Replication Fork Stability in
Mammalian Cells
Ingegerd Elvers
Department of Genetics, Microbiology and Toxicology
Stockholm University
Cover photo Ingegerd Elvers and mang
© Ingegerd Elvers, Stockholm 2011
ISBN 978-91-7447-270-7
Printed in Sweden by Universitetsservice US-AB, Stockholm 2011
Distributor: Department of Genetics, Microbiology and Toxicology
Nothing in life is to be feared,
it is only to be understood.
- Marie Curie
Abstract
Maintaining replication fork integrity is vital to preserve genomic
stability and avoid cancer. Physical DNA damage and altered nucleotide or
protein pools represent replication obstacles, generating replicative stress.
Numerous cellular responses have evolved to ensure faithful DNA
replication despite such challenges. Understanding those responses is
essential to understand and prevent or treat replication-associated diseases,
such as cancer.
Re-priming is a mechanism to allow resumption of DNA synthesis past a
fork-stalling lesion. This was recently suggested in yeast and explains the
formation of gaps during DNA replication on damaged DNA. Using a
combination of assays, we indicate the existence of re-priming also in human
cells following UV irradiation.
The gap left behind a re-primed fork must be stabilised to avoid
replication-associated collapse. Our results show that the checkpoint
signalling protein CHK1 is dispensable for stabilisation of replication forks
after UV irradiation, despite its role in replication fork progression on UVdamaged DNA. It is not known what proteins are necessary for collapse of
an unsealed gap or a stalled fork. We exclude one, previously suggested,
endonuclease from this mechanism in UV-irradiated human fibroblasts. We
also show that focus formation of repair protein RAD51 is not necessarily
associated with cellular sensitivity to agents inducing replicative stress, in
rad51d CHO mutant cells.
Multiple factors are required for replication fork stability, also under
unperturbed conditions. We identify the histone methyltransferase SET8 as
an important player in the maintenance of replication fork stability. SET8 is
required for replication fork progression, and depletion of SET8 led to the
formation of replication-associated DNA damage.
In summary, our results increase the knowledge about mechanisms and
signalling at replication forks in unperturbed cells and after induction of
replicative stress.
List of original publications
This thesis is based on the following publications, which will be referred to
by their Roman numerals:
I
UV stalled replication forks restart by re-priming in
human fibroblasts
Elvers I, Johansson F, Groth P, Erixon K, Helleday T.
Submitted
II
CHK1 activity is required for fork elongation but not
fork stabilisation after UV irradiation
Elvers I, Johansson F, Djureinovic T, Lagerqvist A, Stoimenov I,
Klaus E, Helleday T.
Submitted
III
UV-induced replication fork collapse in DNA polymerase
η deficient cells is independent of the MUS81
endonuclease
Elvers I, Deperas-Kaminska M, Johansson F, Schultz N, Wojcik
A, Helleday T.
Manuscript
IV
The histone methyltransferase SET8 is required for Sphase progression
Jørgensen S, Elvers I, Trelle MB, Menzel T, Eskildsen M, Jensen
ON, Helleday T, Helin K, Sørensen CS.
J Cell Biol. 2007 Dec 31;179(7):1337-45
V
Uncoupling of RAD51 focus formation and cell survival
after replication fork stalling in RAD51D null CHO cells
Urbin SS, Elvers I, Hinz JM, Helleday T, Thompson LH.
Submitted
Table of contents
Introduction...............................................................................................................9
Genomic integrity maintenance ..................................................................................9
DNA damage response .............................................................................................10
Induction of DNA damage....................................................................................11
Ultraviolet radiation ........................................................................................11
Hydroxyurea....................................................................................................12
DNA damage signalling .......................................................................................14
ATM and ATR ................................................................................................15
Replication protein A in DNA damage signalling...........................................17
Damage signalling is a complex network........................................................18
ATR activation after double-strand break induction .......................................18
CHK1 in DNA damage signalling...................................................................19
Cell cycle arrest ...................................................................................................19
G1/S checkpoint ..............................................................................................20
S phase checkpoint ..........................................................................................20
G2/M checkpoint.............................................................................................21
DNA repair...........................................................................................................22
Nucleotide excision repair...............................................................................23
Damage recognition during NER ...............................................................................23
Damage excision and new DNA synthesis during NER ...................................................24
Homology-directed repair ...............................................................................26
DSB damage recognition during HR ..........................................................................27
Strand invasion and branch migration during HR...........................................................28
Repair synthesis during HR .....................................................................................29
Replication fork stabilisation ....................................................................................30
Replication on undamaged DNA..........................................................................30
Replication initiation and origin firing ............................................................30
Assembly of pre-replication complexes .......................................................................31
The existence of dormant origins ...............................................................................33
Firing of a replication origin.....................................................................................34
Replication elongation.....................................................................................35
SV40 replication elongation .....................................................................................36
Eukaryotic replication elongation ..............................................................................37
DNA damage tolerance and replication resumption............................................38
Natural impediments to replication fork progression ......................................39
One-ended DNA double-strand breaks............................................................40
Resolution of a stalled replication fork............................................................40
Resolution by chicken-foot formation .........................................................................41
Template switching by strand invasion .......................................................................42
PCNA polyubiquitination ........................................................................................43
Translesion synthesis.......................................................................................43
Translesion synthesis across UV-induced lesions...........................................................44
Re-priming and firing of dormant origins .......................................................45
Chromatin Remodelling............................................................................................46
Chromatin ............................................................................................................47
Chromatin remodelling ........................................................................................48
Present Investigation ..............................................................................................50
Aim ...........................................................................................................................50
Methodology.............................................................................................................50
Cell lines and cell culturing .................................................................................50
Replication elongation measuring techniques .....................................................51
Replication fork progression using alkaline DNA unwinding.........................51
Alkaline sucrose gradients...............................................................................52
DNA fibre technique .......................................................................................52
DNA damage and signalling measurement techniques ........................................53
Pulsed-field gel electrophoresis.......................................................................53
Immunofluorescence .......................................................................................54
Results and Discussion .............................................................................................55
UV stalled replication forks restart by re-priming in human fibroblasts
(paper I) ...............................................................................................................55
CHK1 activity is required for fork elongation but not fork stabilisation
after UV irradiation (paper II).............................................................................56
UV-induced replication fork collapse in DNA polymerase η deficient cells
is independent of the MUS81 endonuclease (paper III) ......................................56
The histone methyltransferase SET8 is required for S-phase progression
(paper IV).............................................................................................................57
Uncoupling of RAD51 focus formation and cell survival after replication
fork stalling in RAD51D null CHO cells (paper V) .............................................58
Concluding Remarks and Future Perspectives..........................................................59
Acknowledgements .................................................................................................62
References................................................................................................................63
Papers I – V
Abbreviations
(6-4)PP
AraC
ATM
ATR
CDK
CHK1
CHK2
CPD
dsDNA
GG-NER, GGR
HR
HU
NER
NHEJ
ORC
ORI
PFGE
pre-RC
pre-IC
PTM
RNR
RPA
ROS
RPIIo
ssDNA
TC-NER, TCR
TLS
UV
XP
XP-V
γH2AX
pyrimidine pyrimidone (6-4) photo product
cytarabine, cytosine arabinoside
ataxia telangiectasia mutated
ATM and Rad3 related
cyclin-dependent kinase
checkpoint kinase 1
checkpoint kinase 2
cyclobutane pyrimidine dimer
double-stranded DNA
global genome repair
homologous recombination
hydroxyurea
nucleotide excision repair
non-homologous end joining
origin of replication complex
origin of replication
pulsed-field gel electrophoresis
pre-replication complex
pre-initiation complex
post-translational modification
ribonucleotide reductase
replication protein A
reactive oxygen species
RNA polymerase II, elongating
single-stranded DNA
transcription-coupled repair
translesion synthesis
ultraviolet radiation
xeroderma pigmentosum
XP variant cells or phenotype
phosphorylated histone H2AX
Introduction
Replication is the process by which DNA is copied to generate identical
daughter chromosomes, thereby passing on the genetic material to the
daughter cells. The DNA synthesis phase is part of the cell division cycle,
which is necessary for tissue growth and renewal. Some tissues stop
proliferating while others, such as the skin, continue to grow during our
entire lives. Replication, being vital to all aspects of life, is tightly regulated
by the cell. As DNA is constantly exposed to exogenous and endogenous
damaging factors, DNA lesions are frequently occurring. If not repaired
before replication, these perturbed structures may become an obstacle during
the progression of replication. DNA lesions may cause stalling and collapse
of a replication fork structure, and can also give rise to mutations eventually
leading to cancer.
Several DNA repair pathways have evolved to remove DNA damage
and restore the genetic material. Maintaining the integrity of the genome is
essential to avoid cancer, but to some extent, DNA damage may be tolerated
by the cell. However, replication of damaged DNA may induce changes in
the genetic sequence inherited by the daughter cells, the original sequence no
longer identifiable.
DNA damaging agents are frequently used in cancer therapy, which may
seem contradictory as they can also cause cancer. The aim of this study has
been to use well-known DNA damaging agents to further understand the
mechanisms and signalling involved in replication fork stability in
mammalian cells. Additional understanding in this field may be useful in the
struggle to prevent cancer, and to cure such disease, but also in the general
understanding of the function of cells, the diverse components forming our
bodies.
Genomic integrity maintenance
Every single day, tens of thousands of DNA lesions are formed in a
mammalian cell (Beckman and Ames, 1997; Lindahl, 1993). Most lesions
are small, such as the spontaneous oxidation of a nucleotide, while other
lesions may induce loss of genetic material if not repaired correctly. Indeed,
if left unrepaired, DNA damage may cause misregulation of, or be a physical
9
obstacle to, the transcription of genes. Furthermore, if encountered by the
replication fork, DNA lesions may induce mutations or cause loss of genetic
information, and hence they pose a severe threat to the cell and the organism.
Several specialised DNA repair pathways have evolved to take care of the
numerous lesions before replication occurs. In addition, DNA damage may
activate other cellular responses including cell cycle arrest and apoptosis. To
some extent, damage may also be tolerated during replication.
Numerous proteins are involved in maintaining genomic integrity. The
physical connection between sister chromatids, termed cohesion, is achieved
by ring-like cohesion protein complexes which hold the sister chromatids
together. Cleavage of this connection is vital for proper segregation of sister
chromatids during anaphase (Uhlmann et al., 1999). By facilitating DNA
repair, cohesin is important for genomic integrity (Strom et al., 2004; Unal et
al., 2004).
Failure to maintain the genomic material may lead to accumulation of
DNA damage. Loss of genome integrity may also lead to changes in gene
expression, as a result of deletions or duplications. Genomic instability is
associated with diseases such as cancer, Fragile X, and Huntington’s disease.
DNA damage response
Being constantly exposed to many different types of stress and DNA
damaging agents, the cell has evolved several pathways to repair DNA
lesions. A damaged protein can be replaced by a new one, but since the cell
doesn’t have any backup copies of its nuclear genome, DNA repair is vital.
Several different repair pathways, specialising in different types of damage,
are present in eukaryotic cells in combination with fine-tuned sensing
mechanisms. Many types of DNA damage can become more harmful for the
cell if a replication fork runs into the site of damage before repair has taken
place. Ultraviolet radiation emitted by the sun is an important environmental
source of DNA damage. The damaged bases may be detected by the DNA
repair system, and repaired. However, if not repaired before replication, this
will cause an obstacle to the replication machinery. Another agent
influencing replication forks is hydroxyurea, which depletes the nucleotide
10
pool by inhibiting ribonucleotide reductase, thereby stalling replication
forks.
When damage is detected, the cell cycle may need to be halted to allow
repair of the damage. The signalling for repair of DNA and restoration of
chromatin is an intricate system, tightly regulated by the cell. The above
mentioned DNA damage inducing agents as well as damage signalling and
DNA repair pathways are described in more detail below.
Induction of DNA damage
Paracelsus (1493-1541) stated that all compounds can be poisonous, the
toxicity being only a matter of dose. This is a central dogma in the field of
toxicology, as the concentration of a compound affects the response in the
cell. Below, two commonly used agents for induction of DNA damage are
discussed; ultraviolet radiation and hydroxyurea. Ultraviolet radiationinduced DNA damage and hydroxyurea both block the progression of
replication forks, but by different mechanisms. These two agents can be used
to study different and overlapping aspects of replication fork stability and
repair at replication forks.
Ultraviolet radiation
One of the constant exogenous threats to DNA is ultraviolet (UV)
radiation, an electromagnetic radiation emitted by the sun. The energy
carried by the ultraviolet radiation is taken up by the matter with which the
wave collides when reaching the earth's surface. Ultraviolet radiation is
subdivided into UVA (400-320nm), UVB (320-295nm), and UVC (295100nm). UVC and most of the UVB radiation is absorbed by the ozone
layer, and the majority of UV radiation reaching the surface of earth consists
of UVA.
While UVA and UVB primarily cause protein damage, the energy peak
of UVC radiation is preferentially absorbed by DNA. Hence UVC irradiation
induces mainly the same types of lesions as the other ultraviolet radiation
types together, but no protein damage. This makes UVC irradiation a useful
tool for studying DNA repair after sun exposure. The most frequent DNA
11
lesion observed after UV irradiation is cyclobutane pyrimidine dimers
(CPDs), where two adjacent pyrimidines in the same strand become
covalently linked to each other (Kuluncsics et al., 1999), destabilising the
helical structure. This covalent bond can theoretically occur in several
different conformations, but in DNA sterical hindrance allows mainly the
cis-syn CPD to form. In addition, two pyrimidines can be linked by a
covalent bond forming between the C6 of the 5' pyrimidine and the C4 of the
3' pyrimidine, a structure identified as a pyrimidine-pyrimidone (6-4)
photoproduct ((6-4)PP) (Varghese and Wang, 1967). UVC irradiation
induces CPDs and (6-4)PPs in the ratio 3:1 (Mitchell et al., 1990). Both
possible lesions, originating from two adjacent thymidines are shown in
figure 1. (6-4)PPs cause a higher distortion to the DNA backbone compared
to a CPD. A (6-4)PP induces a 44º bending of the backbone (Kim and Choi,
1995), while a cis-syn CPD causes less backbone bending, still allowing
some Watson-Crick base-pairing
(Park et al., 2002). This difference
in backbone distortion makes the (64)PPs more easily recognised by the
cellular DNA repair machinery, and
hence this lesion is more quickly
repaired following UV exposure
even though (6-4)PPs comprise only
a fraction of the total amount of
DNA lesions induced (Mitchell and
Nairn, 1989; Tijsterman et al.,
1999). Additionally, 8-oxo-Guanine
(8-oxoG) is produced following
UVA exposure (Kino and
Sugiyama, 2005).
Figure 1. UV-induced formation of a CPD (left) and
a (6-4)PP (right) in DNA
Hydroxyurea
Hydroxyurea (HU) is occasionally used as an antitumor agent and in the
treatment of HIV infections, reviewed in (Szekeres et al., 1997). Its effect
comes from the inhibition of ribonucleotide reductase (RNR), a rate-limiting
12
enzyme of DNA synthesis. RNR reduces ribonucleotides to
deoxyribonucleotides, the source for deoxynucleotides in the cell. HU is a
radical scavenger and prevents the electron transport from a tyrosine in the
R2 subunit of RNR, which is required for reduction of the 2'-hydroxy group,
as described below. This effect can be used to slow down and eventually
stall replication forks in living cells. As HU is a general radical scavenger, it
may also disturb other free radical chemistry.
Currently, the known RNRs are divided into three classes. Class I is the
only class described to carry out ribonucleotide reduction in mammals. It has
been proposed that the existence of the three classes of ribonucleotide
reductases and their differences in regulation is reflecting the changes in
oxygenic environment during evolution (Poole et al., 2002).
The class I RNRs are tetramers consisting of two nonidentical
homodimers, the larger subunit R1, and the smaller subunit R2. Both are
required for enzymatic activity, although the active site is restricted to the R1
subunit. Ribonucleotide reductase capacity is controlled through cell-cycle
regulated transcription and proteolysis of the two subunits (Bjorklund et al.,
1990; Chabes et al., 2003; Engstrom et al., 1985). For the enzyme to
function, a stable tyrosyl radical is formed in the R2 subunit. This radical is
to be transferred via several other amino acid residues to a cysteine residue
in the active site of the R1 subunit, generating a thiyl radical. The thiyl
radical may transfer the extra electron onto the C3 of a ribose, from where it
is transported via the ribose C2 onto another cysteine residue of the active
site of the R1 subunit, resulting in reduction of the C2. This involves the
formation of several intermediates, which are stabilised by interactions with
other amino acid side chains of the active site.
The same RNR enzyme can reduce ADP, CDP, GDP, and UDP. dUMP
is later methylated and phosphorylated to generate dTTP. The activity of
class I RNRs is controlled by the binding of deoxyribonucleotides to an
allosteric site of the R1 subunit, regulating which ribonucleotides are
reduced (Eliasson et al., 1996).
Hydroxyurea scavenges the free radical of RNR, by a mechanism not yet
completely understood. The result is a decrease in the concentration of the
dNTP pool and stalled replication forks, which eventually collapse
producing double-strand breaks (Lundin et al., 2002). Interestingly,
13
following genotoxic stress, the yeast small RNR subunit has been shown to
redistribute to the cytoplasm, while both human subunits are relocalised
from the cytoplasm to the nucleus in response to UV irradiation (Xue et al.,
2003; Yao et al., 2003).
DNA damage signalling
The DNA damage response (DDR) is a complex network of proteins
activated by induction of DNA damage or replicative stress. In a generalised
view, DNA damage can be recognised by sensor proteins that recruit and
activate transducer proteins that phosphorylate effector or mediator proteins,
inducing cell cycle arrest (Petrini and Stracker, 2003). In addition to cell
cycle arrest, transcription is downregulated in damaged areas by regulation
of DNA methyltransferase DNMT, which methylates CpG islands in
promoters (O'Hagan et al., 2008).
Besides signalling for DNA repair, this signalling cascade may induce
cell cycle arrest to allow time for repair of DNA, or cell death via apoptosis.
In addition, the DNA damage response may induce cellular senescence (von
Zglinicki et al., 2005), a permanent cell cycle arrest that is associated with
ageing (reviewed in (Herbig et al., 2006)). Recently, ageing has been
discussed as an antagonist of cancer, as ageing induced by cell death can be
seen as an alternative pathway to the accumulation of mutations and
subsequent transformation into a malignant cell that characterises cancer
development (recently reviewed in (Hoeijmakers, 2009)). Furthermore, there
are a number of genetic disorders caused by deficiencies in genomic
maintenance that are characterised by increased ageing (Lombard et al.,
2005; Schumacher et al., 2008), e.g. ATM deficiency which causes
premature ageing (Wong et al., 2003). Interestingly, mice expressing a
mutant version of the DNA damage signalling protein BRCA1 show
premature ageing accompanied by an increased risk for cancer (Cao et al.,
2003). In contrast, mice with elevated levels of active p53, also showing
premature ageing, are protected from cancer (Tyner et al., 2002). Expression
of p16INK4a, associated with senescence, elevates with age (Collins and
Sedivy, 2003). Some of the proteins and interactions of the DNA damage
response are summarised in figure 2.
14
UV, replicative stress
DNA DSB
replication uncoupling
MRN
ATM
RPA accumulation
Rad17
9-1-1 clamp
recruitment
ATM recruitment
ATRIP
TopBP1
ATR activation
ATM activation
H2AX phosphorylation
H2AX phosphorylation
Claspin
MRN phosphorylation
RPA phosphorylation
MDC1 recruitment
CHK2 phosphorylation
CHK1 phosphorylation
CtIP
53BP1 recruitment
repair
RAD51 recruitment
cell cycle arrest
homologous recombination
replication
TLS
apoptosis
Figure 2. Schematic and simplified view of the DDR. Dotted lines indicate crosstalk between ATM and
ATR after double-strand break induction.
ATM and ATR
The ataxia-telangiectasia mutated (ATM) and ATM and Rad3-related
(ATR) kinases play a major role in the DNA damage response and are vital
to maintain genomic stability (Shiloh, 2003). They belong to the
phosphatidyl-inositol 3-kinase related kinase (PIKK) family, as do DNAdependent protein kinase catalytic subunit (DNA-PKcs), suppressor of
morphogenesis in genitalia-1 (SMG-1), mammalian target of rapamycin
(mTOR), and transformation/transcription domain-associated protein
(TRRAP). DNA-PK is primarily involved in non-homologous end joining
(NHEJ) mediated repair of DNA double-strand breaks (DSBs) (reviewed in
(Meek et al., 2008)).
In response to a DSB, the MRE11-RAD50-NBS1 (MRN) sensory
complex activates and recruits ATM (Lee and Paull, 2005; van den Bosch et
al., 2003). Under normal conditions, ATM exists as inactive dimers in the
cell (Bakkenist and Kastan, 2003). Upon recruitment to damaged DNA,
ATM is acetylated by TIP60, stimulating ATM autophosphorylation which
results in its disassociation into active monomers (Bakkenist and Kastan,
2003; Sun et al., 2005). The active ATM monomers phosphorylate multiple
effector proteins, including the MRN complex (Gatei et al., 2000; Kim et al.,
1999). The roles for ATM include damage signalling for repair after ionising
15
radiation (IR) and other DSB-inducing agents, and cell cycle control
(recently reviewed in (Derheimer and Kastan, 2010)).
When the replicative polymerases are stalled, either from slowing down
by hydroxyurea treatment or by a direct stalling on one strand by a physical
DNA lesion or adduct, the MCM helicase continues to unwind DNA in front
of the fork. This exposes long stretches of single-stranded DNA (ssDNA),
which are stabilised by rapid binding of replication protein A (RPA). RPA
then acts as a sensor protein and recruits ATR through interaction with
ATRIP, which directly binds RPA-coated ssDNA (Zou and Elledge, 2003).
Like ATM, ATR belongs to the PIKK protein family and these two related
kinases play fundamental roles in the DDR. Although ATM is generally
activated by DSBs and ATR signals in response to replicative stress, the two
proteins have overlapping roles, especially if one is missing or
downregulated (Bartek and Lukas, 2003; Lindsay et al., 1998; Zachos et al.,
2003).
Both ATM and ATR phosphorylate BRCA1 as well as p53 (Cortez et
al., 1999; Siliciano et al., 1997; Tibbetts et al., 1999), thereby promoting
nucleotide excision repair (NER) (Ford and Hanawalt, 1995; Ford and
Hanawalt, 1997). ATR also promotes NER by phosphorylation of and
physical interaction with XPA after UV exposure (Shell et al., 2009; Wu et
al., 2007; Wu et al., 2006) and by promoting repair in S phase (Auclair et al.,
2008). Conversely, NER intermediates activate ATR leading to
phosphorylation of H2AX (Hanasoge and Ljungman, 2007). In addition,
although ATR is the main signalling kinase after UV exposure, ATMdeficient cells show a deficiency in repair of UV-induced lesions (Hannan et
al., 2002).
The checkpoint kinases CHK1 and CHK2 play major roles in the
signalling after DNA damage and replicative stress. Although ATM
primarily signals through CHK2 and ATR through CHK1, there is some
crosstalk between the two pathways (Gatei et al., 2003). In line with the
roles of ATM and ATR, CHK2 is constitutively expressed throughout the
cell cycle and is activated in response to DNA damage (Lukas et al., 2001).
In contrast, the structurally different but functional analogue CHK1 (Bartek
and Lukas, 2003) is preferentially expressed during S and G2 (Lukas et al.,
16
2001), having a constitutive activity that is amplified in response to DNA
damage induction (Kaneko et al., 1999; Zhao et al., 2002).
Replication protein A in DNA damage signalling
RPA is a conserved heterotrimeric protein consisting of three subunits of
different sizes (70, 32, and 14 kDa, respectively), found in all eukaryotes
(Wold, 1997). Binding specifically to single-stranded and not doublestranded DNA (Nasheuer et al., 1992; Wobbe et al., 1987; Wold and Kelly,
1988), RPA plays a crucial role in DNA replication, DNA repair, and DNA
recombination (Wold, 1997). Stabilisation of ssDNA by RPA is required for
the loading of replication factors onto replication origins (Walter and
Newport, 2000), and RPA is involved in the switch from polymerase α to
polymerase δ (Yuzhakov et al., 1999). In addition, in NER the polarity of
RPA helps in the positioning of XPF and XPG at the lesion (de Laat et al.,
1998b). Human RPA has been reported to show a preference for ssDNA but
not for DNA ends in in vitro experiments using oligonucleotides (Ristic et
al., 2003). However, when plasmid DNA was used, no preference was
observed toward either ss or dsDNA (Van Dyck et al., 1998).
In response to replication damage, ATRIP is recruited to RPA-coated
ssDNA, stimulating ATR activation (Cortez et al., 2001; Costanzo et al.,
2003; Zou and Elledge, 2003). The 32 (-34) kDa-subunit of RPA is
phosphorylated by CDKs in the S and M phases of the cell cycle, and
dephosphorylated in late M (Din et al., 1990; Dutta and Stillman, 1992;
Oakley et al., 2003). This phosphorylation affects the binding of RPA to
DNA and replication factors (Oakley et al., 2003). In addition to this, RPA is
also phosphorylated in response to DNA damage (Carty et al., 1994; Liu and
Weaver, 1993; Shao et al., 1999; Zernik-Kobak et al., 1997; Zou and
Elledge, 2003). This modification induces a conformational change,
lowering the interaction of RPA with replication proteins, while interactions
with DNA repair proteins are unaffected (reviewed in (Binz et al., 2004)).
UV induces RPA phosphorylation only during replication (Rodrigo et al.,
2000) in an ATR-dependent manner (Olson et al., 2006) but also involving
DNA-PK (Cruet-Hennequart et al., 2006). In contrast, ionising radiation
induces phosphorylation of RPA during all cell cycle phases, and is
17
dependent on ATM and DNA-PK (Stephan et al., 2009). The UV-induced
phosphorylation of RPA increases in the absence of polymerase η (CruetHennequart et al., 2006).
Damage signalling is a complex network
The signalling events after DNA damage requires a complex network of
interactions. For example, the Rad9-Rad1-Hus1 (9-1-1) clamp activates
ATR by recruiting TopBP1 to the site of DNA damage, thereby facilitating
the phosphorylation of CHK1 (Delacroix et al., 2007; Kumagai et al., 2006;
Lee et al., 2007). However, efficient accumulation of 9-1-1 at damaged DNA
requires ATR-mediated phosphorylation of Rad17 (Bao et al., 2001;
Medhurst et al., 2008), which together with four small RFC subunits acts as
a clamp loader of 9-1-1 at sites of DNA damage (Bermudez et al., 2003;
Ellison and Stillman, 2003; Zou et al., 2002). Like ATR, Rad17 is recruited
by RPA (Zou et al., 2003). Although ATR and Rad17/9-1-1 are recruited to
damaged DNA independently of each other (Kondo et al., 2001; Zou et al.,
2002), they interact with each other in the regulation of CHK1
phosphorylation. Interestingly, Xenopus TopBP1 is required for 9-1-1 clamp
recruitment to stalled replication forks (Yan and Michael, 2009).
ATR activation after double-strand break induction
ATR recognises ssDNA regions present after uncoupling of the
replication fork due to replicative stress (Byun et al., 2005) or formed by
resection of DNA ends (Adams et al., 2006; Jazayeri et al., 2006; Zou and
Elledge, 2003).
In response to DSBs, ATM activates ATR through phosphorylation of
TopBP1, strengthening the interaction between ATR and TopBP1 (Yoo et
al., 2007). This phosphorylation is mediated through MRE11 (Yoo et al.,
2009). Furthermore, after IR exposure, ATM and MRE11 generate ssDNA
regions that, upon RPA binding, recruit ATR to the site of a DSB, leading to
activation of CHK1 (Cuadrado et al., 2006; Jazayeri et al., 2006; Myers and
Cortez, 2006).
18
In yeast, the 9-1-1 complex is also recruited to DSBs, activating
ATR (Kondo et al., 2001; Melo et al., 2001).
CHK1 in DNA damage signalling
ATR phosphorylates Claspin, creating a docking site for CHK1
recruitment allowing phosphorylation by ATR (Kumagai and Dunphy,
2003). CHK1 suppresses origin firing after induction of DNA damage, and is
essential for cell cycle arrest in S and G2 phases (Zachos et al., 2003). CHK1
also prevents hydroxyurea-stalled replication forks from collapse (Syljuasen
et al., 2005). In response to DNA damage, ATR-dependent CHK1 activation
releases CHK1 from chromatin (Smits et al., 2006), allowing it to
phosphorylate downstream targets such as CDC25A inducing a cell-cycle
arrest (Zhao et al., 2002). In response to UV exposure, ATR signals through
CHK1 giving an S phase arrest and preventing replication initiation
(Heffernan et al., 2002) as well as slowing down replication forks (Guo et
al., 2000; Heffernan et al., 2002; Liu et al., 2000). The slowing of replication
forks also requires RAD51 (Henry-Mowatt et al., 2003), and in addition to
this, CHK1 interacts with RAD51 to promote homologous recombination at
stalled replication forks (Sleeth et al., 2007; Sorensen et al., 2005). CHK1 is
also required for proper ubiquitylation of PCNA for translesion synthesis
(Yang and Zou, 2009).
Cell cycle arrest
The cell cycle is driven by non-catalytic, regulatory cyclin proteins in
complex with cyclin-dependent kinases (CDKs). The CDKs are active only
when bound to a cyclin, and can then phosphorylate target proteins. The
activity of Cyclin/CDK complexes is regulated by cyclin levels, CDK
inhibitors, and regulatory phosphorylations (Morgan, 1995; Sherr and
Roberts, 1999; Waga et al., 1994). In response to DNA damage, the cell
cycle is arrested to allow repair. Three major checkpoints where the cell
cycle can be stalled have been found – the G1/S, the intra-S, and the G2/M
checkpoints (figure 3).
19
a
b
CyclinB
CDK1
G0
M
CyclinA
CDK1
CyclinD
CDK4/6
G1
G2
ATM
p53
CHK2
ATR
p38
CHK1
MK2
p16
p21
SMC1
CDC25 phosphorylation
S
CyclinA
CDK2
CyclinE
CDK2
G1/S
checkpoint
intra-S
checkpoint
G2/M
checkpoint
Figure 3. (a) Overview of the cell cycle and the major cyclin and CDK complexes. (b) DNA damage
checkpoints summary. The impact of the three CDC25 proteins in different checkpoints are discussed in
(Bucher and Britten, 2008)
G1/S checkpoint
During G1, CyclinD/CDK4/6 phosphorylates the retinoblastoma (Rb)
protein, releasing E2F and allowing progression through G1 (Sherr, 1996;
Sherr and Roberts, 1999; Weinberg, 1995). At the end of the G1 phase,
Cyclin E levels increase and the CyclinE/CDK2 complex form (Figure 3 a),
being activated through dephosphorylation by CDC25A (Xu and Burke,
1996). Both Cyclin E and CDC25A are transcriptional targets of E2F
(Bartek and Lukas, 2001; Dyson, 1998; Vigo et al., 1999). The active
CyclinE/CDK2 complex marks the transition into S phase.
In response to DNA damage, the cell cycle is temporarily halted before S
phase begins by phosphorylation of CDC25A (Hoffmann et al., 1994). A
slower but more permanent arrest is achieved by p16INK4a induction and p21
activation by p53, leading to inhibition of the Cyclin/CDK complex, thereby
preventing replication of the damaged DNA (Dulic et al., 1994; Serrano et
al., 1993; Xiong et al., 1993). This is known as the G1/S checkpoint (Figure
3 b) (reviewed in (Massague, 2004)).
S phase checkpoint
Once replication has begun, the intra-S checkpoint is activated both by
DNA damage and replication stress (Figure 3 b) (reviewed in (Bartek et al.,
20
2004)), leading to inactivation of CDC25 phosphatases and downregulation
of origin firing (Larner et al., 1999). Although the ATR-CHK1 pathway is
considered to be the main pathway induced after replication-associated DNA
damage, the ATM-CHK2 pathway may also induce CDC25A
phosphorylation, targeting it for destruction (Falck et al., 2001; Sorensen et
al., 2003). ATM also phosphorylates the cohesin subunit SMC1 (Kim et al.,
2002; Kitagawa et al., 2004; Yazdi et al., 2002), constituting a CHK2independent pathway signalling for S phase arrest (Yazdi et al., 2002). This
pathway also requires NBS1 and BRCA1 (Kitagawa et al., 2004; Yazdi et
al., 2002). In addition to the actions of CHK1 and CHK2, the MAPKactivated protein kinase-2 (MK2) is also believed to play a role in the
destruction of CDC25 phosphatases (Falck et al., 2001; Reinhardt and Yaffe,
2009; Sanchez et al., 1997) as part of the intra-S checkpoint. p38-activated
MK2 directly phosphorylates CDC25B and C, inducing cell cycle arrest after
UV exposure (Manke et al., 2005). MK2 is also activated after other DNA
damaging agents (Reinhardt et al., 2007).
G2/M checkpoint
Both the G1/S and the intra-S checkpoints ensure that replication is
stalled until DNA damage has been repaired. DNA damage that has
remained undetected by these checkpoints or has occurred during the G2
phase, may activate the G2/M checkpoint (Figure 3 b). This checkpoint stalls
the cell cycle and prevents entry into mitosis in response to DNA damage
(reviewed in (Bucher and Britten, 2008)). During late G2 phase,
CyclinB/CDK1 complexes start to assemble but are kept inactive by the
Wee1 and Myt1 kinases (Booher et al., 1997; Parker and Piwnica-Worms,
1992). After faithful finishing of DNA replication, CDC25 phosphatases
remove this inhibitory phosphorylation, allowing entry into mitosis
(Dunphy, 1994). In response to DNA damage CHK1, and to some extent
CHK2, phosphorylates the CDC25 phosphatases preventing CyclinB/CDK1
complexes from assembling (Falck et al., 2001; Jin et al., 2008; Schmitt et
al., 2006; Xiao et al., 2003; Zhao et al., 2002). In addition to the cell cycle
arrest caused by DNA damage, anaphase will not begin until all
21
chromosomes are aligned on the metaphase plane (recently reviewed in (Ito
and Matsumoto, 2010).
Both ATM and ATR are crucial for the G2/M checkpoint-induced cell
cycle arrest. CHK1 inactivation abolishes the G2/M checkpoint, resulting in
cell death (Chen et al., 2003). Interestingly, the unspecific kinase inhibitor
caffeine has been shown to inhibit ATM-dependent G2/M arrest after
nitrogen mustard treatment without affecting the other checkpoints (Das,
1987).
DNA repair
Several specialised repair pathways have evolved in the cell. Smaller
base damages such as alkylations and oxidations, as well as apurinic (AP)
sites and single-strand breaks, are repaired by base excision repair. In this
repair pathway, the damaged base is cut out by a glycosylase, after which the
abasic site formed is cleaved by an AP endonuclease and up to a few new
bases are synthesised. In the case of larger lesions that are also restricted to
one strand, such as UV-induced lesions inducing a distortion to the DNA
backbone, the nucleotide excision repair (NER) pathway opens up a bubble
in the DNA and removes about 24-32 bases around the damage, followed by
repair synthesis and ligation. DNA double-strand breaks are repaired either
by non-homologous end joining (NHEJ) or homologous recombination
(HR). In NHEJ the two double-strand ends are trimmed and quickly joined
together, often resulting in a small deletion. HR uses a homologous
sequence, preferably the sister chromatid, as a template for correct repair of
the DNA double-strand break, but since this pathway requires a template it is
mainly used following DNA replication. HR also plays a role in resolving
replication forks stalled at a lesion. The fifth main mammalian repair
pathway is mismatch repair, repairing mismatches induced during replication
of the genome. Some damage can also be removed from DNA by direct
reversal mechanisms such as photoreactivation.
Below, NER and HR are discussed in detail.
22
Nucleotide excision repair
Nucleotide excision repair (NER) is a repair pathway that removes UVinduced CPDs and (6-4)PPs, bulky adducts etc. Deficiencies in NER are
seen in disorders such as xeroderma pigmentosum (XP), cockayne syndrome
(CS), and trichothiodystrophy (TTD), which are all heritable disorders
associated with sunlight sensitivity. The XP patients can be subdivided into
eight complementation groups, out of which seven are defective in different
proteins involved in NER. These proteins are named, as their
complementation groups, XPA to XPG. Similarly, the CS patients lack either
the CSA or the CSB protein, both involved in NER. TTD patients are a
heterogenous group, characterised by brittle hair and growth retardation, due
to defective synthesis of certain genes. Most of the patients show mutations
on their XPD alleles, leading to impaired NER. Schematically, NER can be
divided into five steps; (a) recognition of the lesion, (b) DNA unwinding
creating an open bubble structure, (c) incisions resulting in DNA singlestrand breaks and removal of the damaged region followed by (d) new DNA
synthesis to cover the gap and (e) ligation. In total, about 24-32 bases are
removed (de Laat et al., 1999). The recognition step may differ and NER has
hence been subdivided into two separate pathways, global genome
nucleotide excision repair (GG-NER or GGR) and transcription-coupled
nucleotide excision repair (TC-NER or TCR).
Damage recognition during NER
Removal of UV-induced (6-4)PPs and CPDs exemplifies the difference
in damage recognition by TC-NER, which recognises lesions blocking
transcription, and GG-NER, detecting and repairing lesions that induce a
distortion on the DNA helix. As mentioned previously, UV irradiation
induced (6-4)PPs cause a much stronger distortion on the DNA backbone
compared to the more common lesion, CPDs. These two lesion are removed
with equal efficiency from the transcribed strand while (6-4)PPs are
removed considerably faster than CPDs by GG-NER (Tijsterman et al.,
1999).
XPC-hHR23B has a binding preference for UV-damaged over
undamaged DNA, and can bind both single-stranded and double-stranded
23
DNA (Masutani et al., 1994; Reardon et al., 1996; Shivji et al., 1994),
changing the conformation of DNA around the lesion (Sugasawa et al.,
1998). The binding of XPC-hHR23B to a DNA lesion leads to assembly of
repair proteins and removal of the damaged part in GG-NER. Lesions
inducing only a small backbone distortion, such as CPDs, can be identified
by the E3 ubiquitin ligase complex UV-DDB, consisting of DDB1 and
DDB2 (Takao et al., 1993). The smaller subunit DDB2, also known as XPE
(Nichols et al., 1996), is absent in rodents (Tan and Chu, 2002; Tang et al.,
2000). DNA damage binding by the UV-DDB complex is followed by
recruitment of the SWI/SNF chromatin-remodelling protein complex XPChHR23B (Fitch et al., 2003). The direct recognition of (6-4)PPs explains
why these lesions are removed more quickly from the genome.
However, transcriptionally active DNA has been shown to be repaired
faster than the overall repair rate. This is explained by the specific
recognition of damage being an obstacle to transcription, resulting in repair
by TC-NER. This subpathway of NER results in rapid removal of damage on
the template strand of transcriptionally active DNA, and the repair is
triggered by the stalling of RNA pol II by a lesion (Tornaletti and Hanawalt,
1999). Repair signalling following the stalling of elongating RNA pol II
(RNAPIIo) requires several specific factors, including the CSA and CSB
proteins. CSA is part of a ubiquitin ligase complex also involving Cullin 4A,
which is neddylated by the COP signalosome (CSN) in response to UV
irradiation (Fousteri and Mullenders, 2008). CSB is related to the SWI/SNF
family of ATP-dependent chromatin remodelers, displaying nucleosome
remodelling activity in vitro, and is also active in the initiation of
transcription of certain genes as a response to UV irradiation (Citterio et al.,
2000).
Damage excision and new DNA synthesis during NER
Following recognition of the damage, the same repair factors are
assembled on DNA in both TC-NER and GG-NER. Among the first repair
proteins to be loaded at the site of damage is XPA, believed to verify the
damage, the single-stranded DNA coating protein replication protein A
(RPA), and the nine-subunit protein complex TFIIH (Yokoi et al., 2000),
which plays a role in both GG-NER, TC-NER, and transcription. Similarly
24
to XPC, XPA preferentially binds damaged DNA and recognises various
DNA damages, preferring (6-4)PPs over CPDs (Asahina et al., 1994; Jones
and Wood, 1993; Robins et al., 1991). The TFIIH complex includes the
helicase subunits XPB (ERCC3) and XPD (ERCC2), and is required in both
transcription initiation and nucleotide excision repair to open up the DNA
helix (Evans et al., 1997; Gerard et al., 1991). Both XPB and XPD have
helicase activity: XPB unwinds DNA in the 3' → 5' direction and XPD in the
opposite direction (Schaeffer et al., 1994; Schaeffer et al., 1993). This opens
up a bubble around the lesion, an action requiring TFIIH (Evans et al., 1997)
and in GG-NER also XPC-hHR23B (Evans et al., 1997; Mu et al., 1997).
XPC-hHR23B is not required for TC-NER, possibly because the DNA helix
is already partly opened due to ongoing transcription.
The opening of DNA is followed by cleavage of single-stranded DNA
by the two structure-specific endonucleases XPG (ERCC5) and ERCC1XPF (ERCC4), on the 3' and 5' sides of the bubble, respectively.
XPG is a member of the FEN-1 family of endonucleases, a protein
family known to incise DNA in loops and at other junctions between singlestranded and double-stranded DNA (Harrington and Lieber, 1994), and has
been shown to make a 3' incision in nucleotide excision repair (O'Donovan
et al., 1994). XPG is also required for formation of the open complex (Evans
et al., 1997) and an active-site mutant of XPG has been found to stabilise a
pre-incision complex with XPC-hHR23B, TFIIH, XPA, and RPA (Mu et al.,
1997) although removal of XPG does not prevent TFIIH and XPA from
binding RNAPIIo (Laine and Egly, 2006).
Stabilisation of the open complex by XPG is independent of its catalytic
activity (Mu et al., 1997), and although XPG has to be present for efficient
cleavage by ERCC1-XPF the catalytic activity of XPG is not required
(Wakasugi et al., 1997). Notably, even though some cleavage by XPG in the
absence of ERCC1-XPF has been reported (Matsunaga et al., 1995; Mu et
al., 1996), there is substantial evidence indicating that the cleavage by
ERCC1-XPF occurs before the incision by XPG (Staresincic et al., 2009).
The ERCC1-XPF heterodimer binds to single-stranded DNA protruding in
either a 3' or 5' direction and can incise several different DNA substrates,
including bubbles and stem loops, although a minimal loop size of 4-8
nucleotides is required (de Laat et al., 1998a). The incisions made by
25
ERCC1-XPF are always on the 5' side of the border between duplex and
single-stranded DNA (Sijbers et al., 1996).
Following incision of the damage, new DNA is synthesised by a DNA
polymerase. In vivo and in vitro studies using different chemical inhibitors
suggest that Polδ and Polε, but not Polα, function during NER synthesis
(Coverley et al., 1992; Dresler and Frattini, 1986; Hunting et al., 1991;
Popanda and Thielmann, 1992), and this is supported by the need for PCNA
in repair synthesis to convert UV irradiated nicked repair intermediates to
fully repaired templates (Shivji et al., 1992). The replication factor C (RFC)
protein complex is also required for nucleotide excision repair (Shivji et al.,
1995). It is possible that Polβ seals small gaps left by DNA Polδ or Polε
(Popanda and Thielmann, 1992). After finished DNA synthesis, the resulting
nick at the 5' end of the newly synthesised patch is sealed by DNA ligase I
(Aboussekhra et al., 1995; Araujo et al., 2000).
Homology-directed repair
Homologous recombination (HR) is a general name for several DNA
repair pathways used by the cell to accurately repair DNA double-strand
breaks (DSBs). HR is also used during replication to repair one-ended DSBs
induced by replication forks running into single-strand breaks, and to
overcome lesions stalling the replication fork.
DSBs can be induced by exogenous sources such as γ-irradiation, or
enzymatically for instance by endonucleases or by DNA topoisomerase II.
DSBs induced by a restriction enzyme are rather 'clean', while radiationinduced DSBs are surrounded by other damages such as single-strand breaks
and oxidative damage, or may even be due to a cluster of single-strand
breaks. The nature of the break will affect the repair, however much of the
knowledge about DSB repair comes from studies using the HO endonuclease
in yeast, and from experiments using high doses of γ-irradiation. A
schematic illustration of DSB repair pathways is shown in figure 4. After
induction of the DSB (figure 4 a), the two ends may be ligated in a the quick
but error-prone non-homologous end-joining repair pathway (figure 4 b).
Alternatively, homologous recombination may repair the break without any
loss of information, using the complementary sequence of a sister chromatid
26
as a template for new DNA synthesis. Occasionally, a homologous
chromosome can be used as a template in homologous recombination,
potentially resulting in loss of heterozygosity. HR is initiated by resection of
the 5' ends of the break, creating 3' ssDNA overhangs (figure 4 c), enabling
strand invasion (figure 4 e) and repair by classical HR (figure 4 h - j) or
synthesis-dependent strand annealing (figure 4 f - g). If the DSB is induced
in a repetitive area, the ssDNA overhangs may be directly ligated with each
other in a process termed single-strand annealing (figure 4 d), a homology
directed repair that requires resection but not recombination.
a
b
c3'
d3'
3'
5'
j 3'5'
3'
5'
h3'
5'
3'
5'
3'
5'
5'
5'
3'
5'
e3'
5'
3'
5'
i 3'
5'
3'
5'
f 3'
5'
3'
5'
g3'
5'
3'
5'
Figure 4. The different possible outcomes of double-strand break repair in eukaryotic cells.
DSB damage recognition during HR
The first recognition signal of DNA double-strand break repair in yeast
is the phosphorylation of the SQ motif on histone H2A (γH2A) around the
break. In mammals, the phosphorylation motif is present on the histone H2A
variant H2AX, comprising a minor fraction of the H2A present in the cell
(Kinner et al., 2008; Rogakou et al., 1998). Consequently, the H2A
phosphorylation signal is seen at 50 kbp on either side of a double-strand
break in yeast (Unal et al., 2004), while γH2AX is seen at a 2 Mbp area
around a DNA double-strand break in mammals (Rogakou et al., 1998). In
yeast, new cohesin molecules are loaded onto DNA following a doublestrand break, and the enrichment of cohesion correlates with the γH2A
domain (Strom et al., 2004; Unal et al., 2004). H2AX can be phosphorylated
27
by ATM, ATR, and DNA-PKcs, as part of the DNA damage response. They
further phosphorylate and activate proteins involved in the cell cycle
checkpoint control, to stop the cell cycle progression until the damage is
repaired. The phosphorylation signal on histone H2AX works as a platform
for the recruitment of repair factors to the site of the break. In homologous
recombination, some of the first proteins to be recruited include MRE11,
RAD50, and NBS1, which together form the MRN complex known to
promote 5' → 3' resection in yeast (Ivanov et al., 1994), although the
exonucleolytic activity of MRE11 proceeds in the 3' → 5' direction (Paull
and Gellert, 1998). The resection creates 3' single-stranded overhangs of
several hundred bases (White and Haber, 1990) (figure 4 c), and is
immediately followed by RPA binding to and protecting the single-stranded
DNA (Wang and Haber, 2004). As more DNA is resected, RPA binding
spreads from the double-strand break (Wang and Haber, 2004).
Strand invasion and branch migration during HR
The 3' overhang generated by DNA resection may invade a homologous
sequence (fig 4 e). This requires several proteins, including RAD51, a
homologue of the E. coli protein RecA, and cofactors including RPA
(Baumann and West, 1997). RAD51 binds ssDNA and dsDNA with similar
affinity and forms helical filaments with ssDNA and dsDNA in the presence
of ATP (Benson et al., 1994; Sung, 1994), and has been shown to catalyse
strand exchange between single-stranded and double-stranded DNA in vitro
(Baumann et al., 1996). However, binding of RPA impedes yeast Rad51p
binding, and in vitro studies have shown that if RPA is allowed to bind DNA
before the addition of Rad51p, other factors are needed for branch migration
(Shinohara and Ogawa, 1998). One such factor shown to improve the actions
of Rad51 is Rad52, a protein conserved among eukaryotes (Bezzubova et al.,
1993; Muris et al., 1994). Yeast Rad52 has been shown to bind ssDNA,
dsDNA, and RPA (Shinohara et al., 1998). Human RAD52 proteins interact
with each other, forming heptameric rings with a central channel, where
DNA is bound along the positively charged peripheral groove (Stasiak et al.,
2000). About 4 nucleotides of single-stranded DNA are suggested to bind
each Rad52 subunit (Singleton et al., 2002), yielding a beads on a string-like
pattern (Van Dyck et al., 1998). RAD51 binds RAD52, (Kurumizaka et al.,
28
1999; Milne and Weaver, 1993), and yeast Rad52p has been suggested to
counteract the inhibitory effect of RPA on ssDNA binding by Rad51p (Sung,
1997). RAD51 binding displaces RAD52 from DNA (Van Dyck et al.,
1998). Another factor proposed to facilitate the invasion mediated by
RAD51 is the SNF2-related protein RAD54 (Eisen et al., 1995), inducing
supercoiling of the intact template which facilitates separation of the two
strands to allow invasion (Petukhova et al., 1999; Sigurdsson et al., 2002).
BRCA2 is a large nuclear protein binding RAD51 through its BRC
repeats (Wong et al., 1997), and is involved in mediating RAD51-dependent
strand invasion by transporting RAD51 into the nucleus and regulating its
DNA binding activity (Davies et al., 2001; Moynahan et al., 2001).
Consequently, BRCA2 mutations predispose to breast and ovarian cancers
(Venkitaraman, 2002), supposedly because BRCA2 deficient cells use nonhomologous end joining or single-strand annealing to repair DNA doublestrand breaks, both being error-prone repair pathways (Tutt et al., 2001; Xia
et al., 2001). However, RAD51 foci do form at stalled replication forks even
in the absence of BRCA2 (Tarsounas et al., 2003). Due to its requirement in
homologous recombination, RAD51 can be used as an indicator of
homologous recombination.
Repair synthesis during HR
Once the ssDNA overhang has invaded the dsDNA template, repair
synthesis by replicative polymerases occurs (figure 4 e). In the classical
homologous recombination scheme, a double Holliday junction forms
(figure 4 h) that is resolved resulting in a non-crossover (figure 4 i) or
crossover (figure 4 j) event. Alternatively, the formed heteroduplex may be
opened up, allowing the invading, prolonged strand to reanneal with its
original complementary strand in a process designated synthesis-dependent
strand annealing (SDSA) (figure 4 f, g). Both pathways are reviewed in
(Helleday et al., 2007).
Another double-strand break repair mechanism reported in yeast and
mammalians, is the RAD51 independent single-strand annealing (SSA),
occurring in regions of repeated sequences (Lin et al., 1984). Strand
resection following a double-strand break in such a region results in
overhangs that are partially homologous, and may be annealed (figure 4 d).
29
Removal of the 3' non-homologous overhangs and DNA synthesis to fill in
shorter single-stranded DNA sections results in a repaired DNA molecule
containing a deletion.
Replication fork stabilisation
The double-helical nature of DNA provides the basis of replication,
where the separated strands are used as templates. Accurate and complete
copying of the genome is essential to maintain genomic stability in the cell,
and is hence subjected to tight regulations. The initiation of replication
happens at one point on the bacterial chromosome and on several spots
simultaneously in eukaryotic genomes. The number of origins fired and at
what stage during replication, is tightly regulated in the eukaryotic cell.
Replication on undamaged DNA
Once the cell has passed the G1/S checkpoint, the double helix is
unwound and DNA replication is initiated from origins of replication. This is
illustrated in figure 5. Two replication forks, or replisomes, will emerge in
opposite directions from the origin, creating a bubble on the DNA (figure 5
b). The sister replisomes are uncoupled from each other (Yardimci et al.,
2010).
An origin from which replication has been initiated may not be fired
again until the next round of replication. This is partly regulated through the
levels of cyclin-dependent kinases controlling cell cycle progression.
Replication initiation and elongation are well studied, although the
mechanisms are not yet fully understood, while the termination of replication
forks still leaves a lot to explore.
Replication initiation and origin firing
In bacteria, replication is initiated at one single point in the
circular chromosome. This point is called the origin of replication
(ORI). Mammalian cells, having a much greater amount of DNA, start
their replication at several points on each individual chromosome.
30
Replication is controlled by the selection of such potential replication
origins and the firing of a subset of them. Some origins are fired early
during replication, and others late. The origin of replication in budding
yeast Saccharomyces cerevisiae, is often intergenic (Raghuraman et
al., 2001; Wyrick et al., 2001) and is about 100 bp in length including
a consensus sequence (Palzkill and Newlon, 1988). In fission yeast,
Schizosaccharomyces pombe, replication origins often appear in
clusters where normally only one origin is used in each replication
round (Dubey et al., 1994; Patel et al., 2006). The typical S. pombe
replication origin is about 1000 bp in length and AT-rich (Segurado et
al., 2003) but lacking a clear consensus sequence (Clyne and Kelly,
1995; Dubey et al., 1996; Kim et al., 2001). The increase in A and T is
probably energetically favorable for the cell since A-T pairs are held
together by fewer hydrogen bonds than G-C basepairs, and are thereby
easier to pull apart in the DNA melting process necessary for
replication initiation. Even though no consensus sequence has been
identified in metazoans, certain intergenic regions have been found to
be enriched in the origin-binding proteins, as well as some CpG
islands (Keller et al., 2002; Ladenburger et al., 2002), in striking
contrast to the finding that human replication origins in general are
AT-rich (Falaschi et al., 2007).
Assembly of pre-replication complexes
Pre-replication complexes (pre-RCs) can only assemble at the potential
replication origins when levels of cyclin-dependent kinases (CDKs) are low
and APC/C protein levels are high; that is, from late mitosis and until early
G1 phase (Dahmann et al., 1995; Piatti et al., 1996). Origin firing on the
other hand can only occur following APC/C inactivation and increased CDK
levels (Diffley, 2004). This prevents re-accumulation of pre-RC complexes
at fired origins before the end of S phase. Formation of the pre-RC at
potential replication origins involves the assembly of several proteins, that
are not highly conserved (Falaschi et al., 2007). In total, the pre-RC consists
31
of more than ten different proteins (Gillespie et al., 2001), some of which are
discussed below.
First, the ATPase origin of replication complex (ORC) binds DNA. S.
cerevisiae and S. pombe ORC both interact preferentially with A-rich DNA
sequences, which are typical for the S. cerevisiae autonomously replicating
sequence (ARS) consensus sequences (ACS) (Van Houten and Newlon,
1990) whereas in S. pombe the ORC uses an AT-hook to bind the origin
sequence (Chuang and Kelly, 1999). In contrast, no consensus sequence
have been found for human origins of replication, and the binding of human
ORC to origins could depend on other protein factors or on local chromatin
structure (Bell, 2002). Consistent with the lack of a conserved sequence in
replication origins, in vitro studies have shown that the ORCs from higher
eukaryotes have very little DNA sequence specificity (Vashee et al., 2003).
After binding DNA, the ORC uses the energy of ATP hydrolysis to
recruit several other proteins (Bowers et al., 2004), working as a platform for
the assembly of these additional replication factors. Early proteins to be
assembled at the origins include Cdc6p / Cdc18p / p62 (Cdc6); Tah11p
(Sid2p) / Cdt1p / CDT1, and the Mcm2-7 / Mcm2-7 / MCM2-7 proteins in S.
cerevisiae / S. pombe / human, respectively. Cdc6, CDT1, and MCM2-7
together form the pre-replication complex (pre-RC). MCM2-7 is a sixsubunit ATPase with helicase activity while the main actions of Cdc6 and
CDT1 seem to be regulatory. The expression levels of these proteins are
controlled to prevent re-replication. Cdc6 is an ATP-binding protein
(Neuwald et al., 1999) that is required for association of Mcm2-7 with
chromatin (Coleman et al., 1996). When the cell enters S phase, ScCdc6p is
ubiquitinated and degraded by the proteasome (Drury et al., 1997).
Similarly, the levels of SpCdt1p have been shown to peak in G1 and
decrease when the cell passes through S phase (Bell and Dutta, 2002). In
metazoans, an additional regulatory protein named geminin (McGarry and
Kirschner, 1998) has been described. Human geminin interacts with CDT1,
preventing the loading of the MCM proteins (Wohlschlegel et al., 2000), and
thereby inhibiting the formation of pre-replicative complexes. Consistent
with this, geminin depletion induces re-replication in human cells
(Melixetian et al., 2004). Together, Cdc6 and CDT1, and in metazoans also
geminin, are involved in preventing re-replication while the main function of
32
the MCM complex appears to be its helicase activity. 20 to 40 MCM
complexes have been found to bind at individual origins in Xenopus sperm
DNA (Edwards et al., 2002). The six MCM proteins in the hexamer have
high sequence similarity but they all contain unique and conserved
sequences that distinguish them from each other, and interestingly all studied
eukaryotes appear to have all six MCM protein analogs, one of each class
(Mcm2-7p) (Bell and Dutta, 2002). Additional MCM proteins have also
been found in higher eukaryotes, reviewed in (Maiorano et al., 2006).
The existence of dormant origins
Not all origins bound by the ORC are fired (Raghuraman et al., 2001;
Wyrick et al., 2001), and the exact regulation of this is not known, although
some argue that firing of an origin is a stochastic event (Ge et al., 2007;
Goldar et al., 2008). Loading of the MCM helicase onto an origin is referred
to as origin licensing, since only these origins can be activated (DePamphilis
et al., 2006). Chromatin loops are physically attached to the nuclear matrix,
and the effect of this on DNA replication is reviewed in (Anachkova et al.,
2005). Recently is has been shown that replication fork speed can change the
binding of loops and thereby the firing of origins (Courbet et al., 2008).
Furthermore, human ORC has been shown to interact with the histone acetyl
transferase HBO1, which by acetylating histones can increase the negative
charge on chromatin, opening up the local chromatin structure and possibly
making firing more likely by increasing the accessibility of DNA (Iizuka and
Stillman, 1999). HBO1 also interacts with mammalian MCM proteins
(Burke et al., 2001). Similarly, loss of the budding yeast histone deacetylase
RPD3 or targeting of the histone acetyl transferase Gcn5p results in
accelerated origin firing (Vogelauer et al., 2002). Consistent with this,
targeting of Drosophila Rpd3 close to a replication origin resulted in a
decrease in origin firing (Aggarwal and Calvi, 2004), and an increased
histone acetyl transferase activity is seen as DNA replication is initiated in
mammalian cells (Weiss and Puschendorf, 1988).
Following genotoxic stress, origin firing is decreased through activation
of the checkpoint protein CHK1 (Petermann et al., 2010b; Zachos et al.,
2003). Interestingly, it was recently shown dormant origins close to the
33
induced damage are still fired despite the general decrease in origin
activation (Ge and Blow, 2010).
Firing of a replication origin
Following pre-RC assembly, the origin may be fired, generating two
sister replisomes moving in opposite directions (figure 5 a - b). Vital for the
firing of an origin is the loading of the cell division cycle protein Cdc45p
onto DNA (Kukimoto et al., 1999; Mimura and Takisawa, 1998). The
loading of Cdc45p onto DNA correlates with replication initiation (Aparicio
et al., 1999; Zou and Stillman, 2000), and requires CDK activity (Zou and
Stillman, 1998). Cdc45p loading precedes the loading of DNA polymerases
and is required for the assembly of PCNA and replication protein A (RPA) at
replication forks (Mimura et al., 2000). These proteins are required for
replication elongation, and are discussed below. Following assembly, the
resulting structure at the replication origin is called a pre-initiation complex
(pre-IC) as the origin can now be fired and replication initiated. Pre-ICs are
seen on early fired origins before they are fully assembled on late origins
(Mendez and Stillman, 2003). As the cell progresses through S phase, human
CDC45 is released from chromatin, which might prevent re-replication
(Saha et al., 1998).
Two kinases are involved in the control of replication initiation; CDK1
mentioned above and Cdc7-Dbf4 kinase (DDK). DDK phosphorylates MCM
proteins and other replication factors at replication origins (Foiani et al.,
1995; Nougarede et al., 2000; Sclafani, 2000), and CDK1 phosphorylates
MCM proteins, RPA, and DNA polymerase alpha (Polα) (Dutta and
Stillman, 1992; Foiani et al., 1995; Hendrickson et al., 1996).
Phosphorylation of pre-RC components by CDK1 also prevents reassembly
and re-replication (Coverley et al., 1998; Hendrickson et al., 1996).
In bacteria, replication starts by the denaturation of DNA at the origin of
replication by DnaA, which is called an initiator factor. This allows the
binding of the helicase DnaB in complex with the helicase loading factor
DnaC. DnaA belongs to the AAA+ initiator factor protein family of
ATPases, and in metazoans, melting of the double-stranded DNA helix at the
origin is also believed to be achieved by AAA+ ATPases. Both the ORC
complex and the MCM helicase complex belong to this family (Iyer et al.,
34
2004). The ability of AAA+ ATPases to melt DNA at the origin might be
conserved in all three domains of life (Costa and Onesti, 2008). It is still
unclear whether it is the MCM helicase or the ORC complex that is
responsible for melting the DNA at the replication origin.
a
A
pre RC
Bb
progressing replication forks
Pol ε
Cc
Topo I
Pol δ
RPA
MCM complex
PCNA
RFC
primer
Pol α
Figure 5. (a) Pre-RC assembly allows origin firing. (b) Sister replisomes travel in opposite directions on
DNA. (c) In each replication fork, the leading and lagging strand are coupled together by interactions
between PCNA, replication factor C, and the MCM helicase complex. Figure from (Groth, 2011).
Replication elongation
The unwinding of DNA requires the MCM helicase complex, whose
helicase activity is dependent on the presence of single-stranded DNA (Lee
and Hurwitz, 2001). Also, the helicase activity of the MCM complex have
been reported to be stimulated by thymine-rich sequences (You et al., 2003),
a feature commonly seen at replication origins, see above. The MCM
complex is regulated by phosphorylations (Ishimi et al., 2000; Lei et al.,
35
1997) and by Cdc45p, which might be the critical factor converting the
MCM helicase complex to an active DNA helicase, allowing DNA
replication (Masuda et al., 2003). As Cdc45p is loaded at an origin,
unwinding starts resulting in an opened bubble in DNA (Walter and
Newport, 2000) from which replication proceeds in both directions (figure
5). While MCM complexes are present in large excess at replication origins,
only a fraction of the chromatin-bound MCM helicase complexes recruit
Cdc45p, explaining why Cdc45p is limiting for DNA replication (Edwards et
al., 2002). Cdc45p will stay associated with the MCM complex at each
replication fork (Gambus et al., 2006), being required not only for
replication initiation but also for replication elongation (Hopwood and
Dalton, 1996). The unwinding activity by the MCM helicase complex at
each replication fork requires the prior removal of histones from the
nucleosomes. This is thought to be achieved by the chromatin remodelling
complex ˝facilitates chromatin transactions˝ (FACT), also referred to as SPN
(yeast) or DUF (Xenopus), reviewed by (Tabancay and Forsburg, 2006),
known to promote human RNA polymerase II elongation in vitro
(Orphanides et al., 1998).
SV40 replication elongation
Much of our original understandings about eukaryotic DNA replication
come from biochemical studies of replication of plasmid DNA containing
the Simian Virus 40 (SV40) replication origin. These studies, reviewed in
(Waga and Stillman, 1998), revealed the combined action of two essential
DNA polymerases, Polα and Pol delta (Polδ). Polα, also designated the
primase, makes short RNA primers and extends them on DNA to about 30
nucleotides. This double-stranded DNA template is recognised and bound by
replication factor C (RFC), which then expels Polα and facilitates the
loading of proliferating cell nuclear antigen (PCNA) (Stillman, 2008).
PCNA is a three subunit ring-shaped clamp that after loading can slide along
DNA, and by binding Polδ, ensuring the progression of DNA replication. On
the SV40 leading strand, priming is only needed to initiate replication at the
origin of replication, while at the lagging strand, short Okazaki fragments are
formed and Polα is needed for initiation of replication of each new fragment.
In eukaryotic cells, Okazaki fragments are about 100-200 base pairs, while
36
in prokaryotes they are about ten times that size.
Eukaryotic replication elongation
Eukaryotes have another essential DNA polymerase, Pol epsilon (Polε),
initially designated Polδ II (Burgers et al., 1990). Originally, contradicting
data was presented regarding the need for the catalytic activity of this
polymerase during replication in S. cerevisiae and S. pombe (D'Urso and
Nurse, 1997; Feng and D'Urso, 2001; Kesti et al., 1999; Navas et al., 1995).
However, in vivo inhibition of human Polε in cell nuclei blocks genomic
DNA synthesis (Pospiech et al., 1999). The presence of a proofreading
exonuclease activity in Polε, as in Polδ, indicates that the DNA synthesis
activity by Polε is actually required for proper replication of genomic DNA.
Recent experiments from the Kunkel and Johansson labs have shed some
light on the separable roles of Polδ and ε. By placing a reporter gene in
either direction and at either side of an origin of replication, a given strand in
the reporter gene will be replicated as either leading or lagging strand,
depending on the orientation of the gene. In yeast mutated in Polδ, mutations
were primarily found on the lagging strand, while the opposite was seen in
yeast mutated in Polε (Nick McElhinny et al., 2008; Pursell et al., 2007).
This idea is supported by the finding that Polδ and Polε edit replication
errors on opposite DNA strands, indicating that they would replicate these
opposite strands (Shcherbakova and Pavlov, 1996), and the reporting of
strand-specific replication of plasmid DNA (Karthikeyan et al., 2000).
Consistent with these findings, Polδ but not Polε proofreads errors made by
Polα (Pavlov et al., 2006). However, it is possible that Polδ can partly take
over if the polymerase activity of Polε is absent. Polδ and Polε are believed
to travel together on DNA, bound by separate PCNA molecules held
together by Replication factor C (RFC) (figure 5 c).
Summarising the actions on the lagging strand, there is a constant flow
of initiation and short DNA elongation by Polα, a switch to and further
elongation by Polδ, and maturation and ligation of the Okazaki fragment also
by Polδ together with the flap structure-specific endonuclease FEN1 which
removes the RNA primer, and DNA ligase which seals the gaps. In addition
to the three replicative polymerases α, δ, and ε, several additional DNA
polymerases have been described in eukaryotes. Pol beta (Polβ) lacks
37
proofreading exonuclease activity and fills in short gaps (Singhal and
Wilson, 1993), and functions in base excision repair (Mullen and Wilson,
1997) and possibly also nucleotide excision repair (Popanda and Thielmann,
1992). Pol gamma (Polγ) has both polymerase and 3' → 5' exonuclease
activity, and is responsible for replication of mitochondrial DNA. There are
also several enzymes with polymerase activity across DNA lesions, so-called
translesion synthesis (TLS) polymerases.
DNA damage tolerance and replication resumption
Of the thousands of damages induced in a cell every day, some will not
be repaired before replication takes place. In addition to the exogenous and
endogenous sources of DNA damage, replication forks encounter natural
replication pause sites, and fork movement may be slowed down for
example by depletion of the nucleotide pool. Several mechanisms have
evolved to handle or bypass lesions at the replication fork. A replication fork
stalled for a short time may restart after removal of the nucleotide depriving
agent (Petermann et al., 2010a). If stalled for a long time, the replication fork
collapses into a DNA double-strand break, detectable by pulsed-field gel
electrophoresis (Lundin et al., 2005; Petermann et al., 2010a; Saintigny et
al., 2001). However, there is also evidence that after a physical block to
replication fork progression, replication is resumed on the 5' side of the
lesion, leaving a daughter-strand gap that is later filled in (paper I, (Lopes et
al., 2006)).
The mechanisms for continuation of replication despite DNA damage,
are often termed DNA damage tolerance. It is important to note that the
mechanisms for damage tolerance at replication forks, with the exception of
the repair of DNA single-strand breaks, do not remove the damage.
Although these processes were earlier referred to as post-replication repair
(PRR), it is more accurate to designate them as damage tolerance
mechanisms, as repair of the DNA molecule does not take place.
38
Natural impediments to replication fork progression
The progression of a replication fork may be halted by natural
obstructions, such as unusual DNA structures or other proteins (reviewed in
(Mirkin and Mirkin, 2007)). A replication fork may also collide with a
transcription unit, although this is generally avoided by the cell (Brewer and
Fangman, 1988; Deshpande and Newlon, 1996). Head-on collision causes
replication fork pausing (Deshpande and Newlon, 1996) and demise,
inducing transcription-associated recombination (Prado and Aguilera, 2005).
As the DNA polymerase moves about ten times faster than the RNA
polymerase, a replication fork may also collide with the rear end of a
transcription unit, slowing down the replication fork (Deshpande and
Newlon, 1996). It has been suggested that the mRNA may be used as a
primer for continued DNA synthesis on the leading strand after in-line
collision with transcription (Pomerantz and O'Donnell, 2008).
a 3'
5'
3'
5'
b
h
c
i
j
d
e
k
o
l
p
m
f
g5'
3'
5'
3'
n
3'
5'
3'
5'
q 3'
5'
3'
5'
Figure 6. Homologous recombination repair pathways at replication forks. Only non-crossover products
are depicted.
39
One-ended DNA double-strand breaks
Single-strand breaks are induced by γ-irradiation etc, and are also
induced during the nucleotide and base excision repair pathways. A
replication fork running into a single-strand break will form a one-ended
DNA double-strand break (figure 6 c), a lesion that can also be formed by
cleavage of a small lesion blocking the progression of the replication fork
(figure 6 b). In yeast, such a structure has been shown to induce breakinduced replication (BIR), reviewed by (Kraus et al., 2001), while in
mammalian cells, there is little evidence for BIR. Instead, mammalian repair
of one-ended DNA double-strand breaks highly resembles the repair of twoended DNA double-strand breaks by homologous recombination (figure 6,
left column). The repair is initiated by the resection of DNA of the single
arm, forming a 3' overhang (figure 6 d) that is immediately covered by RPA.
Also, single-stranded gaps of the intact DNA molecule are filled (Helleday,
2003). This enables the single-stranded 3' overhang to invade the doublestranded template (figure 6 e), allowing DNA synthesis and leading to the
formation of a Holliday junction (Zou and Rothstein, 1997) (figure 6 f). This
is mostly resolved without crossing-over (figure 6 f, open arrowheads)
(Richardson et al., 1998), leading to sister-chromatid exchanges (figure 6 g).
Consistent with this, cells defective in single-strand break repair have
elevated levels of sister-chromatid exchanges (Caldecott et al., 1992;
Thompson et al., 1990).
Resolution of a stalled replication fork
In an intricate pathway to avoid the fork collapse, a stalled replication
fork may reverse due to positive torsional strain, forming a structure with
four joined double-strands. As this schematically resembles a chicken-foot, it
has been named so (figure 6 i). This switching of templates allows bypass of
the lesion, and possibly leaves time for repair of the damage (Hanawalt,
2007; Johansson et al., 2004). Alternatively, template switching may occur
through invasion of a sister chromatid (Branzei and Foiani, 2007).
40
Resolution by chicken-foot formation
The chicken-foot structure allows DNA synthesis with the
complementary, newly synthesised strand as a template. However, the
formation of a chicken-foot involves the formation of an intermediate
structure where only the non-damaged, newly synthesised strand has
reversed (figure 6 h). This structure, a half chicken-foot, may be cleaved by
endonucleases, creating a one-ended DNA double strand break (figure 6 c)
repaired as previously described.
Alternatively, if the chicken-foot is formed (figure 6 i), the Holliday
junction it constitutes (figure 6 j) may be resolved by two pathways. Either
replication is restored before the Holliday junction is resolved, generating a
double Holliday junction (figure 6 o), or the Holliday junction is cleaved
resulting in a one-ended DNA double-strand break (figure 6 k). As described
previously, this one-ended double-strand break will invade the intact
template, restoring replication and creating a second Holliday junction
(figure 6 l). Cleavage of this (figure 6 m) generates a restored replication
fork (figure 6 n). Alternatively, a restored fork where replication can
continue may also be obtained dissolution of the double Holliday junction (6
p → 6 q), by the BLM / TopoIII complex (reviewed in Cheok et al roles of
the bloom’s syndrome helicase 2005). In line with this, cells deficient in the
BLM helicase show elevated levels of sister chromatid exchanges, and BLM
suppresses crossing-over in vitro (Wu and Hickson, 2003). However, the
regulation between crossing-over and non crossing-over during resolution,
and between resolution and dissolution, is not yet fully determined.
The resolution of a stalled replication fork and the repair of one-ended
DNA double-strand breaks involve to a large extent the same proteins as for
repair of two-ended DNA double-strand breaks by homologous
recombination. RAD51 foci can hence be used to measure homologous
recombination at replication forks. Consistent with this, depletion of
replication protein A (RPA) inhibits the formation of RAD51 foci at
replication forks following replication stress (Sleeth et al., 2007).
41
Template switching by strand invasion
Template switching by strand invasion has mainly been described in
yeast (Branzei and Foiani, 2007), although it has been shown to occur also in
mammalian cells (Blastyak et al., 2010). Template switching is mainly
discussed as a means of sealing gaps left behind a (possibly re-primed)
replication fork (figure 7 a, f). Without reversal of the replication fork, the 3'
end next to the lesion invades the double-stranded sister chromatid, using the
complementary, newly synthesised strand as a template for DNA synthesis
(figure 7 b, g). This creates a pseudo double Holliday junction (figure 7 c, h),
that can be dissolved to restore normal replication fork progression (figure 7
d, k). However, the lesion stalling the replication fork will remain
unrepaired.
It is unclear to what extent template switching also occurs to allow
progression of a stalled replication fork (figure 7 e). In such a case, strand
invasion (figure 7 i) creates a pseudo single Holliday junction (figure 7 j)
that could be resolved by dissolution to a stabilised replication fork (figure 7
k) still containing the initial damage.
e
a
f
b
i
g
j
c
h
d
k
Figure 7. Model for template switching to allow lesion bypass at replication forks
42
PCNA polyubiquitination
PCNA polyubiquitination on lysine 164 has been implicated in
error-free template switching (reviewed in (Ulrich, 2007) and is
performed by E2 Ubc13-Mms2 and E3 ligase Rad5 in yeast (Hofmann
and Pickart, 1999). Mammalian orthologs of yeast Rad5, HLTF and
SHPRH, induce polyubiquitination of PCNA, thereby preventing
spontaneous mutations (Motegi et al., 2008; Motegi et al., 2006; Unk
et al., 2006). Interestingly, these two enzymes are not essential for
PCNA polyubiquitination, indicating the existence of an alternative
E3 ligase (Krijger et al., 2011).
Translesion synthesis
In eukaryotes, ten DNA polymerases have been described, reviewed in
(Hubscher et al., 2002), out of which five are less stringent than the normal
replicative polymerases. These are described to be involved in translesion
DNA synthesis (TLS) where the replicative polymerase stalled at the site of
a lesion, is exchanged for a DNA polymerase with a more open active site
that is able to replicate past the lesion, potentially inducing a mutation but
preventing the fork from collapse. Due to low processivity the TLS
polymerase quickly falls off the template. The normal replicative polymerase
is then restored on DNA and replication continues.
Translesion synthesis can be divided into two separate processes,
insertion opposite the damaged base and extension of the DNA strand from
the damage. Some TLS polymerases can perform both activities, while
others can only do one. Of the five TLS polymerases described in
eukaryotes, Pol zeta (Polζ) is the only one that does not belong to the Y
family, but is instead part of the B family of DNA polymerases. Polζ is a
heterodimer of the catalytic subunit Rev3 and the regulatory subunit Rev7.
The Y family of TLS polymerases consists of polymerases eta (Polη),
iota (Polι), kappa (Polκ), and REV1, although the latter differs from the
others by not being an actual polymerase, but a dCMP transferase. Due to its
structure it is, however, considered to be part of the Y family together with
Polη, Polι, and Polκ, all being characterised by a more open structure at their
43
active site (Trincao et al., 2001) and with a conserved N-terminal catalytic
domain and an unconserved C-terminal part (Lehmann et al., 2007).
Translesion synthesis across UV-induced lesions
Polη is perhaps the most characterised of the TLS polymerases. This
polymerase appears to differ from the other, non-specific low fidelity TLS
polymerases in that it has specially evolved to replicate correctly past the
most common UV-induced thymine dimer, the cis-syn TT CPD, although it
may also bypass other lesions. When the replication fork runs into a DNA
lesion, it will initially be stalled while the helicase continues, creating a
stretch of ssDNA in front of the stalled fork. This ssDNA is immediately
bound by RPA, and is thought to be the signal responsible for ubiquitination
of PCNA seen at replication forks stalled at a UV-induced lesion
(Kannouche et al., 2004; Lehmann et al., 2007). The ubiquitination mark on
K164 on PCNA increases its affinity for Polη, and as the normal replicative
polymerase falls off the strand, Polη bypasses the CPD (Masutani et al.,
1999a; Masutani et al., 2000), dissociating from DNA after insertion of only
a few nucleotides (McCulloch et al., 2004). Deubiquitination of PCNA is
performed by USP1, which is degraded following UV-irradiation (Niimi et
al., 2008). Since Polη preferentially inserts A, the bypass of a TT-CPD is
normally error-free, but Polη has higher error rates at the 3' thymine than at
the 5' thymine (McCulloch et al., 2004). Although Polη is required for errorfree bypass of CPDs, TLS after UV exposure also involves Polζ (Takezawa
et al., 2010), REV1 (Clark et al., 2003; Gibbs et al., 2000; Takezawa et al.,
2010), and Polι (Dumstorf et al., 2006; Tissier et al., 2000). Interestingly,
depletion of Polκ delayed TLS after UV irradiation (Takezawa et al., 2010).
(6-4)PPs are believed to be bypassed by the combined actions of Polι
and Polζ (Johnson et al., 2000; Tissier et al., 2000; Zhang et al., 2001).
In addition to CPDs and (6-4)PPs, UV-irradiation also induces 8-oxoG
lesions, which are primarily repaired by short-patch base excision repair. If
not repaired before replication of DNA, 8-oxoG has been suggested to be
bypassed by replicative polymerases (Tolentino et al., 2008) or by TLS
polymerases including Polη (Avkin and Livneh, 2002; Maga et al., 2007). It
is possible that Polη is not required for bypass of 8-oxoG but may be
44
involved if present.
Recent data from yeast indicates that translesion synthesis-dependent
filling of gaps after replication on damaged DNA occurs to some extent after
chromosomal replication is finished (Daigaku et al., 2010; Karras and
Jentsch, 2010). In line with this, there is direct evidence of both REV1 and
Polζ being involved not only in bypass during replication but also in
mutagenic DNA synthesis during post-replication filling of gaps after S
phase (Jansen et al., 2009a; Jansen et al., 2009b; Waters and Walker, 2006).
Individuals mutated in Polη display high sensitivity to UV irradiation,
and are categorised as xeroderma pigmentosum variant (XP-V). Unlike
patients from the other XP complementation groups, which are deficient in
NER, Polη deficient individuals have functioning NER (Hessel et al., 1992),
and their UV sensitivity was puzzling for a long time. It was not until 1999
that XP-V patients were found to have a mutated Polη (Johnson et al., 1999;
Masutani et al., 1999b).
Following UV exposure, Polη is phosphorylated by ATR (Gohler et al.,
2011). This phosphorylation is dependent on physical interaction with
RAD18 but is independent of PCNA, and is involved in the checkpoint
response after UV exposure (Gohler et al., 2011).
Re-priming and firing of dormant origins
A stalled replication fork (figure 8 c) may restart by direct resumption of
DNA synthesis if the fork-blocking agent is removed (Petermann et al.,
2010a). Also, if the general replication is slowed down, the cell may fire
more origins to keep the total replication speed up. To prevent re-replication,
only origins licensed before replication, can be fired. Origin firing is
suppressed by CHK1 (Petermann et al., 2010b; Zachos et al., 2003), which is
phosphorylated by ATR in response to replicative stress (Guo et al., 2000;
Liu et al., 2000).
However, if a replication fork becomes completely stalled by a DNA
lesion (figure 8 d), general increase of origin firing may not be the ultimate
choice. Rather, in yeast cells, when a replication fork encounters a DNA
lesion and one strand becomes stalled, leading and lagging strand are
uncoupled and replication continues on the undamaged strand (Lopes et al.,
45
2006), leaving a single-stranded stretch of DNA on the other strand. This
ssDNA signals for ubiquitination of PCNA (Kannouche et al., 2004;
Lehmann et al., 2007). However replication is resumed on the damaged
strand, leaving behind a ssDNA gap opposite the lesion (Lopes et al., 2006).
The formation of ssDNA gaps during replication of damaged DNA has also
been reported or interpreted previously in bacteria, yeast and mammalian
cells (Lehmann, 1972; Prakash, 1981; Rupp and Howard-Flanders, 1968;
Svoboda and Vos, 1995), leading to the suggestion of re-priming past the
lesion to allow replication to continue (Lopes a
et al., 2006) (figure 8 e). Indeed, the ssDNA
formed after uncoupling of the leading and
lagging strands may also be sufficient for Polα b
to bind and allow replication to resume from a
new Okazaki fragment, possibly also on the
leading strand. If present, translesion synthesis c
polymerases fill in the gap, however this
activity may be uncoupled from replication
(Daigaku et al., 2010; Karras and Jentsch,
d
2010). It should be noted that although
replication origin firing is generally decreased
after replicative stress, local activation of
e
dormant origins in close proximity to the
damage may occur (Ge and Blow, 2010).
Figure 8. The nature of a replication fork stalling agent affects the outcome. (a, b) Replication fork
elongation on unperturbed DNA. (c) Replication fork stalling by nucleotide deprivation causes uncoupling
of the helicase. (d) Replication fork stalling by a physical damage initially stalls only one strand. (e) Repriming may occur to allow continuation of replication past a physical lesion, leaving behind a ssDNA gap.
Chromatin Remodelling
Before the findings by Gregor Mendel (1822-1884), traits were believed
to be inherited through blood, being diluted as they were passed on. Mendel
was able to show that phenotypical attributes are inherited as separate pieces
of information, later termed genes. Coinciding in time with Mendel's work,
the existence of nucleic acids was shown by Friedrich Miecher (1844-1895).
46
However, it was not until the early 1900s that Thomas Morgan Hunt (18661945) was able to show that inherited traits are coded for by genes inherited
on chromosomes. For a long time, the DNA of chromosomes was considered
to hold all the vital information needed. However, the field of epigenetics is
now growing, as the non-DNA parts of a chromosome, such as nucleosome
variants and modifications, have been shown to contain information
affecting for example the expression of nearby genes.
Chromatin
Chromatin is the complex of DNA and proteins that together form
chromosomes. The proteins serve multiple functions in association with
DNA: they protect the vulnerable double-helix against ROS, they make
condensation of the negatively charged DNA possible, and they are used to
organise the genetic material, as a cell only uses a fraction of its entire DNA.
This is a complex task involving multiple protein families. In bacteria,
mitochondria and chloroplasts, all lacking a nucleus, chromatin is generally
found assembled in a certain area, called the nucleoid, reviewed in
(Thanbichler et al., 2005). Here, organisation of DNA is achieved by
histone-like proteins, which regulate the accessibility of the genetic material.
The circular chromosome has been found to be arranged with a central core,
and protruding loops, guaranteeing that even in case of a double-strand
break, all the energy stored in supercoiling of the molecule won't be lost.
In eukaryotes, chromatin structure requires the histone protein family.
Histones are relatively small proteins assembled in octamers, around which
DNA is wrapped, forming nucleosomes. Such octamers are built up by socalled core histones; two H2A-H2B heterodimers and an (H3-H4)2 tetramer.
146 bp of DNA is wrapped around the octamer in less than two turns,
entering and leaving the core particle at about the same place. The created
structure, captured on an electron micrograph in 1973 by Ada L Olins, is
known as beads on a string. In mammals and plants, but not in yeast, linker
histone H1 binds between two nucleosomes, stabilising the structure
(Robinson and Rhodes, 2006). Scaffold proteins lead to further compaction
of chromatin.
47
In the eukaryotic nucleus, the chromosomes are organised as
compartmentalised structures, or territories, reviewed in (Cremer et al.,
2000). Within each territory, certain sections of DNA are more accessible,
while other parts are more condensed. The less closed parts are designated
euchromatin, while the more compacted structures are called
heterochromatin. Although euchromatin is often found in the outer
boundaries of the domains building up a territory, and the closed
heterochromatin is often detected in the less accessible inner parts, there are
numerous exceptions from this, reviewed in (Foster and Bridger, 2005).
Another level of organisation comes from the positioning of the individual
chromosomes. Generally, gene-rich chromosomes are often located in the
middle of the nucleus where more transcription factors etc are also found,
while gene-poor chromosomes are often seen in the outer parts of the
nucleus (Boyle et al., 2001). However, important exceptions to this general
rule include transcriptionally active areas located close to nuclear
membranes pores (Casolari et al., 2004).
Chromatin remodelling
The organisation of chromatin is regularly altered by the cell, due to
differences in desired transcription activity, need for DNA repair etc. Such
chromatin remodelling can be divided into stable modifications such as
general changes in hetero- and euchromatin, and rapid, dynamic changes
such as transient, quick signalling for repair where the repair signal has to be
removed once repair is finished. Chromatin remodelling involves
recruitment of nucleosome remodelling complexes, e.g. SWI/SNF, and posttranslational modifications (PTMs) of histone proteins, primarily on the
histone tails. These PTMs regulate both histone-histone interactions within
and between nucleosomes, and histone-DNA interactions. There are several
PTMs such as phosphorylations, acetylations, methylations, ubiquitinations
etc. In some cases the modification itself can be a sterical hindrance for other
modifications to occur, or change interactions by directly influencing the
charge of the modified amino acid, whilst in other cases the modification(s)
act as a recruiting platform for other proteins.
48
Acetylation of lysines by histone acetyl transferases (HATs) is perhaps
the most well studied histone modification, and removes the positive charge
of the lysine being modified. This reduces the affinity between the histone
and the negatively charged DNA, thereby leading to a more open chromatin
structure. Consequently, deacetylation by histone deactylases (HDACs) in an
area causes a more condensed organisation, leading to decreased
accessibility of DNA. The acetylation pattern influences DNA repair
(Masumoto et al., 2005), replication (Alexiadis et al., 1997), and
transcription (Nightingale et al., 1998; Vettese-Dadey et al., 1996).
Methylation of histones by histone methyltransferases does not alter the
charge of the modified amino acid, but rather function through the
recruitment of downstream effector proteins, such as the H3K9 methylation
mark recognised by the chromodomain of heterochromatin protein 1
(Bannister et al., 2001; Lachner et al., 2001). Interestingly, a lysine side
chain can be mono-, di-, or trimethylated, and the three methylation states
can have different biological outcomes, while methylation of arginine, which
can occur once or twice on the side chain, seems to always activate
transcription (Berger, 2007).
49
Present Investigation
Aim
The aim of this thesis was to gain further insight into the cellular
mechanisms and signalling pathways induced by damaged replication forks.
As a tool for this, much focus has been put on xeroderma pigmentosum
variant (XP-V) cells, deficient in the translesion synthesis polymerase Polη,
and the corresponding restored cell line. Some focus has also been put on the
stabilisation of forks stalling after being slowed down, and the signalling
required for stabilisation of forks during unperturbed replication, preventing
damage.
The specific aims were
to study the resumption of DNA synthesis after a physical fork-
stalling lesion (paper I)
to study the role of CHK1 in replication fork stabilisation after
induction of physical fork-stalling lesions (paper II)
to study the role of the MUS81 endonuclease in UV irradiation
induced replication fork collapse in human cells (paper III)
to study the role of SET8 during unperturbed replication (paper IV)
to study the importance of the RAD51 paralog RAD51D during
replication (paper V)
Methodology
Cell lines and cell culturing
Cells were grown in Dulbecco’s minimal essential medium with 10 %
foetal calf serum and 1 % streptomycin-penicillin, in an incubator at 37 °C
with 5 % CO2. The medium used for the restored XP30RO+Polη cell line
was supplemented with 100 μg/ml of zeocin.
The human Polη deficient fibroblast cell line XP30RO (XP-V),
originally derived from a patient and now with an empty pcDNA vector, and
50
the restored XP30RO+Polη cells stably expressing Polη, were kind gifts
from Dr A.R. Lehmann. The Chinese hamster ovary (CHO) cell lines AA8,
51D1, and 51D1.3, were kind gifts from Dr L.H. Thompson.
Table 1. Origin and characteristics of the cell lines used in the study.
Cell line
Origin and cell type
Characteristics
Reference
U2OS
Human osteosarcoma
Wild type
(Ponten and Saksela, 1967)
AA8
51D1
51D1.3
XP30RO
Hamster endothelial
Derived from AA8
Derived from 51D1
Human fibroblast
Wild type
RAD51D knockout
RAD51D complemented
Polη deficient
(Thompson et al., 1980)
(Hinz et al., 2006)
(Hinz et al., 2006)
(Stary et al., 2003; Volpe
XP30RO+Polη
Derived from XP30RO
(empty pcDNA vector)
Polη complemented
and Cleaver, 1995)
(Stary et al., 2003)
Replication elongation measuring techniques
Replication fork progression using alkaline DNA unwinding
The alkaline DNA unwinding (ADU) replication fork progression
technique utilises the fact that DNA in an alkaline solution will unwind from
strand breaks (Erixon and Ahnstrom, 1979). Such strand breaks are present
at a replication fork, and by pulse labelling DNA and allowing the
replication forks to progress from the labelled area for different times, the
average distance that the replication fork has progressed can be measured, as
described previously (Johansson et al., 2004). The fraction of labelled singlestranded DNA is greater the slower the replication fork is progressing.
However, if replication continues past a DNA damage leaving a daughter
strand gap, unwinding will initiate also from this gap.
Briefly, cells are pulse labelled for 30 min with 3H-TdR, exposed to
different doses of UVC irradiation, and incubated in the presence or absence
of inhibitors. Replication fork progression is terminated by the addition of
0.5 ml ice-cold 0.15 M NaCl and 0.03 M NaOH unwinding solution, and
following a regulated incubation unwinding is stopped by the addition of 1
ml 0.02 M NaH2PO4. DNA is fragmented to pieces of about 3 kb, and singleand double-stranded DNA is separated on hydroxyapatite columns at 60 °C
(Erixon and Ahnstrom, 1979). The fraction of single-stranded DNA is
determined in a scintillation counter which converts the radioactive decay to
51
light flashes. Plotting the fraction of single-stranded DNA against chase time
indicates how the labelled area grows into double-stranded DNA as the fork
progresses from the pulse labelled area. This is a quick assay for measuring
replication fork elongation, although it does not distinguish between
different sources of strand breaks.
Alkaline sucrose gradients
Alkaline sucrose gradients were used to study the growth of DNA
molecules synthesised shortly after UV exposure. Briefly, UVC irradiated
cells are chased for 30 min to allow the replication forks to run into a lesion,
before pulse labelling with 3H-TdR for 30 min. Cells are then incubated in
pre-warmed media before harvesting and mild fragmentation of DNA using
γ-irradiation. Irradiated cells were lysed on newly made gradients and
centrifuged to allow separation of small and large DNA fragments. The sizes
of labelled DNA fragments is determined by dropwise transfer of the
gradient to paper strips that are allowed to dry before the radioactive decay is
measured, as described previously (Bienko et al., 2010). This system allows
us to study the growth of molecules replicated shortly after UVC exposure. It
is interesting to note that primary Polη deficient cells have been shown to
replicate their DNA in short stretches that later grow into larger molecules
(Lehmann et al., 1975), while the short fragments formed in immortalised
Polη deficient cells have not been converted to large fragments even at 12 h
after labelling (Paper I). In restored cells, the DNA fragments were of a
considerable size already one hour after labelling (Paper I), indicating that
replication is quicker in immortalised fibroblasts compared to primary cells.
DNA fibre technique
The DNA fibre technique is a useful tool to study replication structures
after DNA damage induction. In contrast to the ADU replication fork
elongation and the sucrose gradient techniques, single molecules are scored,
making it vital to analyse coded samples. In addition to the information
about replication elongation and replication rate, the DNA fibre technique
52
can also be used to study changes in origin firing after damage induction or
in the presence of inhibitors.
Briefly, cells are labelled with halogenated nucleotides (CldU and IdU),
and lysed on glass slides where the DNA is then spread by tilting the slides.
Following fixation, CldU and IdU can be visualised by staining the slides
with specific antibodies (Groth et al., 2010), yielding fibres that are red
and/or green depending on the progression of the fork. Analysing the relative
distribution of different fragments allows us to determine changes in new
origin firing, active forks, etc. The length of the fibre is converted from μm
to kb using the conversion factor 1 μm = 2.59 kb (Henry-Mowatt et al.,
2003), which can be used to measure replication speed. The speed of the
replication fork is generally lower during the IdU treatment in control
samples, possibly because a higher concentration of IdU is used to replace
CldU from the nucleotide pool. Incubating the untreated control cells for 2 h
in IdU yielded very long fragments, likely due to new origin firing and
replicon merging. Such long incubations are not ideal for control samples. In
addition to replication speed, the DNA fibre assay can be used to study
resumption of DNA synthesis (replication fork restart) after induction of
DNA damage.
DNA damage and signalling measurement techniques
Pulsed-field gel electrophoresis
Pulsed-field gel electrophoresis (PFGE) is a method used to separate
large chromosomal fragments on an agarose gel, allowing the visualisation
of DNA double-strand breaks. Furthermore, labelling the cells with 14C-TdR
allows quantification of DSBs specifically formed in replicating (pulse
labelled) or bulk (homogeneously labelled) DNA. PFGE can be used to
study the induction and the repair of DSBs in a large number of cells but is
not a very sensitive method, and the migration of replication structures on
PFGE is not fully determined.
Briefly, cells are homogeneously (24 h) or pulse (0.5 h) labelled with
14
C-TdR directly before exposure to the DNA damaging agent. At chosen
time points, cells are harvested and melted into agarose plugs that are
incubated with proteinase K for 48 h in room temperature (Lundin et al.,
53
2005) before DNA separation on PFGE and staining with ethidium bromide
for visualisation and quantification of DSBs. If DNA was labelled with a
radioactive nucleotide, the gel can then be dried or the DNA can be
transferred onto a membrane to allow quantification of DSBs appearing in
the overall genome or specifically at replication forks.
Immunofluorescence
Studying the formation or disappearance of repair protein focus
formation using immunofluorescence (IF) is a more sensitive method
compared to PFGE, and allows us to study co-localisation of protein foci in
individual cells. Like the fibre assay, analysis using IF requires coding of all
samples to prevent subconscious scoring.
Briefly, cells are grown on cover slips at least 18 hours before induction
of DNA damage. siRNA treatments are preferably not done on cover slips,
but rather in a well before re-seeding of cells onto cover slips. Following
repair, cells are fixed for 20 min in paraformaldehyde of a concentration (1 –
3 %) depending on the cell line. A lower concentration may be required to
reduce background but this shortens the life length of the slides.
For replication stress studies it is useful to pulse label cells with CldU
directly before exposure to e.g. UV, allowing selective quantification of cells
that were in S phase when irradiated, and also to study co-localisation of
repair proteins with areas that were replicating when irradiated. The CldU
foci pattern depends on the S phase stage of the cells.
γH2AX is a common but rather unspecific marker for DSBs and can be
complemented with 53BP1 foci formation quantification for a more accurate
measure of DSB induction. RAD51 was used as an indicator of homologous
recombination repair, although it is an early HR protein and does not
indicate whether the HR event was successful. It should be noted that
background levels of repair protein foci, especially γH2AX, may differ
greatly between cell lines.
54
Results and Discussion
UV stalled replication forks restart by re-priming in human
fibroblasts (paper I)
After replication fork stalling induced by hydroxyurea, replication forks
can actively restart after a short block or use new origin firing to restart after
a longer block. After replication fork stalling by physical agents, replication
cannot be actively restarted until the damage is removed. However, we show
here that replication forks are elongated despite the stalling lesion, despite
fork stalling and a decrease in origin firing.
In this study, using Polη deficient and restored cells, we see UV
irradiation dependent a general reduction in new origin firing and replication
fork movement in the DNA fibre assay, similar in both cell lines. The tiny
difference in fork speed between Polη deficient and proficient cells after UV
exposure could be explained by efficient bypass by other TLS polymerases
in the absence of functional Polη. However, using the alkaline DNA
unwinding method we see an apparent stalling of replication forks in UV
irradiated Polη deficient cells and they remain stalled for 12 hours, while UV
irradiated restored cells display a smaller and only an initial stalling. In
addition, using alkaline sucrose gradients, we demonstrate that new DNA is
replicated in short fragments after UV exposure and that these fragments are
not sealed within 12 hours in the Polη deficient cells. Together, these data
suggest that in response to UV exposure, replication forks are initially stalled
but re-primed close to the damage, allowing replication to continue. This
leaves a gap from which DNA unwinding can start, however the gap is
obviously too small to be detected using the DNA fibre assay. To study the
fate of these replicative gaps, we used pulsed-field gel electrophoresis,
showing an accumulation of DNA double-strand breaks in Polη deficient
cells, likely caused by replication fork collapse. We also show colocalisation of the DNA damage marker γH2AX and recombination protein
RAD51 to the areas that were replicating when irradiated.
This provides evidence of a restart mechanism that is different from the
replication fork restart seen after nucleotide deprivation by hydroxyurea. It is
not surprising that a physical block requires replication restart by different
means compared to a replication fork slowed down due to shortage of
55
nucleotides. It is interesting to speculate in what would be needed for repriming and it is possible that re-priming requires no factors additional to
what is already present at the replication machinery, since re-priming
essentially happens continuously on the lagging strand. Although it is likely
that re-priming occurs also in Polη proficient cells, this remains to be
determined.
CHK1 activity is required for fork elongation but not fork
stabilisation after UV irradiation (paper II)
The ATR-CHK1 signalling pathway is important for replication fork
stability after long hydroxyurea treatment and also regulates origin firing.
Here, we studied the impact of CHK1 on replication fork stability and
replication elongation after exposure to UV. Since the unspecific kinase
inhibitor caffeine, inhibiting ATM and ATR, has been shown to sensitise
Polη deficient cells to UV, we hypothesised that this might be due to an
inability to stabilisation the replication fork via CHK1. To test this, we
looked at continuous replication fork elongation using the alkaline DNA
unwinding technique. We see that caffeine and CHK1 inhibitor further
decrease replication fork speed in UV irradiated Polη deficient and restored
cells. We also see that both agents sensitise the cells to UV irradiation.
However, by pulse labelling cells directly before UV irradiation and then
separating DNA using pulsed-field gel electrophoresis, we can specifically
study the induction of replication associated DNA double-strand breaks.
Using this setup, we show that UV-induced replication fork collapse in Polη
deficient cells is increased by the presence of caffeine, but unaffected by
CHK1 inhibitor. This indicates that CHK1 is important for the continuous
replication fork elongation, but not replication fork stabilisation after UV
exposure.
UV-induced replication fork collapse in DNA polymerase η deficient
cells is independent of the MUS81 endonuclease (paper III)
The MMS and UV sensitive 81 (MUS81) endonuclease has been
implicated in replication fork collapse after treatment with crosslinking
56
agents. We hypothesised that MUS81 may also be involved in replication
fork collapse after exposure to UV. To test this hypothesis, we depleted Polη
deficient and restored cells of MUS81 using siRNA. We see that MUS81
depletion sensitises Polη deficient cells to UV, while restored cells are
unaffected by the MUS81 status. Interestingly, MUS81 depletion did not
affect the formation of replication-associated DNA double-strand breaks
visualised by pulsed-field gel electrophoresis in Polη deficient cells pulse
labelled directly before irradiation. We also studied the formation of γH2AX
and RAD51 foci at replication forks as well as the induction of 53BP1 foci
in response to UV irradiation in these cells. MUS81 depletion had no impact
on the formation of these foci, clearly showing that replication fork collapse
after UV exposure is independent of MUS81.
The histone methyltransferase SET8 is required for S-phase
progression (paper IV)
Histone methyltransferase SET8-mediated monomethylation of histone
H4-K20, as well as a few other histone post-translational modifications,
targets 53BP1 to chromatin after DNA damage. However, the function of
SET8 in mammalian cells was poorly understood. Here, we show that
siRNA-mediated depletion of SET8 causes an accumulation of cells in S
phase, and induction of DNA double-strand breaks. We see that despite the
inhibition of SET8 expression, 53BP1 and other repair proteins are recruited
to the site of DNA damage.
We found it interesting that DNA damage is induced in unperturbed cell
cycles in cells depleted of SET8, and asked what signalling pathways are
affected. We see that CHK1 is activated in SET8 depleted cells, and that
CHK1 inhibition prevents the S-phase delay induced by siRNA-mediated
SET8 depletion. We also see that inhibition of replication proteins or
replication fork stalling, prevents the accumulation of DNA damage induced
by inhibition of SET8 expression. This is explained by SET8 directly
interacting with PCNA, and its requirement for replication fork progression,
as visualised by the alkaline DNA unwinding technique.
57
Uncoupling of RAD51 focus formation and cell survival after
replication fork stalling in RAD51D null CHO cells (paper V)
The homologous recombination repair protein RAD51 assembles at
collapsed replication forks. Here, we studied the involvement of the RAD51
paralog RAD51D in signalling after replication fork stalling, using RAD51D
null cells. We see that RAD51D deficient cells have an increased sensitivity
to camptothecin, but the same sensitivity to replication stalling agents
hydroxyurea and aphidicolin as restored cells. Using pulsed-field gel
electrophoresis, we see no difference in the induction of replicationassociated DNA double-strand breaks after hydroxyurea exposure, in
RAD51D and wild type cells. Despite the unaffected cell survival after
hydroxyurea exposure, RAD51 foci form in control but not in RAD51D
deficient cells. Since other cell lines deficient in homologous recombination
are highly sensitive against inhibition of PARP, we investigated the response
to a PARP inhibitor in RAD51D deficient and restored cells. We see that
PARP inhibition causes a dramatic killing of RAD51D deficient cells and no
induction of RAD51 foci. In contrast, the survival of control cells is
unaffected and RAD51 foci are induced. We found it intriguing that the
induction of RAD51 foci is not related to cell survival in these RAD51D
deficient cells.
58
Concluding Remarks and Future Perspectives
Faithful replication and maintenance of replication fork stability is
essential in preserving genomic stability and avoiding cancer. In the present
thesis we have studied different aspects of replication fork stability and
mechanisms for replication restart after damage induction. We have focused
on UV irradiation since this is an agent which we are exposed to on an
almost-everyday basis, and have also studied other replication-interfering
agents and mechanisms involved in replication stability in unperturbed
replication.
The DNA fibre technique is an elegant method to study replication
elongation and replication restart by analysing single molecules. It gives a
lot of information about replication termination and new origin firing.
However, it has a fairly low resolution, and cells cannot be labelled for very
long, even a 2 hour labelling gives very long fragments in untreated cells.
Thus, it has to be combined with other methods to give a better view of
replication fork elongation. Here, we chose to combine it with the alkaline
DNA unwinding technique and the method of alkaline sucrose gradients,
which both measure whole cell populations. None of the methods is
sufficient to alone describe the dynamics of replication fork progression on
UV-damaged DNA, but the combination gives new views. Pulse labelling
the cells to allow separation of replication-associated DNA double-strand
breaks on pulsed-field gel electrophoresis provides a system to study
replication fork stability. However, it should be kept in mind that replication
bubbles migrate differently in the gel compared to linear DNA. Furthermore,
pulsed-field gel electrophoresis is not an optimal system to study doublestrand breaks after UV exposure since NER induces background levels of
DSBs as an artefact of this method. However, combined with each other,
these methods enable us to study cellular mechanisms and responses after
induction of DNA damage
The re-priming theory has been suggested before but it has not
previously been shown to occur in mammalian cells. It would be of great
interest to study in detail the mechanisms and differences on the leading and
lagging strands. This might be possible using plasmids where the pyrimidine
dimer can only be induced in one place. Furthermore, it remains to be proven
59
whether re-priming is a mechanism only for lesions that cannot be bypassed,
or if it occurs also after induction of damages that can be bypassed by the
present translesion synthesis machinery. The slowing of replication forks in
UV irradiated control cells seen using the alkaline DNA unwinding
technique may represent gaps that are later sealed, or a general reduction of
replication speed on a damaged template despite functional translesion
synthesis. Also, it is surprising to note that the replicative gaps induced in
Polη deficient cells remain unsealed even twelve hours after UV exposure
despite the induction of RAD51 foci seen at six hours after irradiation. Also,
HR defective cells show no delay in replication elongation and gap closure
compared to wildtype cells (Johansson et al., 2004). It remains to be fully
determined whether RAD51 plays a role during a late closure of replicative
gaps, or if homologous recombination is dispensable for this closure. It
would be interesting to immunoprecipitate BrdU-labelled DNA that was UV
irradiated before BrdU pulse labelling and sonicated after replication has
been allowed to continue, to see what repair proteins are involved in the gap
filling after UV exposure. It is possible that combining fluorescent in-situ
hybridisation (FISH) with the fibre technique would also help in elucidating
the role of recombination or template switching and other mechanisms in the
post-replication repair of gaps.
The study of CHK1 signalling indicated a difference in the requirement
for CHK1 in replication fork stability and replication fork elongation. As
CHK1 is one of the main signal transducers in the ATR pathway, and CHK1
is required for replication fork stability after long hydroxyurea exposures, it
may seem surprising that CHK1 activity was not required for stability of
replication forks after UV irradiation. However, it is possible that signalling
for fork stabilisation differs depending on whether the replication fork was
stalled by slowing down, as in the case of nucleotide deprivation, or by a
direct physical block that allows fork continuation on the 5' side of the
lesion. Really, considering the re-priming theory, it is highly likely that
stabilisation of a nucleotide deprivation-stalled replication fork differs from
stabilisation of the replicative gap left behind a re-primed fork. It would,
however, be of great interest to compare the effect of the CHK1 inhibitor to
the effect of inhibitors of ATM, ATR, and other damage signalling proteins.
Elucidating the role of caffeine would be very useful considering how often
60
this unspecific inhibitor is used, however this task is likely to be more
complicated than we first believed.
The importance of the MUS81 endonuclease after different damages is
interesting, not only to determine the role of this endonuclease, but also to
investigate the involvement of different proteins in the physical collapse of a
stalled replication fork. Considering the substrate specificity of MUS81, it is
not too surprising that it is not required for replication fork collapse after
physical blockage, even though the name suggests a role. MUS81 may,
however, have a role in repair of collapsed replicative gaps after UV
exposure, such as in the resolution of Holliday junctions.
These three projects, although being based on cells from a xeroderma
pigmentosum variant patient, has been useful to gain an understanding of the
general mechanisms of replication fork maintenance rather than revealing
new information about the disease. However, this general understanding is
vital to prevent DNA damage and to come up with new cancer treatments,
which will benefit a larger group.
The project involving the histone methyl transferase has a slightly
different angle of understanding the signalling at replication forks following
DNA damage. Post-translational modifications (PTMs) are important not
only for the regulation of transcription, but also for repair signalling
following induction of DNA damage. This new and exciting field has a lot to
reveal about transient modifications working as a local platform for
recruitment of repair proteins or initiation of a signalling cascade.
The project regarding RAD51D concluded that formation of RAD51
foci has no obvious connection to cell survival after treatment with
recombination-inducing agents. This somewhat unexpected result
complicates the studies of cellular responses, and is therefore important for
our understanding of fork stability.
This thesis has aimed to contribute to the general understanding of
replication fork maintenance in mammalian cells. As always, answering one
question enables us to ask ten more.
61
Acknowledgements
My supervisor and co-supervisors.
Professor Thomas Helleday, thank you for your enthusiasm and for being an endless source
of new ideas, and also for giving me the space to work on my own ideas.
Klaus Erixon, my first co-supervisor. Thank you for always being eager to discuss new data.
Fredrik Johansson, my second and main co-supervisor. Thank you for believing in me, for
introducing me to the XP-V project, and for being so supportive especially during those last
hectic months.
Niklas Schultz, my bystander co-supervisor, thank you for all your help with the confocal
microscope.
Thank you all for supporting me during my time as a PhD student, for encouragement
and discussions!
The Helleday team.
Cissi, mitt stöd. Petra, bästa rumskompisen – tack för hjälpen med fibertekniken! Ivo, I miss
you! Vänskapen med er tre är ovärderlig. Evgenia, för att du är så hjälpsam. Diana, vi började
bra som bag ladies. Ann-Sofie och Esther, tack för att ni läste avhandlingen! The DNA repair
subgroup, for all interesting discussions.
Thanks to everyone in the Helleday and Jenssen groups for creating a supportive work
atmosphere!
GMT, Stockholm University.
Hanna, för alla skratt och dina Norén-perspektiv, tack för din vänskap! Anne L, nån gång
kanske vi kan köra BPDE-projektet? Dominik, for once carrying me through a forest during a
stormy night after I broke my foot – new adventures are waiting. Kristian, tack för vänskapen
och grattis till påökningen! Anne B, du livar alltid upp! Daniel, tack för datorhjälp och
träningssällskap! Marta, thanks for joining the MUS81 project and for all always smiling!
Niklas, det är skönt med en vettig människa här... Katten, för att du alltid får mig att le! Sara
P, numera S, nu hoppar vi! Eva P, för att allt plötsligt funkar när du rör vid det.
The Research School in Gene Expression, for financial support. Thanks to the eminent
caretakers, Ann-Sofie and Niklas. Forskarskolevännerna, för alla roliga och intressanta
workshops, med sena middagar och mystiska badmöjligheter!
Svampen, för att ni gör sena labbnätter uthärdliga, och sena icke-labbnätter fantastiskt
roliga!
The GDSC, University of Sussex.
Professor Alan R Lehmann, thank you for welcoming me for an internship in your lab, and
for discussing my data with me.
Dr Simone Sabbioneda, thank you for your patience, for answering all my questions, and for
teaching me the alkaline sucrose gradient technique.
Marcel, Katie, Ann-Sofie, Carol, Phil, Diana, Ted, Ewan, Harley, Yari, and everyone else at
the GDSC and outside the lab. Those pub nights, snowball fights, and the trip to Belgium
were very valuable to me.
Thank you for adopting me when I arrived as an orphan in a new country!
Vännerna, som håller mig uppe.
EEGEEHKB, min klan som snart utökas – tack för uppmuntran, internskämt, och Stenbrohelger! Sis, nu blir det kanske mer tid till pärlande. Mormor, världens finaste! Anna, trots alla
projekt tar du dig alltid tid! Henke, för nattpromenader och en lugnande syn på det mesta. Ida,
för att du är så pepp bara som du är! Hubbe, för att du förstår mig. Mang, tack för att du tog
framsidefotot! Tältgänget, jag längtar till sommaren. Skidgänget, jag längtar till vintern!
Alla ni som kämpar för en bättre värld! Ni är så stora!
62
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