish (Figure 2B).
This level of chromosomal fragmentation is consistent with cells that have undergone
premature
chromatin condensation. Given their DNA replication problems, we
hypothesized that the ticrr mutants have an impaired ability to activate the S/M
checkpoint and thus enter mitosis with partially replicated DNA. To address this
possibility, we used FACS to determine the DNA content of pH3-positive cells (Figure
2C). It is well established that histone H3 phosphorylation begins in late G2, peaks
during metaphase and then declines through anaphase (Hendzel et al. 1997).
Accordingly, in the wild-type embryos, histone H3 phosphorylation is restricted to the
cells with 4N DNA content (Figure 2C). In contrast, we find that a high percentage of the
pH3-positive cells in the ticrr mutants have less than 4N DNA content. Taken together,
the cell cycle defects observed in 40 hpf ticrr mutants show that ticrr is required in the
absence of exogenous DNA damage for S-phase progression and also for activation of
the S/M checkpoint. We propose that failure of these two processes causes cells to
enter mitosis with partially replicated genomes, resulting in the chromosomal
abnormalities and loss of anaphase cells observed in the ticrr mutants.
Abnormal anaphase
100%
abnormal
anaphase
L anaphase
E metaphase
* prometaphase
* prophase
wt
mut
4
M<4N
3
*4N
r2
0
DNA Content (PI)
Figure 2. Ticrr is required for S/M checkpoint function.
(A) Quantification of the proportion of pH3 positive cells in various mitotic phases, based on
chromatin and spindle morphology, (mean +SD; n=100 cells counted in each of 4 wt and mut
embryos) established a defect in mitotic progression and the presence of abnormal anaphase cells.
Representative cells with anaphase bridges are shown. (B) Metaphase spreads of cells from
colchicine treated embryos showed mitotic cells with fragmented chromosomes. A representative
wild-type metaphase spread and examples of abnormal metaphase spreads from mutants are
shown. (C) FACS measurement of anti-pH3 and PI staining of cells from wild-type and mutant
embryos showed a population of mutant pH3-positive cells with less than 4N DNA content (blue)
in ticrrmutants, establishing entry into mitosis before completing DNA replication.
If ticrr mutants are defective in DNA replication and the S/M checkpoint, the DNA
replication defect should be the primary defect and the mitotic abnormalities a secondary
consequence. To test this, we turned to an earlier developmental timepoint when the
ticrr defects were just beginning to arise. Since we first detect apoptosis in the hi1573
mutants at 26 hpf, we used 24 hpf embryos to screen for cell cycle defects. At this
timepoint, the ticrr mutant embryos are morphologically indistinguishable from the wildtypes. Thus, ticrr mutants were identified by PCR genotyping a small fraction of cells
dissociated from individual embryos. The remaining cells from 20 embryos of each
genotype were then pooled for cell cycle analysis. FACS showed that there is already an
increase in the percentage of cells with between 2N and 4N DNA content in the 24 hpf
ticrr mutants compared to the wild-type controls, suggesting that cells are proceeding
slowly through S-phase (Figure 3A). Moreover, the ticrr mutants had a lower average
level of BrdU incorporation in these S-phase cells than the wild-types (Figure 3B). Thus,
the defect in DNA replication clearly precedes the initiation of apoptosis. Notably,
analysis of the DNA content of pH3-positive cells showed that, unlike the 40 hpf
embryos, pH3-positive cells with sub-4N DNA content are not observed in the 24 hpf ticrr
mutants indicating that there is no premature mitotic entry at this timepoint (Figure 3C).
These data show that the DNA replication defect is the earliest detectable phenotype
and thus is not a consequence of the premature mitotic entry.
....................
.......
....
.I.
100
wt
Li G2/M
MS
75
i50Ef
'~0
~25-U
BrdU-]
lG1
DNA Content (PI)
DNA Content (PI)
30
wt
mut
mut
4
3
*<4N
*4N
D15
.io
C
Wu
6-l
CL.
10
TO-
DNA Content (PI)
Figure 3. A DNA replication defect but not premature mitotic entry occurs in 24 hpf
ticrrmutants.
(A) Cell cycle profiles from 24 hpf hil573 wild-type (+/+) and mutant (-/-) zebrafish embryos
and quantitation of cells with G1, S or G2/M DNA content (mean ±SD, n=3 biological
replicates), showed an increase in S-phase cells in mutants. (B) BrdU and PI FACS analysis of
cells from pools of BrdU pulse labeled, 24 hpf wild-type and ticrr mutant embryos.
Quantification of the BrdU+ population showed no change in the percentage of cells replicating
DNA (mean ±SD, n=20,000 cells in each of 3 biological replicates), but the level of BrdU
incorporation per cell is significantly decreased in the mutants (mean BrdU signal = Mean[BrdU+
signal]/Mean[BrdU- signal]; Student's t-test p<0.05). (C) FACS measurement of anti-pH3 and PI
staining of cells from 24 hpf wild-type and mutant embryos showed no evidence of an S/M defect
in the ticrrmutants at this timepoint, as judged by the absence of pH3 positive cells with less than
4N DNA content.
TICRR binds TopBP1
The phenotypes of the ticrr mutant embryos are highly reminiscent of those
resulting from depletion of TopBP1/Dbpl1/Cut5/Mus101. Specifically, these deficiencies
have been shown to disrupt DNA replication, the S/M checkpoint and DNA damage
checkpoints (Garcia et al. 2005). This raised the possibility that Ticrr acts in the TopBP1
pathway. In particular, we wondered whether Ticrr might be the vertebrate ortholog of
90
the budding yeast protein, SId3, which interacts with Dpbl 1 and is required for DNA
replication initiation (Kamimura et al. 2001; Tanaka et al. 2007; Zegerman and Diffley
2007). To test this hypothesis, we examined whether the Ticrr protein shows any
sequence homology with Sid3, or any other protein that might yield insight into its
biochemical function. We found putative Ticrr orthologs in other metazoans; the
conservation across these proteins maps to specific stretches, yielding high confidence
in its significance, but the overall conservation is poor. There was no significant
homology with any yeast protein, including SId3. However, given how rapidly the
conservation falls off amongst the metazoan Ticrr proteins and also the poor sequence
conservation of SId3 across the fungal kingdom, it seemed plausible that Ticrr could be a
functional analog of Sld3, and yet bear no significant sequence similarity. Notably,
despite their large size, the Ticrr proteins have no known functional motifs that could
help to infer their biochemical activity.
In the absence of any functional insight from the protein sequence, we
investigated the biochemical properties of the Ticrr protein. For these studies, we
switched to human cells due to the availability of reagents for known replication and
checkpoint regulators. Human TICRR is an uncharacterized gene on chromosome 15
(C15orf42). The NCBI Reference Sequence database contained a predicted full-length
TICRR mRNA sequence (NM_152259.3), inferred from partial cDNA and genomic
sequences, and the encoded human TICRR protein is predicted to be 211 kDa. We
generated a full-length 5,753-bp TICRR open reading frame by amplifying two
overlapping cDNA fragments from HeLa cell mRNA. We also used a 30 kDa, carboxyterminal fragment of the TICRR protein to raise multiple polyclonal antisera in mouse
and rabbits. These successfully recognized overexpressed TICRR and also an
endogenous protein of approximately 250 kDa by both western blotting and
immunoprecipitation (Figure 4; data not shown). This band was verified to be TICRR by
partial knockdown using TICRR shRNAs (data not shown).
Having established that we can detect the endogenous TICRR protein, we
assessed its subcellular localization through biochemical fractionation of human cells.
TICRR was entirely recovered from nuclear extracts of asynchronously growing HeLaS3 cells and was not present in cytosolic extracts. Moreover, TICRR fractionated with
the insoluble nuclear material and was resistant to extraction from nuclei by low salt (150
mM NaCl) and nonionic detergent (1% NP-40) but could be extracted with high salt (0.3
M NaCl) or ionic detergents (0.1% SDS or 0.5% deoxycholate; Figure 4A and data not
shown). These results strongly suggested that TICRR associates with chromatin.
Dpbl
1 TopBP1
and Sid3 are both chromatin-associated, thus, we further tested whether
TICRR was bound to chromatin by treating the insoluble nuclear material with nuclease.
For this analysis, we used a component of the origin recognition complex, ORC2, as a
positive control. We found that treatment of the chromatin/nuclear matrix material with
micrococcal nuclease caused partial release of TICRR and this mirrored the level of
release of ORC2 (Figure 4B). These results demonstrate that a large fraction of TICRR
is associated with chromatin, as has been previously described for TopBP1 (Garcia et al.
2005).
92
MNase:
[Nac
-
M)
0
0
Figure 4. Human TICRR is a chromatin-associated protein.
(A) Nuclei preparations were subjected to sequential salt extractions with the indicated NaCl
concentrations. Orc2 is a chromatin-bound protein, while Lamin is structural component of the
nucleus. (B) Western blotting of biochemical fractions from HeLa cells showed that the majority
of TICRR is present in the chromatin-enriched fraction (chrom) and can be partially released by
micrococcal nuclease (Mnase) treatment. Orc2 and alpha-tubulin are chromatin and cytoplasmic
markers. WCE = whole cell extract, Si = soluble cytoplasmic fraction, S2 = soluble nuclear
fraction.
We next asked if the endogenous TICRR and TopBP1 proteins interacted. For
this analysis, proteins were extracted from nuclei using ionic detergent (0.1% Sodium
Dodecyl Sulfate) and then immunoprecipitated with either pre-immune or anti-TICRR
polyclonal rabbit antisera. Subsequent western blotting established that TopBP1 was
present in the anti-TICRR, but not in pre-immune, immunoprecipitates (Figure 5A).
Moreover, we were able to conduct a reciprocal IP/immunoblotting experiment to show
that TICRR co-immunoprecipitated with TopBP1 (Figure 5B). Since both TICRR and
TopBP1 are chromatin-associated proteins, it seemed possible that DNA bridged the
interaction between these two proteins. However, we were able to show that these two
proteins continued to co-IP even if the lysates were pre-treated with ethidium bromide or
DNase I (Figure 5A). Thus, we conclude that human TICRR and TopBP1 are associated
proteins and this interaction can occur in the absence of DNA.
93
.......
.....
..
+EtBr
Input
(5%) Ip.
EJL
+DNase I
B,
(ly
_
CO
"D
Figure 5. Human TICRR interacts with TopBP1.
(A) Immunoblotting showed TopBP1 coimmunoprecipitates with TICRR but not preimmune
(PI) antibodies. The association was not affected by treatment with ethidium bromide (EtBr) or
DNase I indicating its independence from chromatin binding. (B) Reciprocal IP-western blotting
confirmed that TICRR is present in TopBP1, but not IgG, immunoprecipitates.
Given the relatively poor sequence homology of TICRR across species, it was
important to determine whether the TopBP1 binding ability of human TICRR was
conserved in the zebrafish Ticrr protein. Since antibodies were not available for
zebrafish Ticrr, we cloned the full length zebrafish Ticrr coding sequence and overexpressed GFP-tagged versions of either zebrafish Ticrr (GFP-zTicrr) or human TICRR
(GFP-hTICRR) in human cells along with human TopBP1. Notably, human TopBP1 was
recovered with similar efficiency in GFP-zTicrr and GFP-hTICRR immunoprecipitates,
even though the GFP-zTicrr protein was poorly expressed (Figure 6). This cross-species
binding shows unequivocally that the TICRR-TopBP1 interaction is conserved between
human and zebrafish.
.......................
IP: a-GFP
GFP
Input
'Uzebrafish
9
r,
Figure 6. Human TopBP1 interacts with
TopBPI.
IP-western blotting showed that GFP-tagged
zebrafish Ticrr associated with human
TopBP1 when co-expressed in human cells.
The human TopBP1 is a large multifunctional protein whose predominant feature
is the presence of eight BRCT domains (Figure 7A). The yeast homologs of TopBP1,
Dpbl1 and Cut5, have only four BRCT domains that are highly conserved with BRCTs I,
11,IV and V of the vertebrate TopBP1 proteins (Hashimoto et al. 2006). Prior studies
have shown that individual or pairs of BRCT domains mediate interactions with specific
proteins, and various known TopBP1 functions have been mapped to specific BRCT
domains. The N-terminal half of X. laevis TopBP1, which includes the four highly
conserved BRCTs, is both necessary and sufficient for the initiation of DNA replication
(Hashimoto et al. 2006). Moreover, BRCT motifs I and 11appear to be particularly
important in both replication and checkpoint functions: these are required for Dpbl
1 TopBPI
to bind to Sld3 (Tanaka et al. 2007; Zegerman and Diffley 2007) and for interaction with
the 9-1-1 checkpoint complex (Furuya et al. 2004; Delacroix et al. 2007; Lee et al. 2007).
If the interaction between TopBP1 and TICRR is relevant to their DNA replication
functions, then TICRR would be predicted to interact with the N-terminal BRCTs of
TopBP1. To address this, we created a panel of human TopBP1 mutants in which
specific BRCT domains were either deleted singly or pairwise (AI+ll, All1, AIV+V, AVI and
AVII+VlIl) and tested their ability to bind to TICRR in co-transfection assays (Figure 7B).
All five of the TopBP1 mutants were expressed at similar levels to the wild-type TopBP1.
TICRR co-immunoprecipitated with four of these mutants, AlII, AIV+V, AVI and AVII+VIII,
and with
wild-type TopBP1.
In contrast,
no TICRR
was recovered
in the
immunoprecipitate of the Al+ll deletion mutant even though TICRR was present at high
levels in these cells. Taken together, our data show that TICRR associates with TopBP1
in vivo and this interaction requires the two N-terminal BRCT domains that have been
associated with TopBP1's role in both checkpoint signaling and DNA replication.
A
41W 1-6W
0 BRCT domain
Input (2%)
IP: ot-GFP
\X
I\
eX\%
4
)
0A
&4~
4P
I>
%
XtI
0if1
Figure 7. TICRR interacts with the first two BRCT domains in TopBP1.
(A) Schematic diagram of TopBP1 protein. (B)Anti-GFP immunoprecipitates and whole cell
lysates (input) from cells transfected with GFP-tagged TopBP1 deletion mutants were screened
for TICRR and TopBP1 proteins by immunoblotting.
We were intrigued to find that TICRR association maps to the region of TopBP1
that is required for Sid3 binding. In yeast, formation of the Sid3- Dpbl
known to
require CDK
1 TopBPI
complex is
phosphorylation of Sld3 and this, together with the
96
phosphorylation of SId2, accounts for the CDK-dependence of DNA replication in this
organism (Tanaka et al. 2007; Zegerman and Diffley 2007). CDK is also essential for
replication in human cells, but there is some doubt as to whether pre-IC components will
be the relevant target (DeGregori et al. 1995). Given these questions, we used two
complementary approaches to examine the role of CDK phosphorylation in the TICRRTopBP1 interaction. First, we generated a population of HeLa cells that were
synchronously re-entering G1 from mitosis, and then cultured them in the absence or
presence of the pan-CDK inhibitor roscovitine for 8 hours (Figure 8A). Consistent with
the known CDK-dependence of S-phase entry, FACS analysis showed that the
untreated cells (-ROSC) were beginning to enter S-phase while the roscovitine treated
cells (+ROSC) remained blocked in G1 (Figure 8A). Notably, TopBP1 was recovered at
comparable levels in the TICRR immunoprecipitates of both the untreated and treated
cells (Figure 8A), suggesting that binding occurs in the presence of CDK-inhibition. In
the
second approach, we
incubated TICRR-TopBP1
immunoprecipitates
from
asynchronous cell extracts with, or without, lambda phosphatase and then assayed the
association of TICRR and TopBP1 by western blotting. Notably, phosphatase treatment
significantly increased the mobility of the TICRR protein (Figure 8B), indicating that
TICRR is phosphorylated in vivo and that we had successfully removed this modification.
Despite this change, there was no detectable difference in the levels of the TICRRTopBP1 complex in phosphatase treated versus untreated cells (Figure 8B). Importantly,
this result was not altered
by DNAase treatment of the extracts prior to
immunoprecipitation (data not shown), indicating that this complex formation occurs in
the absence of chromatin binding. Taken together, these synchronization and
phosphatase experiments strongly suggest that the interaction between TICRR and
TopBP1 can occur in a CDK-independent manner.
.......
............
A
+Noc
Release
18 hrs
Rosc
2 hrs
Analyze
B
8 hrs
Input IP ai-TICRR
Phosphatase
Phosphatase Inhib
AS
-Rosc
+Rosc
-
+
P1
-
+
-
+
a
DNA (PI)
AS
-Rosc
+Rosc
Input
IP:
Figure 8. TICRR can bind TopBP1 in a CDK independent manner.
(A) HeLa cells were synchronized in mitosis and then allowed to re-enter the cell cycle in the
absence (-Rosc) or presence (+Rosc) of roscovitine as depicted. FACS analysis showed that the Rosc. cells were entering S-phase, while the + Rosc. cells remained in G1. IP-western analysis
shows that the TICRR-TopBP 1 complex was recovered at similar levels in the -Rosc. and +Rosc.
samples. AS=asynchronous. (B) Asynchronous HeLaS3 cell extracts were immunoprecipitated
with anti-TICRR antibodies and then either incubated with lambda phosphatase, phosphatase
inhibitors or both. The mobility of the TICRR protein was significantly increased in the
phosphatase treated sample, versus the controls, but showed no difference in the levels of
associated TopBP .
Ticrr-deficiencyinhibitspre-IC formation
The TopBP1 orthologs in X. Iaevis, budding and fission yeast are known to be
essential for the transition of the pre-replication complex (pre-RC) into the pre-initiation
complex (pre-IC), an intermediate in the initiation of DNA replication (Garcia et al. 2005).
Having established that ticrr is essential for normal DNA replication in zebrafish, and that
TICRR and TopBP1 associate in the absence of DNA, we asked whether ticrr is similarly
required to form the pre-IC. To address this question, we employed the wild-type and
ticrr mutant zebrafish. For this analysis, we used embryos at 40 hpf, the developmental
timepoint at which the ticrr deficient cells have a profound replication defect, as judged
by S-phase accumulation and strongly reduced BrdU incorporation. Pools of wild-type
and mutant embryos were dissociated and used to generate extracts from whole cell or
chromatin-enriched fractions. These cells were then assayed by western blotting for
chromatin association of the Mcm2-7 complex (using a pan-MCM monoclonal antibody)
and the GINS complex (using an antibody against Psfl) that are core components of the
pre-RC and pre-IC, respectively (Figure 9). There was no difference in the chromatinassociation of the Mcm2-7 complex in wild-type versus ticrr mutant embryos. In contrast,
Psfl showed a significant level of chromatin association in the wild-type embryos but
was nearly absent (although still detectable on long exposure) from the chromatinenriched faction in the ticrr mutant, even though the Psfl protein was present at normal
levels in the whole cell extracts. Taken together our data show that, in concert with
TopBP1, Ticrr is required for the transition from pre-RC to pre-IC explaining its essential
role during DNA replication.
Total
Chromatin
Figure 9. Ticrr is essential for the chromatin
association of the pre-IC component Psfl.
4-
Total cell lysates and chromatin-enriched fractions were
prepared from 40 hpf wild-type (+/+ and +/-) and ticrr
mutant (-/-) zebrafish embryos. Immunoblotting for core
pre-RC and pre-IC components, the MCMs and Psfl
respectively, showed that these are expressed at normal
levels, but only the MCMs are chromatin loaded, in the
ticrr mutants.
cr
99
Discussion
Using a screen for zebrafish mutants that fail to arrest mitotic entry after
exposure to IR, we identified two mutant lines, each with a distinct mutation in the ticrr
gene. In addition to the G2/M checkpoint defect, these lines have a profound apoptotic
phenotype in the absence of exogenous damage that revealed a more general role in
cell survival during development. Consistent with this observation we found that ticrr
mutants at 40 hpf failed to incorporate BrdU, demonstrating that loss of ticrr also causes
a defect in DNA replication. Instead of arresting in S-phase, many cells in the ticrr
mutants proceed into mitosis and display an array of chromosomal abnormalities,
including fragmented chromosomes and anaphase bridges, that likely account for the
loss of anaphase cells through mitotic catastrophe. Thus, ticrr is required to prevent
mitotic entry both following exogenous DNA damage and also when DNA replication is
impaired. These phenotypes are highly reminiscent of those arising when TopBP1 and
its orthologs are inactivated. Accordingly, we found that the human and zebrafish TICRR
proteins both interact with TopBP1 in human cells and ticrr is required for transformation
of the pre-RC to the pre-IC in zebrafish. Taken together, these data suggest that TICRR
functions with TopBP1 in DNA replication and both the S/M and G2/M checkpoints.
TICRR in DNA Replication
The replication machinery and its mechanisms of regulation have been studied in
numerous organisms. These processes are generally conserved among eukaryotes but
a unifying model remains elusive because the order of assembly of pre-IC proteins
seems to differ between X. laevis and yeast, and known key components such as Sld3
have not been identified in higher eukaryotes. The initiation of DNA replication is best
100
understood in S. cerevisiae. Here, Dpbl 1 TopBP1, SId2, SId3, GINS, and Cdc45 are all
required to transform the pre-RC into the pre-IC. Sld3 and Dpbl 1 TopBP1 form a complex
that is induced by the CDK phosphorylation of Sld3 and requires the two N-terminal
BRCT domains of Dpbl1 (Tanaka et al. 2007; Zegerman and Diffley 2007). Although the
formation of the Dpb11-Sld3 complex is critical for replication initiation in S. cerevisiae,
an analogous event has not been defined in higher eukaryotes.
Much of our understanding of vertebrate DNA replication comes from studies in
X. laevis (Bell and Dutta 2002; Sclafani and Holzen 2007). In this organism Cut5TOPBP1 is
also required for the recruitment of DNA polymerases onto chromatin (Hashimoto and
Takisawa 2003; Kubota et al. 2003). Thus, the general function of Dpbl 1 TopBP1 in pre-IC
formation seems to be conserved in vertebrate TopBP1. Our data show that TICRR
displays some of the core properties of the Sld3 protein. First, Ticrr is essential for DNA
replication in unperturbed cells and it is specifically required for formation of pre-ICs, but
not pre-RCs. Second, TICRR associates with TopBP1 through the N-terminal BRCT
motifs I and 11that are conserved with the SId3-interacting BRCTs of Dpb 11TopBP1
However, our data do not address whether the interaction between TICRR and TopBP1
is direct, and other observations are less consistent with the idea that TICRR is a true
ortholog of Sld3. First, we note that there is no detectable sequence similarity between
these proteins. This is not a particularly conclusive finding, since the Sld3 and TICRR
proteins are both poorly conserved even within their own kingdoms, but it does raise
questions about both the relationship and mechanism(s) of action of these proteins. The
second, and more striking, finding is the apparent discrepancy in the role of CDK
phosphorylation in Sld3 versus TICRR regulation. Specifically, CDK phosphorylation of
SId3 is required for it to bind to Dpbl
1TopBP1
and this, together with the phosphorylation of
Sld2, accounts for the CDK-dependence of DNA replication in yeast (Tanaka et al. 2007;
Zegerman and Diffley 2007). In contrast, we find that the TICRR-TopBP1 interaction is
101
completely unaffected by culturing in the presence of the pan-CDK inhibitor roscovitine
or treatment with phosphatase. We note that there are potential limitations to these
approaches: the roscovitine-induced block to S-phase entry (figure 5F) could reflect a
reduction, but not full loss, of CDK activity; and/or the phosphatase may be unable to
access the phosphorylated residue(s) mediating the TICRR-TopBP1 interaction, even
though it effectively targets other phosphorylation sites (figure 5G). Despite these
caveats, our data are most consistent with the notion that TICRR associates with
TopBP1 in a CDK-independent manner. On first consideration, this finding seems to
imply that TICRR is not human Sld3. However, there is evidence to suggest that the
underlying basis for the CDK-dependence of DNA replication differs in human versus
yeast cells. First, DeGregori and coworkers have shown that E2F1 expression
completely bypasses the CDK-dependence of S-phase in human cells (DeGregori et al.
1995). This argues that there is no absolute requirement for CDK phosphorylation of
either pre-RC or pre-IC components in this organism. Consistent with this conclusion,
CDK phosphorylation is required for formation of the yeast Sld2-Dpbl1 complex but not
the vertebrate counterpart RECQL4-TopBP1 (Matsuno et al. 2006; Tanaka et al. 2007;
Zegerman and Diffley 2007). Clearly, additional studies are required to explore how
similar or different Sld3 and TICRR are to one another in the context of both DNA
replication and, as described below, checkpoint response.
TICRR in the SIM and G2/M Checkpoints
The S/M checkpoint plays a vital role in ensuring complete replication prior to
mitotic entry. Our data show that Ticrr is essential for the integrity of this checkpoint in
vivo. Thus, Ticrr joins a short list of proteins that play a dual role in both replication
regulation and S/M checkpoint response. The existing dual replication/checkpoint
proteins can be divided into two different subclasses, based on their role in the S/M
102
checkpoint. The Mcm2-7 complex, Cdtl and Cdc45 are representative members of the
first subclass. The analysis of yeast conditional mutants shows that the loss of MCMs,
Cdtl or Cdc45 prior to the initiation of replication allows cells with unreplicated DNA to
enter mitosis, but the loss of these proteins in replicating cells does not impair the S/M
checkpoint (Tercero et al. 2000; Labib et al. 2001). These findings suggest that these
pre-RC and pre-IC components are not directly involved in the S/M checkpoint. Instead,
they are required to create the replication structures which signal that S-phase is
ongoing and not yet complete. Studies using X. laevis extracts confirm that replication
structures are a pre-requisite for S-phase checkpoint signaling in vertebrates and further
suggest that the necessary feature is the RNA primer generated by DNA polymerase a
(Michael et al. 2000).
TopBP1 is an example of the second class of dual replication/checkpoint
proteins. Studies with conditional and deletion mutants show that TopBP1's checkpoint
function can be separated from its role in pre-IC formation (McFarlane et al. 1997; Saka
et al. 1997; Hashimoto et al. 2006; Yan et al. 2006). McFarlene and colleagues identified
a temperature sensitive mutant of Cut5 in S. pombe that fails to arrest when treated with
hydroxyurea at the semi-permissive temperature, but has normal replication capabilities
(McFarlane et al. 1997). Two independent studies in Xenopus egg extracts also showed
that the replication and checkpoint functions of TopBP1 could be separated (Hashimoto
et al. 2006; Yan et al. 2006). Addition of an oligonucleotide duplex to Xenopus egg
extracts bypasses the requirement of replication for checkpoint activation in response to
stalled replication forks (Yan et al. 2006). Using this system, it was shown that the Nterminal half of the TopBP1 protein rescues the replication defect caused by depletion of
TopBP1 in egg extracts, but is incapable of activating the checkpoint in response to
stalled replication forks, suggesting that the checkpoint defect is not a consequence of
loss of a complete replication complex and TopBP1 is directly involved in checkpoint
103
signaling. In fact, TopBP1 directly stimulates Chk1 phosphorylation by ATR in an in vitro
kinase assay and the C-terminus half of TopbP1 is required for this function.
Biochemical data shows that TopBP1 is recruited to the site of replication stress or DNA
damage (Furuya et al. 2004; Delacroix et al. 2007; Lee et al. 2007) and once recruited
promotes ATR activation (Kumagai et al. 2006). Together, these data strongly suggests
that TopBP1 is a bona fide checkpoint protein.
Given the precedent of the existing dual replication/checkpoint proteins, it
remains an open question whether Ticrr is directly involved in the S/M checkpoint or
whether this function simply reflects its role as an essential replication regulator. Based
on the following observations, we favor the former hypothesis. First, our data show that
TICRR interacts with TopBP1, raising the possibility that it cooperates in TopBP1dependent processes beyond DNA replication, such as checkpoint signaling. Second,
our FACS data show that ticrr-deficient cells enter mitosis with partially replicated DNA.
This clearly differs from the yeast MCM, Cdtl and Cdc45 mutants, which do not
prematurely enter mitosis once S-phase has begun (Tercero et al. 2000; Labib et al.
2001). Third, we originally identified ticrr through a screen for G2/M checkpoint
regulators. This showed that ticrr-deficient cells fail to arrest in mitosis in response to
treatment with ionizing radiation. Importantly, our FACS analysis shows that these
mitotic cells have 4N DNA content. Thus, we believe that this MAI phenotype reflects a
bona fide defect in the G2/M checkpoint and is not simply an indirect consequence of the
replication and/or S/M checkpoint defects. Consistent with this view, there is no evidence
in the literature that the G2/M checkpoint is dependent on appropriate replication
initiation. Moreover, included in our zebrafish screen were other known replication gene
mutants that did not display the MAI phenotype. Finally, we again note that TopBP1 has
a well-documented role in the radiation-induced G2/M checkpoint (Garcia et al. 2005).
Given all of these observations, we speculate that TICRR is a previously unknown
104
partner for TopBP1 in its myriad roles as a core regulator of the DNA replication, S/M
checkpoint and the G2/M checkpoint machinery.
Materials and methods
Zebrafish maintenance, collection and genotyping.Zebrafish were maintained
as previously described (Amsterdam et al. 2004). Primers used for genotyping were as
follows:
hil 573 - forward primer (F): AAGCAAGCTACATCTCAAAGCA;
hi1573 -reverse primer (R): CGGAAAACCCTGAAGTGTGAT;
hi3202A - (F) CAGATCCCCTGGTTATAAGTGTTGC;
hi3202A (R) CAGTTATGGCCTCTCAGA ATGTCGT;
reverse primer for provirus: GCTAGCTTGCCAAACCTACAGGT.
Plasmid construction
Human TICRR
A fragment of the 5' end of the human ticrr cDNA was generated by RT-PCR (F:
CACCATGCTGCTGCTGGACAC;
R: GGCCGGGTGAGGAGGAGTCTTTTA) from HeLa
cell mRNA and cloned into pENTR/D-TOPO to generate NC1. A short 5' sequence was
added to this fragment by ligating two annealed oligos (GGCCGCATGGCATGCTGTCA
CAAAGTAATGCTGCTGCTGGACACCGC
and GGTGTCCAGCAGCAGCATTACTTTG
TGACAGCATGCCATGC) into NC1 to generate NC2. The 3' end of the ticrr cDNA was
amplified by RT-PCR (F: GGATCCACTTTGGATTCGGAGGTCCTG,
CTGGCTGACTCCTTGATCT)
R: TTGGCGCGC
and ligated into NC2 to generate NC3. The full-length
cDNA cloned into pENTR/D-TOPO (NC3) was then transferred to an expression vector
(pDSLPCX-FTM-XB) using the LR Clonase (Invitrogen) Reaction 1l.
105
Zebrafish Ticrr
The zebrafish ticrr cDNA was generated by RT-PCR from mRNA of a single adult
zebrafish (F: ATGGCCTCTCAGAATGTCG;
R: CCGAACGTTAAAATCCATGC).
The
cDNA cloned in pENTR/D-TOPO was then transferred to an expression vector (pDESTEGFP-C1) using the LR Clonase (Invitrogen) Reaction 11.
Human TopBP1
The human TopBP1 cDNA was PCR amplified from pCR-XL-TOPO-TOPBP1 (Open
Biosystems;
BC126209.1;
F:
CACCATGTCCAGAAATGACAAAGAACC;
R:
TTAGTGTACTCTAGGTCGTTTGA) and transferred into pENTR/D-TOPO (Invitrogen)
and then pDEST-EGFP-C1 using the LR Clonase (Invitrogen) Reaction 11.In frame
deletions within the cDNA encoding TopBP1 were generated by PCR. Final products
were verified by DNA sequencing. The TopBP1 BRCT domain deletion mutants carry
the following in frame deletions within the 1522 a.a. ORF: A98-306 (BRCTs 1+11), A354452 (111), A547-760 (IV+V), A922-1011 (VI), A1267-1489 (VIl-VII). Except for A922-1 011,
the deleted amino acids were replaced by two glycines.
FACS analysis
For BrdU labeling in zebrafish, dechorionated embryos were incubated with 10mM
BrdU/1 5% DMSO on ice for 15 minutes, washed and incubated at 28.5*C for 15 minutes.
Mutant embryos were identified by either developmental phenotype or PCR genotyping
of a fraction of individual, fixed embryos. Cells were disaggregated by triturating
embryos in 0.25% Trypsin/lmM EDTA using a p200 and fixed in 70% ethanol at -20*C
overnight. Suspensions of cells from 20 wild-type or mutant embryos were pooled and
prepared for pH3/propidium iodide or BrdU/propidium iodide FACS analysis as
described (Pozarowski and Darzynkiewicz 2004). FACS analysis was conducted by
106
FACScan (Becton-Dickinson). DNA content was quantified by ModFit LT (Verity
Software), and pH3 and BrdU was quantified by FlowJo (Tree Star, Inc.).
Human cell culture and chromatin fractionation
HeLa, HeLa-S3, and 293FS cells were grown in DMEM with 10% FBS. For the
synchronization experiments, HeLa cells were cultured in presence of 100 ng/ml
nocodazole (Calbiochem) for 18 hours. Synchronized mitotic cells were recovered by
shake-off, replated in DMEM containing 10% FBS for 2 hours and then incubated for a
further 8 hrs in the presence or absence of 20 pM roscovitine (Calbiochem). The
isolation of soluble fraction (S1), soluble nuclear fraction (S2), and chromatin-enriched
fractions were conducted as described (Mendez and Stillman 2000). To solubilize the
chromatin-bound proteins, nuclei were treated with 50U of Micrococcal nuclease
(Worthington) for 2 minutes at 37*C.
Immunoprecipitation
Hela-S3 cells were lysed in modified RIPA buffer (50 mM Tris-HCL [pH 8.0], 150 mM
NaCL, 1 mM EDTA, 1% NP-40, 0.1% SDS, 0.1 mM PMSF, protease inhibitor cocktail
(Roche) 1 mM NaF, 10 mM p-glycerophosphate (b-GP), 200 uM Na3VO 4) for 30 minutes
in ice. The soluble supernatant was used for immunoprecipitations. When indicated,
lysates were treated with 2000U/ml DNase I (Roche; 30 min at 25 0C) or 20 pg/ml
ethidium bromide (30 min on ice) prior to TICRR IP. For phosphatase treatment,
precipitates were resuspended in k phosphatase buffer (NEB) with or without 400U k
phosphatase (NEB) and phosphatase inhibitors (50 mM NaF and 10 mM Na3VO 4), and
incubated at 300C for 30 minutes. The immunoprecipitates were washed four times with
RIPA and resuspended in 2X Laemmli Buffer.
107
Zebrafish chromatin preparation
Deyolked zebrafish embryos were triturated in 0.25% Trypsin/1 mM EDTA using a p200.
Disaggregated cells were filtered through a 35pm nylon mesh and washed once with
PBS. The chromatin-enriched fraction was prepared essentially as described (Aparicio et
al. 2009). Briefly, cells were lysed in 10 mM HEPES pH 7.9, 0.2 M KOAc, 0.1% Triton X100, 0.34 M sucrose, 10% glycerol, 1 mM 1,4-DTT, protease and phosphatase
inhibitors. The chromatin-associated fraction was recovered by spinning at 18,000g for
10 minutes and washed twice in lysis buffer before suspension in Laemmli Buffer.
Antibodies
A 6-His-tagged N-terminal fragment of human TICRR (NP_689472.3 amino acids 10941348) was expressed in bacteria, purified over Ni2+ NTA-agarose resin (Qiagen), and
used to immunize BALB/c mice or New Zealand White rabbits (Pocono Rabbit Farm).
Other antibodies were: phosho-H3 (sc-8656-R, Santa Cruz), mouse anti-goldfish CyclinB1 (B112; Katsu et al. 1993), PSTAIRE-Cdc2 (sc-53, Santa Cruz), tubulin (T9026,
Sigma), ORC2 (sc-13238, Santa Cruz), LaminA/C (2032, Cell Signaling), TopBP1
(NB100-217, Novus Biologicals and sc-32923, Santa Cruz), rabbit polyclonal anti-human
Psf1 (Aparicio et al. 2009), mouse monoclonal anti-human Pan-MCM (Austin et al.
1999), normal mouse IgG (sc-2025, Santa Cruz), and GFP (11814460001, Roche).
Acknowledgements
Antibodies for this study were provided by Dr. Juan Mendez, Spanish National
Cancer Research Center (anti-Psfl) and Dr. Stephen Bell, HHMI and MIT (Pan-MCM).
We also thank Stephen Bell, Sebastian Hoersch and members of the Hopkins and Lees
labs for helpful discussions during this study and the preparation of this manuscript. This
work was supported by Ruth L. Kirschstein NRSAs to C.S. and N.C. J.A.L. is a Daniel K.
108
Ludwig Scholar.
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CHAPTER IV: Discussion
112
Key conclusions
We have screened a collection of zebrafish insertional mutants for genes
required for G2/M checkpoint activation in response to IR treatment. The work described
in this thesis focuses on the characterization of a novel gene discovered in this screen,
ticrr. Our work has demonstrated that Ticrr is required for replication initiation,
specifically for the conversion of the pre-RC into pre-IC. In addition to a G2 checkpoint
defect in response to DNA damage, ticrr mutants have a defect in the S/M checkpoint
that monitors DNA replication during normal cell cycle progression (Sansam et al.,
2010). These results indicate that ticrr is a novel vertebrate cell cycle regulator, with
essential roles in replication initiation and the surveillance mechanisms protecting the
cell from genomic insults during replication, as well as during exposure to exogenous
DNA damaging agents.
The human TICRR protein binds TopBP1, a known replication and checkpoint
factor. This interaction is conserved in zebrafish, suggesting that it is functionally
significant. Interestingly, the interaction is mediated through the first two BRCT domains
of TopBP1, previously shown to be essential for its replication and checkpoint function.
We conclude that TICRR is a TopBP1 partner that probably complements its role in cell
cycle regulation. Currently the mechanism of action of TICRR and how it collaborates
with TopBP1 is unknown, but merits further investigation.
A screen in zebrafish identifies Ticrr
We have screened 336 insertional mutant lines to identify novel cell cycle
regulators. Five mutant lines had a mitotic phenotype, while nine mutant lines had a
checkpoint activation failure in response to IR treatment. These candidates represent
4% of the total lines screened. Although we have proven the sensitivity of the screening
113
assay, it is possible that we have missed some candidates. One of the most important
considerations is the timing for screening. We selected 32 hpf as our timepoint for the
screen based on the level of proliferation occurring at this developmental stage.
However, It is possible that some mutants that showed no change in pH3 staining could
have phenotypes at a later developmental stage. On the other hand, some mutants that
were not screened because they are highly necrotic at 32 hpf could be screened at
earlier stages.
In addition, it has been calculated that the zebrafish collection used for the
screen represent 25% of all essential genes in this organism (Amsterdam et al., 2004). It
is probable that there are other novel genes encoding checkpoint proteins that were not
represented in our collection. Nevertheless, this screen was successful in identifying
ticrr, a novel gene essential for activation of the IR-induced G2/M checkpoint. This gene
is of particular interest for its additional roles during normal cell cycle progression. Ticrr
is conserved among metazoans and we anticipate that it will play a role in checkpoints
and cell cycle progression in other organisms, including humans.
Ticrr is required for DNA replication and functions in a
similar manner to Sld3
ticrr zebrafish have a defect in replication, demonstrated by the abnormal
accumulation of S-phase cells in the mutant embryos and a significant decrease in BrdU
incorporation. In addition, we showed that Ticrr is required for chromatin association of
Psfl (part of GINS, a pre-IC component), but not for the association of the MCM2-7
proteins (a pre-RC component). This data suggests that Ticrr plays an important role in
the conversion of the pre-RC into the pre-IC. This process is best understood in yeast,
and involves the recruitment of a number of proteins to origins in a particular order. In
metazoans, studying the mechanism of replication initiation has been more challenging
114
because origins are not well defined and a consensus sequence has not been identified.
Additionally, identification of orthologs of the yeast replication factors Sid2 and SId3 has
proven very difficult.
RecQL4 has emerged as a candidate for the Sld2 ortholog in vertebrates. The
N-terminal part of RecQL4 shows limited sequence conservation to Sld2 and this protein
is essential for replication initiation in Xenopus egg extracts and human cells (Im et al.,
2009; Sangrithi et al., 2005; Xu et al., 2009). Sld3 homology is poor among yeast
species so it is not surprising that a homolog in higher eukaryotes could not be identified
based on sequence similarity. Nevertheless, the fact that all the other known replication
proteins and the fundamental aspects of replication are conserved suggests that an
ortholog, or at least a functional analog, exists in higher eukaryotes. Intriguingly, Ticrr
shares several characteristics with the yeast Sld3 protein. Similarly to Sid3, Ticrr plays
an important role in the conversion of the pre-RC complex to pre-IC. Furthermore,
human TICRR interacts with TopBP1, the human ortholog of Dpbl 1, through TopBP1
BRCT domains I and 11. This is reminiscent of the yeast Sld3, which interacts with
Dpbl1 through the same BRCT domains, which are highly conserved from yeast to
humans (Tanaka et al., 2007; Zegerman and Diffley, 2007).
Despite the functional similarities between TICRR and SId3, we were unable to
find significant sequence similarity. However, Sanchez-Pulido et al. recently showed that
TICRR does in fact contain a domain that shares significant sequence homology with
Sld3 (Sanchez-Pulido et al., 2010). Using a conserved region in metazoan Ticrr as the
query, they found divergent plant homologs. Then they generated a profile of metazoan
and plant Ticrr orthologs and used the HHpred method to compare it with a profile from
the Sld3 protein family and found highly significant similarity. The authors concluded that
this level of significance indicates that these proteins descend form a common ancestor
prior to the divergence of the main eukaryotic lineages. Interestingly, the domain in SId3
115
that interacts with the TopBP1 yeast homolog Dpbl1 does not lie within this conserved
domain (Tanaka et al., 2007; Zegerman and Diffley, 2007), nor does the region in TICRR
that interacts with TopBP1
(my results, discussed in the Appendix). However,
temperature sensitive mutants in yeast Sld3 that abrogate replication lie within this
conserved domain suggesting that it is functionally significant and it may bind to a
different factor (Sanchez-Pulido et al., 2010). Further studies are needed to elucidate
what is the function of this region.
Kumagai et al. identified the Xenopus ortholog of Ticrr, named Treslin, in a
screen for TopBP1 interacting proteins (Kumagai et al., 2010). Consistent with our
findings, they showed that depletion of Xenopus Ticrr/Treslin protein and the human
counterpart causes a defect in replication and the pre-IC component Cdc45 is not
recruited to chromatin. They also showed that Xenopus Ticrr/Treslin binds Cut5/TopBP1
through the first two BRCT domains in Cut5/TopBP1. This indicates that the function of
Ticrr is highly conserved in vertebrates. We haven't directly tested whether the TopBP1
interaction with TICRR is required for TICRR's function in replication initiation. However,
this seems very likely since a direct role for TopBP1 orthologs in the initiation of DNA
replication has been described (Garcia et al., 2005). The Xenopus and human TopBP1
proteins are required for the recruitment of pre-IC components to chromatin, but not for
the formation of the pre-RC (Hashimoto and Takisawa, 2003; Jeon et al., 2007; Matsuno
et al., 2006; Van Hatten et al., 2002). Furthermore, the replication defect caused by
depletion of TopBP1 in Xenopus egg extracts can be rescued by the N-terminal half of
the protein containing BRCT domains I through V (Hashimoto et al., 2006; Yan et al.,
2006). This implies that TopBP1 replication functions are all contained in the first half of
the protein, which includes the BRCT motifs required for TICRR binding. TICRR mutants
that fail to bind TopBP1 will be particularly useful for testing whether the TICRR-TopBP1
interaction is functionally significant, by assaying whether they can rescue the replication
116
defect caused by TICRR depletion.
The role of CDK activity in regulating TICRR function in
DNA replication
It has been known for a long time that CDK activity is essential for replication
initiation. Although in yeast SId2 and Sld3 are the minimum CDK targets for replication
initiation (Tanaka et al., 2007; Zegerman and Diffley, 2007), in metazoans the CDK
targets that are functionally significant for replication initiation remain elusive. The
possibility that TICRR may be the functional analog of Sld3 raised a critical question:
does CDK regulates TICRR function in an analogous manner, in particular, is TopBP1
binding to TICRR CDK dependent? We identified multiple putative CDK phosphorylation
sites in the TICRR sequence; however, our experiments did not show that CDK activity
is important for the interaction, even though we confirmed that TICRR is phosphorylated
in vivo. Contrary to our findings, Kumagai et. al. reported that Xenopus Ticrr/Treslin
binds TopBP1 in a phosphorylation and CDK2 dependent manner (Kumagai et al.,
2010). Although the discrepancies between these results remain unclear, there are
several possible explanations. First, it is possible that the precise regulation of TICRR
function might have diverged in different organisms. There are notable differences in
pre-IC regulation even between S. pombe and S. cerevisiae. For instance, Sld3
recruitment to origins is not Cdc45 dependent in fission yeast, whereas SId3 in budding
yeast forms a complex with Cdc45 before associating to chromatin and their association
is mutually dependent (Yabuuchi et al., 2006). It has not been addressed if the SId3
interaction with Cut5 (TopBP1 homolog) in fission yeast is dependent on CDK activity,
but 3 putative phosphorylation sites were mutated in a study and disruption of at least
those specific sites didn't affect chromosome replication (Nakijima et al., 2002). It is
important to note that this result doesn't rule out that other phosphorylation sites in the
117
protein are required for replication. Alternatively, the difference could be accounted for
by the fact that I performed experiments using cancer cell lines. Perhaps there is still a
requirement for phosphorylation in normal human cells, but it is bypassed in tumor cells,
where the regulation could be different. Lastly, there are caveats to the experiments that
I performed that could explain why I didn't observed a disruption of the complex under
the conditions used in my experiments. The treatment of cells with roscovitine may not
have fully inhibited CDK activity; and/or the phosphatase may be unable to access the
phosphorylated residue(s) mediating the TICRR-TopBP1 interaction, even though it
effectively targets other phosphorylation sites.
Other proteins with functional similarity to TICRR
In the last year, two other proteins were identified that have similar
characteristics to TICRR. GEMC1 is a novel vertebrate protein identified in Xenopus
(Balestrini et al., 2010). Like TICRR, it interacts with TopBP1 and depletion from
Xenopus egg extracts or from
mammalian cells causes a defect in replication. Cdc45
and the GINS component Sld5 are not recruited to chromatin in GEMC1 depleted cells,
indicating that this protein is also required for pre-IC formation. Similarly to Sld2 and
Sld3, GEMC1 is phosphorylated by Cdk2 and a phosphomimetic form of GEMC1
stimulates TopBP1 binding, Cdc45 recruitment and DNA replication, indicating that these
phosporylations are functionally significant. Nevertheless, the phosphomimetic mutant
could not bypass the requirement for CDK2 in replication, suggesting that there are other
targets required for initiation of DNA replication (Balestrini et al., 2010). DUE-B is yet
another protein recently found to be required for replication initiation that interacts with
TopBP1. Initially identified for its ability to bind DNA regions of predicted helical
instability (DNA unwinding elements) (Casper et al., 2005), it is also required for Cdc45
loading to chromatin. In contrast to SId3, phosphorylation is not essential for DUE-B
118
interaction with TopBP1 (Chowdhury et al., 2010).
An interesting question to investigate in the future is the relationship between
TICRR and these other novel replication factors. Some issues that can be addressed are
the order in which these proteins are recruited to chromatin during replication initiation,
what proteins are required for their recruitment and whether they can bind to each other
to form a functional complex or complexes required for replication initiation. It is possible
that GEMC1 or DUE-B, or both of them, could be performing the role of Sld2 in
vertebrates. The proposed ortholog of Sld2 in higher eukaryotes, RecQL4, is also
required for replication initiation, but studies in Xenopus and human cells suggest that it
doesn't share the same molecular function as its counterpart in yeast. Further studies
are needed in order to understand better the role of these factors, including TICRR,
during the initiation steps of DNA replication and how they might work together. We can
anticipate that the process of replication initiation in vertebrates is more complex than
that of yeast and although some aspects will be very similar, there will be notable
differences.
Possible roles of TICRR in the maintenance of genome
stability
The replication machinery works in conjunction with DNA damage checkpoint
proteins to protect cells from possible genomic insults occurring during DNA replication.
The replication proteins MCM7, RPA, DNA polymerase a, and TopBP1 have been
shown to play a role in ATR-mediated checkpoint signaling. Mutations in replication
proteins may indirectly affect the intra-S phase checkpoint because replication structures
are required for checkpoint activation. On the other hand, certain proteins, such as
TopBP1, have direct roles in mediating checkpoint activation. This dual role of TopBP1
has been demonstrated by defining separate domains encoding the replication and
119
checkpoint functions (Hashimoto et al., 2006; Yan et al., 2006). A recent screen
performed in Drosophila identified several pre-IC components to be required for G2/M
checkpoint signaling, including TopBP1, Cdc45 and RecQL4 (Kondo and Perrimon,
2011), suggesting that these proteins also have a role in checkpoints outside of Sphase. Ticrr fits into the category of proteins with a dual role in replication and
checkpoints.
We observed that zebrafish ticrr mutants have a number of phenotypes
associated with genomic instability. Firstly, cells in the ticrr mutants continue to enter
mitosis after IR treatment. Secondly, untreated cells that do not complete DNA synthesis
due to a defect in replication fail to arrest and instead enter mitosis prematurely. Thirdly,
abnormal mitotic figures and chromosomal fragmentation were detected in ticrr mutants.
These phenotypes suggest that Ticrr plays an important role in checkpoint activation in
response to DNA damage and during normal replication. It is possible, however, that
abnormal mitosis and metaphase arrests in ticrr mutants lead to the persistence of
mitotic cells after DNA damage regardless of the G2/M checkpoint. However, our results
argue against this possibility, as the mitotic index of untreated ticrrzebrafish embryos did
not greatly differ from wild-type zebrafish embryos at the timepoint chosen for the
screen. We also confirmed that cells in ticrr embryos continue to enter mitosis after IR
exposure using nocodazole, a drug that blocks cells in mitosis, which caused further
accumulation of mitotic cells in the mutants. Therefore, the sustained presence of mitotic
cells after DNA damage is unlikely to be a secondary consequence of DNA replication or
S/M checkpoint defects, but caused by a bona fide G2/M checkpoint defect. We
conclude that maternal stores contributions results in a hypomorphic phenotype in ticrr
mutants, in which cell cycle progression is only moderately affected but the G2/M
checkpoint is severely impaired during early stages of development.
Based on the link that other replications proteins have with the ATR pathway,
120
including Ticrr's binding partner TopBP1, it seems plausible that TICRR may be involved
in the ATR signaling pathway. Ticrr could have a role in the activation of ATR or
recognition of downstream substrates. Although TopBP1 directly activates ATR in vitro in
the absence of DNA, the recruitment of TopBP1 to chromatin is required for ATR
activation in vivo (Kumagai et al., 2006; Lee and Dunphy, 2010). Several studies
suggest that the 9-1-1 complex is involved in TopBP1's recruitment to chromatin,
however, these studies do not rule out the possibility that additional proteins may be
required for this process. Ticrr interacts with the same BRCT domains in TopBP1 that
are required for Rad9 binding. It will be informative to know whether a complex
containing both TICRR and Rad9 exist or the binding of these proteins to TopBP1 is
mutually exclusive.
In addition, TopBP1 has been associated with mediating the bridge between
ATM and ATR signaling pathways in response to DSBs. ATM phosphorylation of a
critical residue in the ATR-activation domain of TopBP1 is required for ATM-dependent
activation of ATR in response to DNA damage (Yoo et al., 2007). Furthermore, human
and Xenopus TopBP1 associate with the MRN component Nbsl (Morishima et al., 2007;
Yoo et al., 2009) which mediates the recruitment of ATM to TopBP1 in response to
DSBs and this has been suggested as a mechanism for the activation of ATR in
response to DSBs (Yoo et al., 2009). Interestingly, the Nbsl interaction also occurs
through the first two BRCT domains in TopBP1. Since Ticrr was found to be required for
G2/M checkpoint activation in response to IR, which induces DSBs, an interesting
possibility is that Ticrr may have a role mediating the activation of ATR through its
interaction with TopBP1.
Kumagai et al. observed that depletion of Ticrr/Treslin in human cells causes an
increase in H2AX phosphorylation, indicative of DNA damage, which is consistent with
121
our findings that chromosomal fragmentation occurs in the ticrr zebrafish mutants
(Kumagai et al., 2010). However, Chk1 fails to become phosphorylated in the activating
residues S317 and S345 even after treatment with the drug aphidicolin, which induces
stalled replication forks (Kumagai et al., 2010). These results suggest that, similarly to
zebrafish Ticrr, the human ortholog is involved in checkpoint signaling. However, this
study did not address the phosphorylation status of Chk2 in Ticrr depleted cells;
therefore it is unclear whether this defect is specific for Chk1 and the molecular basis for
this defect. In any case, this data suggest that the function of Ticrr in checkpoints is
conserved in vertebrates.
S. cerevisiae SId3 has been recently identified as a target of the intra-S phase
checkpoint. In response to stalled replication forks, the Rad53 kinase (homolog of Chk2,
downstream of Mecl/ATR) inhibits origin firing by phosphorylating SId3, preventing it
from binding to Dpbl1 (Lopez-Mosqueda et al., 2010; Zegerman and Diffley, 2010). The
yeast SId3 proteins have not been described to regulate other checkpoints. While it is
not known whether Ticrr is targeted by checkpoints in a similar manner, I suspect that
Ticrr will have a more general role in checkpoint signaling. The vertebrate TICRR
proteins are considerably larger than the SId3 proteins, suggesting that they may have
acquired additional interaction partner and additional functions through evolution.
Identification of other Ticrr interactors will further amplify our understanding of the
functions of this protein.
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124
APPENDIX A: Characterizing the functional
significance of the TICRR-TopBP1 interaction
Nelly M. Cruz and Jacqueline A. Lees
125
We identified TopBP1 as a binding partner of the zebrafish and human TICRR
proteins. TopBP1 is a multifunctional protein with key roles in replication and ATRmediated checkpoint signaling. Little is known about the biological significance of
TopBP1's interaction with TICRR. This appendix contains data that further characterizes
the TICRR-TopBP1 interaction and investigates mechanisms by which TICRR regulates
replication initiation in human cells.
Results and discussion
The TopBP1 interaction domain lies within the C-terminal half of TICRR
Having determined the region of TopBP1 that associates with TICRR, we sought
to define the region of TICRR participating in this interaction. Human TICRR is a large
220 KDa protein that based on sequence analyses has no identifiable functional
domains. We prepared GFP tagged fragments of TICRR containing either the N-terminal
half, (residues 1-969), or the C-terminal half (residues 970-1910) of the protein and cotransfected
these
fragments
with
full-length
TopBP1
into
293T
cells.
By
immunoprecipitating the tagged TICRR protein with anti-GFP antibody, we observed that
TopBP1 binds to the C-terminal but not the N-terminal fragment of TICRR (Figure 1).
We then tested several smaller C-terminal fragments of TICRR to define more
precisely the minimum requirements for binding to TopBP1. We observed that deletion
of residues 1565 to 1910 of TICRR, which excises the very end of the C-terminus, did
not affect the binding of TopBP1 (Figure 1). However, any deletion that removed
residues 976-1565 dramatically reduced the interaction with TopBP1 (Figure 1 and data
not shown). Importantly, a fragment consisting of residues 976-1261 binds to TopBP1,
indicating that this region is sufficient for the interaction (Figure 1).
126
976
WT
1261 1565 1910
IP: a-GFP
TopP1 indng
+
-M
2
3
+-n
-:
6
+
-
Figure 1. Residues 976-1261 of TICRR protein are required and sufficient for
TopBPI binding.
(A) Schematic diagram of TICRR truncations and summary of the abilities of the indicated forms
to interact with TopBP1. (B) Anti-GFP immunoprecipitates from cells transfected with GFPtagged TICRR truncation mutants were screened for TICRR and TopBP1 proteins by
immunoblotting.
Two putative CDK phosphorylationsites are highly conserved in TICRR
There are many consensus CDK sites within the TICRR sequence. Interestingly,
the vast majority of these sites are located within the C-terminus of the protein.
Furthermore, we identified a proline-rich region in which many of these sites are
clustered. This region contains 54 serine or threonine residues followed by proline, the
minimal CDK consensus site; 13 of these also contained an arginine or lysine two
positions downstream of the proline, which is the full consensus site for CDK
phosphorylation.
Interestingly, this region spans the TopBP1 interaction domain
identified by our binding assays.
Sanchez-Pulido and colleagues found significant homology between the TICRR
127
and Sld3 sequences (Sanchez-Pulido et al., 2010). CDK phsophorylation of Sld3 at two
sites, Thr 600 and Ser 622, is required for Dpbl 1 TopBP1 binding to SId3 and subsequent
pre-IC formation. As described in Chapter 3, we examined whether phosphorylation is
required for the TICRR-TopBP1 complex formation by treating cells with the CDK
inhibitor roscovitine as well as phosphatase treatment of cellular lysates, but did not
detect a disruption in the interaction. We note that there are limitations to this
experimental approach (discussed in Chapter 3). Therefore, we have decided to use a
more direct approach in order to test if phosphorylation plays a role in TICRR's
replication function. By searching the TICRR protein sequences for highly conserved
CDK consensus motifs, we identified a consensus sequence containing serine 1001 (in
human TICRR) that is present in all metazoan species examined, including the primitive
chordates Branchiostoma floridae and Ciona intestinalis (Figure 2). We aligned this
sequence with the CDK sites in S. cerevisiae Sld3 required for replication and found that
it is highly similar to the consensus sequence of Sld3's Serine 622 site (Figure 2). We
identified a second consensus sequence highly conserved across TICRR metazoan
species 32 residues upstream of Ser 001 (Threonine 969). Although the conservation is
low, there is similarity between this consensus sequence and the sequence
encompassing Sld3's Thr 600 (Figure 2).
To test whether these conserved sites play a role in TopBP1 interaction, we
replaced them
with
alanine residues, which cannot be phosphorylated.
Co-
immunoprecipitation experiments showed that there was no significant difference in the
binding capabilities of TICRR when either site was changed to alanine (Figure 3). In
contrast, the double mutant had reduced binding to TopBP1. It is important to note that
the interaction was not completely disrupted by these mutations (Figure 3). It is possible
that other phosphorylation sites or sequences are contributing to the interaction and loss
of phosphorylation at these two sites is not sufficient for complete disruption of the
128
complex. Alternatively, these amino acids may have an important function in replication
initiation that is independent of TopBP1. As previously discussed, the mechanistic
details of replication initiation differ among species. We are currently developing a robust
system for performing rescue experiments to examine whether these phosphorylation
sites play a role in TICRR's function.
Homo sapiens
Mus musculus
Gallus gallus
Xenopus laevis
Gasterosteus aculeatus
Takifugu rubripes
Danio rerio
Branchiostoma floridae
Capitella sp
Nematostella vectensis
Strongylocentrotus purpuratus
xodes scapularis
Lottia gigantea
Ciona intestinalis
Schistosoma mansoni
Saccharomyces cerevisiae
Homo sapiens
Mus musculus
Gallus gallus
Xenopus laevis
Gasterosteus aculeatus
Takifugu rubripes
Danio rerio
Branchiostoma floridae
Capitella sp
Nematostella vectensis
Strongylocentrotus purpuratus
Ixodes scapularis
Lottia gigantea
Ciona intestinalis
Schistosoma mansoni
Saccharomyces cerevisiae
K74A'
P'
74vAET:3
2RCZAAI:A-~
V2 LVL
K \ "r:qN~
K
:K2-
IvEEZ PEK 'iRDIDLR.
IvEEC'vK ADADL:G
c-AD- DL:
a
-
7LK
IEE
v
CDDO
-KDDEv
L
Figure 2. Sequence alignment of conserved CDK consensus motifs in the TICRR
protein.
Sequence alignements of putative CDK motifs for Thr 969 (A) and Ser 1001 (B). Sld3 sequences
including the CDK sites Ser600 (A) and Thr622 (B) were included in the alignment for
comparison.
129
IP: a-GFP
N
Figure 3. Mutation of Thr 969 and Ser 1001 in the TICRR protein causes a
reduction in TopBP1 binding
Thr 969 and Ser 1001 were changed to alanine redues in the GFP-tagged TICRR protein. AntiGFP immunoprecipitates from cells transfected with GFP-tagged TICRR truncation mutants were
screened for TICRR and TopBP1 proteins by immunoblotting.
To test the roles of TICRR in replication, we used a siRNA that was described by
Kumagai and colleagues (Kumagai et al., 2010) to knockdown the TICRR protein in
human cells. Using these cells we could introduce mutant forms of TICRR and assay
which domains of TICRR are required for replication. This siRNA causes robust
depletion of the TICRR protein 48 hours after transfection (Figure 4A). We confirmed the
replication defect previously observed by Kumagai et al. that is consistent with the
replication defect observed in the zebrafish ticrr mutants. Specifically, we compared the
ability of U2OS cells transfected with siRNA against TICRR (siTICRR) or a control
siRNA (siControl) to incorporate the nucleotide analog bromodeoxyuridine (BrdU). Cells
transfected with siTICRR incorporated relatively little BrdU, in comparison with control
cells even though a significant fraction of the cells contained between 2N and 4N DNA
content (Figure 4B). This demonstrates that TICRR-deficient cells are impaired for DNA
replication.
130
+TICRR
TICRR
TopBP1
+siResTICRR
-
siControl
siTICRR
21.1%
5.91%
-
.
-
--.
DNA content (PI)
Figure 4. Loss of TICRR in human cells causes a replication defect.
(A) U2OS cells were transfected either with a control siRNA or TICRR-specific siRNA, along
with wild-type TICRR or a form of TICRR that is resistant to knockdown. There is robust
decrease of the wild-type TICRR protein but not of siRNA resistant TICRR (sires-TICRR) 48
hours after transfection. TopBP1 protein levels are shown as a loading control. (B) BrdU and PI
FACS analysis of BrdU pulsed labeled siControl and siTICRR cells. Quantification of the BrdU+
population showed a dramatic reduction in replicating DNA in siTICRR cells.
We used FACS to determine the DNA content of phosho-histone H3 positive
cells during normal cell cycle progression in TICRR depleted cells (Figure 5). As
expected, histone H3 phosphorylation is restricted to cells with 4N DNA content in
control cells (Figure 5). In contrast, we find that a high percentage of the pH3-positive
cells in the TICRR deficient cells have less than 4N DNA content, indicative of a S/M
checkpoint defect. This phenotype is equivalent to the defect observed in cells isolated
from ticrr mutant zebrafish embryos, further confirming that TICRR's roles are conserved
among vertebrates.
131
DNA content (PI)
Figure 5. Ticrr is required for S/M checkpoint function in human cells.
FACS measurement of anti-pH3 and PI staining of siControl and siTICRR cells showed a
population of mutant pH3-positive cells with less than 4N DNA content in TICRR deficient cells,
indicative of entry into mitosis before completing DNA replication.
TICRR-depleted cells underwent morphological changes including enlargement
and flattening of the cells on culture dishes (Figure 6). These changes started to become
noticeable 48 hours after transfection of the TICRR siRNA. The size of the nuclei of
TICRR-deficient cells increased by as much as 3-4-fold relative to the control cells.
These morphological changes are the typical phenotypes of senescent cells (Hwang et
al., 2009). Consistent with these morphological changes, TICRR-deficient cells stained
positive in a senescence associated P-galactosidase assay (Figure 6). Importantly, the
control cells did not undergo any morphological changes and did not stain in the pgalactosidase assay (Figure 6). Most likely, this senescence phenotype is a
consequence of DNA damage accumulating in TICRR-depleted cells due to defects in
replication and activation of the S/M checkpoint.
132
siControl
siTICRR
Figure 6. TICRR knockdown induces cellular senescence in U2OS cells
U2OS human cell lines were transfected with control siRNA or TICRR-specific siRNA,
every 4 days for a period of 12 days. Cells were stained with p-galactosidase stain and
viewed under a phase contrast microscope.
In summary, TICRR deficiency causes replication and S/M checkpoint defects,
as well as cellular senescence in U2OS cells. These phenotypes can be assayed for
rescue upon the expression of different mutant forms of TICRR, including phosphomutants and truncations that are defective for TopBP1 binding. These assays will enable
the identification of functional domains within the TICRR protein.
A TICRR mutant that localizes to PCNA and RPA foci
Biochemical experiments have shown that TICRR is a chromatin-associated
protein. We noticed that the GFP-TICRR fusion protein consisting of residues 1565-1910
1910), the C-terminus
(TICRR' 56 5of the protein, localizes to foci within the nucleus while
wild-type GFP-TICRR localization is diffuse throughout the nucleus (Figure 7). A large
19 ' 0 had foci and the size
percentage of cells expressing GFP-TICRR' 565and number of
these varied between cells. Nuclear foci are characteristic of sites of ongoing replication,
as well as the sites of stalled replication forks and DNA damage. We sought to
determine what these foci might be by performing co-localization experiments using
133
indirect immunofluorescence to locate proteins known to localize to nuclear foci. We
stained for phosphorylated H2AX (6H2AX), a marker of double strand breaks and the
replication proteins RPA and PCNA. In the absence of DNA damage, we did not detect
increased H2AX in cells transfected with GFP-TICRR156
1910 ,
indicating that there is no
increase in DNA damage in these cells (data not shown). When treated with yIR, GFP
positive cells accumulated 6H2AX foci in a comparable manner to GFP negative cells
(Figure 8). However, GFP-TICRR'5 51910 foci did not colocalize with ylR-induced 6H2AX
foci. In addition, IR treatment did not induce any obvious changes in the foci pattern of
910 (Figure 8).
GFP-TICRR' 565-1
GFP
DAPI
TICRR:
Figure 7. GFP-TICRR
565 '9'0
Merge
1565
1910
I
1
localizes to foci within the nucleus.
GFP expression of cells tranfected with full length GFP-TICRR or a truncation containing
residues 1565-1910. A schematic representation of the TICRR protein showing the location of the
truncated fragment in orange is included.
134
GFP-TICRR
1565-1910
8-H2AX
Figure 8. GFP-TICRR'*5-'9'* foci do not colocalizes with 8H2AX.
U2OS cells transfected with GFP-TICRR1565-19 10 were treated with ionizing radiation, followed by
methanol fixation and staining for 6H2AX one hour after treatment.
On the other hand, the GFP-TICRR' 5 ,1910 foci co-localized with PCNA and RPA
910
foci in unperturbed cells (Figure 9). This indicates that TICRR' 719~1
is localizing to
either sites of ongoing DNA replication or sites of stalled replication forks. The
hypothesis that these foci represent stalled replication forks seems unlikely because the
GFP-TICRR' 565-1910 positive cells did not accumulate 8H2AX foci in the absence of ylR.
H2AX is phosphorylated at sites of stalled replication forks and forms foci that colocalize
with RPA after replication inhibition (Balajee and Geard, 2004).
We analyzed the cell cycle of cells transfected with GFP-tagged wild-type or
135
TICRR
565 1910
~
(Figure 11). Cells expressing TICRR 1565-' 10 showed an increased
accumulation of cells containing between 2N and 4N DNA content when compared to
GFP negative cells within the same population, or with cells expressing wild-type TICRR
(Figure 10), indicating that expression of the truncated form of TICRR is causing a defect
during replication. A BrdU incorporation assay will be informative to test whether
replication is occurring or whether cells are blocked in S-phase. It is possible that
replication is occurring at a slower rate, leading to the accumulation of cells in S phase.
Alternatively, a regulatory mechanism of origin firing may have been lost, and more cells
are undergoing replication.
This mutant may be exerting its dominant effect by sequestering an important
factor for replication. Given that this fragment of the protein is lacking the region that
binds to TopBP1 it is likely that the primary factor being sequestered is not TopBP1. It
will be informative to elucidate what are the binding partners of this TICRR fragment and
whether these interactions reflect functions of wild-type TICRR during replication.
Currently, we do not know if wild-type TICRR localizes to replication foci, whether it
moves with replication forks during normal replication or if it re-localizes to replication
forks when replication is inhibited. All these questions merit further investigation and the
1910 fusion protein
GFP-TICRR 1565could be a useful tool for dissecting the roles of TICRR
in replication.
136
GFP-TICRR 1565-1910
RPA
GFP-TICRR 1565-1910
PCNA
Figure 9. GFP-TICRR'
565 1910 foci
colocalizes with RPA and PCNA.
U2OS cells transfected with GFP-TICRR 1 .19'. were stained for RPA (A) and PCNA (B).
137
100%
G2/M
~~80%
Us
70%
2
60%
U
50%
0
40%
U
Gi
30%
20%
10%
WT
GFP-
WT
GFP+
Mut
GFP-
Mut
GFP+
DNA content (PI)
Figure 10. Expression of GFP-TICRR
565 '9'0 impairs
S-phase progression.
65
(A) Cell cycle profiles from cells expressing GFP-tagged wild-type TICRR and GFP-TICRR'15 65 ~
1910. Quantification of the GI, S or G2/M DNA content of wild type TICRR or GFP-TICRR
~
19 10
transfected cells reveals an increase in S-phase cells in the population expressing GFPTICRR 565-1910 .
Materials and methods
Plasmid construction
Restricition enzymes were used to cut the TICRR sequnece into fragments that were
cloned into pENTR/D-TOPO, and then into pDest-GFP expression vector using the
Gateway recombianation cloning technology (Invitrogen).
Human cell culture and transfections
293FS and U2OS cells were grow n in DMEM supplemented with 10% FBS.
Lipofectamine RNAiMAX from Invitrogen was used, following the manufacturer's
instructions, to transfect 25nM siRN As into U2OS cells. DNA transfections were
138
performed using Mirus TranslT-LT1 transfection reagent.
FACS analysis
For BrdU labeling, U2OS cells were incubated with 10mM BrdU/1 5% DMSO on ice for
30 minutes, washed and incubated at 28.50C for 15 minutes. Cells were fixed in 70%
ethanol at -20*C overnight. Samples for pH3/propidium iodide or BrdU/propidium iodide
FACS analysis were prepared as described (Pozarowski and Darzynkiewicz 2004).
FACS analysis was conducted by FACScan (Becton-Dickinson). DNA content was
quantified by ModFit LT (Verity Software), and pH3 and BrdU staining was quantified
using FlowJo (Tree Star, Inc.).
Immunoprecipitation
293FS cells were lysed in modified RIPA buffer (50 mM Tris-HCL [pH 8.0], 150 mM
NaCl, 1 mM EDTA, 1% NP-40, 0.1% SDS, 0.1 mM PMSF, protease inhibitor cocktail
(Roche) 1 mM NaF, 10 mM p-glycerophosphate (b-GP), 200 [M Na3 VO 4 ) for 30 minutes
in ice. The soluble supernatant was used for immunoprecipitations with anti-GFP
antibody (11814460001, Roche). The immunoprecipitates were washed four times with
RIPA and resuspended in 2X Laemmli Buffer.
Immunofluorescence
U20S cells were plated on coverslips and fixed with cold methanol for 10 minutes at 200C. For PCNA, soluble proteins were pre-extracted prior to methanol fixation as
described (DiMicco, RD et al. 2006). Antibodies used were a-PCNA (PC10) from Santa
Cruz, a-5H2AX from Upstate (#05-636) and a-RPA (Calbiochem, NA19L). Coverslips
were mounted for imaging with Vectashield containing 1 ug/ml DAPI on a glass
139
microcope slide. Slides were examined using a Zeiss Axioplan 11confocal microscope.
Antibodies
The following antibodies were used in this study: phosho-H3 (sc-8656-R, Santa Cruz),
TopBP1 (NB100-217, Novus Biologicals and sc-32923, Santa Cruz), GFP
(11814460001, Roche), PCNA (PC10, Santa Cruz), 6H2AX (Upstate #05-636) and RPA
(Calbiochem, NA1 9L).
Acknowledgements
I would like to thank members of the Lees lab for helpful comments and discussions.
This work was supported by Ruth L. Kirschstein NRSAs to N.C. J.A.L. is a Daniel K.
Ludwig Scholar.
References
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