Mutation Research 763 (2015) 15–29
Contents lists available at ScienceDirect
Mutation Research/Reviews in Mutation Research
journal homepage: www.elsevier.com/locate/reviewsmr
Community address: www.elsevier.com/locate/mutres
Review
The Ku heterodimer: Function in DNA repair and beyond
Victoria L. Fell, Caroline Schild-Poulter *
Robarts Research Institute and Department of Biochemistry, Schulich School of Medicine & Dentistry, The University of Western Ontario, London, Ontario N6A
5B7, Canada
A R T I C L E I N F O
A B S T R A C T
Article history:
Received 11 April 2014
Received in revised form 7 June 2014
Accepted 25 June 2014
Available online 4 July 2014
Ku is an abundant, highly conserved DNA binding protein found in both prokaryotes and eukaryotes that
plays essential roles in the maintenance of genome integrity. In eukaryotes, Ku is a heterodimer
comprised of two subunits, Ku70 and Ku80, that is best characterized for its central role as the initial DNA
end binding factor in the ‘‘classical’’ non-homologous end joining (C-NHEJ) pathway, the main DNA
double-strand break (DSB) repair pathway in mammals. Ku binds double-stranded DNA ends with high
affinity in a sequence-independent manner through a central ring formed by the intertwined strands of
the Ku70 and Ku80 subunits. At the break, Ku directly and indirectly interacts with several C-NHEJ
factors and processing enzymes, serving as the scaffold for the entire DNA repair complex. There is also
evidence that Ku is involved in signaling to the DNA damage response (DDR) machinery to modulate the
activation of cell cycle checkpoints and the activation of apoptosis. Interestingly, Ku is also associated
with telomeres, where, paradoxically to its DNA end-joining functions, it protects the telomere ends
from being recognized as DSBs, thereby preventing their recombination and degradation. Ku, together
with the silent information regulator (Sir) complex is also required for transcriptional silencing through
telomere position effect (TPE). How Ku associates with telomeres, whether it is through direct DNA
binding, or through protein–protein interactions with other telomere bound factors remains to be
determined. Ku is central to the protection of organisms through its participation in C-NHEJ to repair
DSBs generated during V(D)J recombination, a process that is indispensable for the establishment of the
immune response. Ku also functions to prevent tumorigenesis and senescence since Ku-deficient mice
show increased cancer incidence and early onset of aging. Overall, Ku function is critical to the
maintenance of genomic integrity and to proper cellular and organismal development.
ß 2014 Published by Elsevier B.V.
Keywords:
Non-homologous end joining
Ku
Telomere
DNA repair
Cancer
Aging
Contents
1.
2.
3.
4.
5.
6.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ku structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ku in DNA repair . . . . . . . . . . . . . . . . . . . . . . . . . .
Non-homologous end joining . . . . . . . . . . .
3.1.
Ku removal . . . . . . . . . . . . . . . . . . . . . . . . .
3.2.
Competition between repair pathways . . .
3.3.
Role of Ku in other DNA repair pathways .
3.4.
Ku as a DNA damage signaling molecule . . . . . . .
Ku at telomeres . . . . . . . . . . . . . . . . . . . . . . . . . . .
Ku in disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Immune system disorders . . . . . . . . . . . . .
6.1.
6.2.
Aging . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
6.3.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
.
16
16
17
17
20
20
21
21
21
23
23
23
23
* Corresponding author at: Robarts Research Institute, Western University, 115 Richmond Street North, London, Ontario N6A 5B7, Canada. Tel.: +1 519 931 5777x24164;
fax: +1 519 931 5252.
E-mail address: [email protected] (C. Schild-Poulter).
http://dx.doi.org/10.1016/j.mrrev.2014.06.002
1383-5742/ß 2014 Published by Elsevier B.V.
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
16
7.
Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1. Introduction
Ku was first identified in the early 1980s as an autoantigen
targeted by autoantibodies in the serum of patients diagnosed
with an autoimmune disease known as scleroderma polymyositis
overlap syndrome [1]. The name Ku comes from the first two
letters of the name of the original patient in whose serum it was
identified. Autoantibodies directed against Ku were subsequently
found in several other autoimmune diseases, including systemic
lupus erythematosus, Sjorgren’s syndrome, polymyositis and
scleroderma [2–4]. Early studies using serum from Ku-positive
patients identified Ku as an abundant, mostly nuclear protein
[1,5]. Subsequent reports showed that Ku had unusual DNA
binding properties, binding avidly to the ends of double-stranded
DNA molecules in a sequence-independent manner, and to a
lesser extent, to other forms of DNA discontinuities such as
hairpins gaps and nicks [5–7]. These unusual end-binding
properties made Ku an appealing candidate for a role in DSB
A
23
24
24
repair and V(D)J recombination, which was confirmed when the
two Ku subunits (XRCC5, Ku80 and XRCC6, Ku70) were found to
complement the DNA repair defect of several IR-sensitive cell
lines [8–14]. It is now well established that while not essential to
individual life in the short term, Ku function is critical to the
maintenance of genomic integrity and to the proper cellular and
organismal development. A better understanding of Ku’s diverse
roles at the cellular and organismal level have implications for the
study and the treatment of other human diseases, such as immune
system disorders, cancer and aging.
2. Ku structure
Ku is a highly abundant protein found in vivo as a stable
heterodimer consisting of two subunits, Ku70 and Ku80 (70 and
80 kDa, respectively). Both Ku70 and Ku80 eukaryotic Ku subunits
contain three domains (Fig. 1A): an N-terminal alpha helix/beta
barrel von Willebrand A (vWA) domain; a central core domain
Ku70
P
P
Human
P
AcAc Ac
vWa
AcAc AcAc
SAP
DNA binding core
NLS
Yeast
Ku80
P P
DNA binding core
vWa
Human
DNA-PK binding P
CTD
NLS
Yeast
Prokaryotes
B
DNA
binding
core
Ku80
vWA
Ku70 vWA
Ku70 SAP
Fig. 1. Representation of Ku structure. (A) Schematic diagram of domain representation of the Ku70 and Ku80 subunits. The subunit domain structure of yeast and human Ku
consists of the alpha helix, beta barrel N-terminal vWA domain, a central DNA binding core and a C-terminal helical domain (CTD). The eukaryotic Ku80 CTD contains the
region required for binding DNA-PKCS, while the Ku70 CTD is shorter and contains a SAF-A/B, Acinus and PIAS domain (SAP). The location of the nuclear localization signals
and post translation modifications (phosphorylation and acetylation) on the human Ku protein is indicated. Yeast Ku is comprised of a similar domain structure to human Ku,
except for a truncated C-terminal domain in Ku80. Prokaryotes encode for a single Ku subunit that is homologous to the eukaryotic core DNA binding domain. (B) The crystal
structure of human Ku (PDB: IJEY) colored according to the domain structure. The dimer forms an asymmetrical basket structure with a positively charged ring large enough
to accommodate two turns of the DNA. The Ku80 C-terminus is not included in this crystal structure.
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
required for DNA binding and dimerization; and a helical Cterminal domain.
The Ku70/80 crystal structure (Fig. 1B) shows that the two
subunits dimerize through the central domain to form a ring
capable of accommodating two turns of double-stranded DNA
(approximately 14 base pairs) [15]. This ring, consisting of
intertwined strands of both Ku70 and Ku80, is lined with positively
charged residues positioned to interact with the sugar phosphate
backbone of DNA in a sequence-independent manner. Ku binds
double-stranded DNA ends with high affinity (Kd 10–9 M),
including 50 -30 or 30 -50 overhangs and blunt ends, however has
significantly less affinity for circular DNA and single-stranded DNA
ends [5,16,17]. Ku has a preferred orientation when loaded onto
the DNA, placing Ku70 proximal to the DNA end, and Ku80 on the
distal side, facing away from the end [15].
The N-terminal vWA domain (also called a/b domain) consists
of a six-stranded beta-sheet in a Rossman fold. The amino side of
the beta-sheet makes contact with the DNA groove, although it
contributes little to Ku70/80 dimerization or DNA binding. The
carboxyl end of the fold points away from the break, which makes
it available as a protein–protein interaction surface [15]. This
domain, named for its prototype protein, the von Willebrand
Factor type A, is present in mostly extracellular matrix proteins but
also several intracellular proteins, where it facilitates protein–
protein interactions with a wide variety of ligands [18]. Overall, the
function of this domain is not well characterized but there is
emerging evidence that it has an important role in DSB repair and
telomere regulation. In Saccaromyces cerevisae, mutations in helix
5 of Ku80 abrogated telomere silencing, while mutations in helix 5
of Ku70 had a negative impact on DSB repair in yeast and mouse
cells [19–21]. Specific residues in the Ku70 vWA domain were also
shown to be critically implicated in conferring abasic site
processing by Ku in vitro [22,23]. Additionally, another region of
the human Ku70 vWA domain in helix 4 was found to regulate DNA
damage signaling to apoptosis [21]. Consistent with its predicted
role as a protein–protein interaction surface, the vWA domain has
been shown to mediate an interaction with both the telomere
complex component telomere repeat binding factor 2 (TRF2) and
NHEJ factor aprataxin and PNKP like factor (APLF) [20,24].
Both Ku subunits contain a helical C-terminal domain (CTD),
however this is the most divergent region of the two proteins. The
Ku80 CTD is approximately 15 kDa and contains a helical and a
disordered region. In vertebrates, the extreme Ku80 C-terminus is
involved in the recruitment of DNA-dependent protein kinase
catalytic subunit (DNA-PKCS) to DNA [25,26]. Lower eukaryotes,
such as Saccharomyces cerevisiae and Aradopsis thaliana, encode for
a smaller Ku80 protein, missing this DNA-PKCS binding region,
which correlates with the fact that DNA-PKCS is not present in
these organisms. The Ku70 CTD is composed of a highly flexible
linker region followed by a structured 5 kDa helix–loop–helix
region known as the SAP domain (named after SAF-A/B, Acinus and
PIAS motifs) [27]. The SAP domain has putative DNA binding
properties and has been shown to increase the overall DNA binding
affinity of the heterodimer [28,29]. This C-terminal region is also
subject to post translational modifications. Multiple C-terminal
lysine residues are acetylated to regulate Ku interaction with proapoptotic proteins [30,31]. Acetylation and sumoylation of Ku70’s
C-terminal tail were also implicated in modulating its recruitment
to DNA damage sites and Ku’s DNA binding affinity, respectively
[31,32]. The Ku70 SAP domain was also shown to mediate the
recruitment of homeodomain proteins to DNA ends [33]. Furthermore, Ku is primarily a nuclear protein, and the CTD in both
subunits contains the basic nuclear localization signal motif that
regulates the heterodimer’s nuclear transport [34].
It is currently unclear whether Ku subunits can exist alone in a
monomeric form, but there is strong evidence that Ku is an obligate
17
heterodimer. Cells derived from Ku70 deficient mice show very
low expression of Ku80, and similarly, the expression of Ku70 is
severely reduced in cells derived from Ku80 deficient mice
[14,35,36]. This phenomenon is also observed in yeast strains
null for either Ku subunit, and in fact, these strains are
phenotypically identical to the double Ku subunit knockout strain
[37]. This is likely due to the instability of each subunit in the
absence of their interacting partner, as exogenous re-expression of
the missing subunit restores the protein levels of its heterodimeric
partner [36]. Overall, evidence suggests that the individual Ku
subunits are unstable and require heterodimerization to function.
The Ku subunits share little primary sequence, but do have a
conserved secondary structure, indicating that they may be
derived from a common ancestor. Ku is an evolutionarily
conserved protein, found in both the prokaryotic and eukaryotic
kingdoms, and its overall structure is preserved throughout. As
several homologues have been identified in bacterial and archaea
species, Ku is thought to have prokaryotic origins [38,39]. For the
most part, eukaryotic genomes contain a single copy of a Ku-like
protein which also functions in a bacterial NHEJ pathway, but as a
homodimer [40]. While having little DNA or protein sequence
similarity to its eukaryotic counterparts, prokaryotic Ku homologues display structural homologies to the central core DNA
binding domain region of both eukaryotic Ku subunits and form
homodimers that bind double-stranded DNA ends [41]. Prokaryotic Ku is much smaller however, only approximately 30–40 kDa,
lacking both the vWA and C-terminal domains found in eukaryotes
[38,39].
3. Ku in DNA repair
3.1. Non-homologous end joining
There are three main DSB repair pathways in eukaryotes
(Fig. 2): the classical NHEJ (C-NHEJ), alternative NHEJ or
microhomology-mediated end joining (A-NHEJ or MMEJ) and
homologous recombination (HR)[42–44]. C-NHEJ and A-NHEJ have
the potential to be active in all stages of the cell cycle, however HR,
is only active in the S and G2 phases because it utilizes the
complementarity of the sister chromatid to repair the DSB with
high accuracy [43,44]. C-NHEJ is capable of ligating any two DNA
ends, regardless of sequence. Due to this flexibility, C-NHEJ is the
predominant DSB repair pathway in humans and other higher
eukaryotes [45–47].
C-NHEJ can be divided into five main stages: (I) recognition of
the break by Ku, (II) Synaptic end bridging, (III) DNA end processing
to produce compatible ends, (IV) ligation of the break, and finally,
(V) Ku removal from the restored DNA. The importance of Ku to
DSB repair has been well documented with reports of Ku deficient
cells displaying inaccurate end-joining, dramatic radiosensitivity,
and chromosomal breakage, translocations and aneuploidy
[14,48–52].
The initial step of C-NHEJ is the rapid recognition of the DSB by
Ku. Ku’s abundance (in humans, approximately 500,000 molecules
per cell [5,15,53]) and strong affinity for DNA allows it to associate
with DNA ends within 5 s of damage [54]. Ku binding has a
protective role and in Ku-deficient cells, nucleolytic processing
occurs at DSB ends [55]. Once Ku has encircled the DNA via its
central ring domain, it directly and indirectly interacts with several
NHEJ factors, serving as the scaffold for the entire NHEJ complex.
Some uncertainty remains as to how many Ku molecules are
recruited to a DNA break. Many DNA damage response factors form
distinct foci easily visible at the site of DNA damage. However there
has been difficulty in observing Ku following DNA damage by
microscopy, leading to the conclusion that Ku does not accumulate
in large numbers at a DSB. Furthermore, an in vitro study showed
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
18
A
Homologous Recombination
B
Alternative Non Homologous End
Joining
MRN
Region of microhomology
RPA
Rad51
Rad54
BRCA2
PARP1
XRCC1
Rad52
Polymerase β
Ligase
I/III
C
Classical Non Homologous End Joining
DSB End Recognition
(1)
Ligation
(4)
Ku70 Ku80
Ligase IV
XRCC4
XLF
Ku80 Ku70
DNA
PKcs
(2)
Synaptic End Joining
P
P
(5)
Ku Removal
Artemis
?
P
P
Ub
Ub
Ub
Processing
(3)
Pol μ/λ
WRN
PNK
APLF
Protease
Fig. 2. Schematics of the double-strand break repair pathways in higher eukaryotes. Homologous recombination (HR) and classical non-homologous end joining (C-NHEJ) are
the two main DSB repair pathways. HR employs a template sequence from a sister chromatid or a homologous chromosome to perform accurate repair of DSBs, and is
restricted to the S and G2 phases of the cell cycle. HR is also involved in the resolution of stalled replication forks and mitotic and meiotic recombination [44]. C-NHEJ directly
joins the two ends of a DSB, regardless of the sequence, and is a simple and rapid process available at any time during the cell cycle [45,46]. C-NHEJ is also involved in the
immunoglobulin gene rearrangements of immune cells through V(D)J recombination and class-switch recombination (CSR)[193]. A third pathway is called by a variety of
names: alternative NHEJ (alt NHEJ or A-NHEJ), microhomology-mediated end joining (MMEJ), and B (backup)-NHEJ. A-NHEJ is a highly error-prone pathway functioning in
case of deficiencies in the C-NHEJ pathway [42]. (A) Homologous Recombination. After the introduction of a DSB, the ends are recognized by the MRN complex (Mre11-Rad50NBS1), which recruits proteins Sae2, Sgs1, Exo1 and Dna2 to facilitate resection of the break in the 50 direction and leave 30 single-stranded extensions. The 30 single-stranded
overhangs are bound by the proteins RPA, Rad51, and Rad54 to form a nucleoprotein filament. The next step, aided by the recruitment of Rad52, is the invasion of this filament
strand into the DNA of the homologous chromosome to form a D-loop structure. A polymerase extends the 30 overhang to create a cross structure known as the Holliday
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
that only one or two Ku molecules were able to load onto a
chromatin substrate [56]. A recently developed method for
visualizing Ku foci also demonstrated that on average there are
only two Ku molecules present at a DSB in vivo, presumably one at
each end of the DNA break [57].
Ku is the DNA binding subunit of the DNA-dependent protein
kinase (DNA-PK) complex, which comprises Ku and DNA-PK
catalytic subunit (DNA-PKCS), a member of the phosphoinositide 3kinase-related kinase (PIKK) family [58]. DNA-PKCS activity is very
important for successful DNA repair, as loss of DNA-PKCS activity
through mutations or knockout, strongly diminishes C-NHEJ
efficiency and, at the organismal level, results in profound
immunodeficiency [10]. The low-resolution electron microscopic
structure of the DNA-PK complex indicates that Ku70/80 makes
several contacts with DNA-PKCS, including the N-terminal and Cterminal regions of DNA-PKCS [59]. At DSBs, DNA-PKCS is recruited
by Ku to form the active complex, leading to Ku translocation and
the eventual activation of DNA-PKCS catalytic activity. This
translocation allows DNA-PKCS to be positioned at the tip of break
and mediates the synaptic joining of the two broken ends and the
stabilization of the complex [60]. Another important consequence
of DNA-PK kinase activation are 15 auto-phosphorylation events
on DNA-PKCS [61–65]. Ku70 and Ku80 were shown to be
phosphorylated by DNA-PK in vitro, however the importance of
these events in C-NHEJ function is unclear, as mutation of these
residues does not impact cell survival after ionizing radiation (IR)
[66]. Similarly, DNA-PK phosphorylates several NHEJ factors in
vitro such as DNA ligase IV, X-ray cross complementing protein 4
(XRCC4), XLF (XRCC4 like factor, also known as Cernunnos) and
Artemis but again many of these were dispensable for NHEJ in vivo
[63,67–69].
There is increasing evidence that, along with DNA-PKCS, Ku
itself mediates the DNA end bridging and stabilization of the CNHEJ ligation complex. Several early observations lead to the
conclusion that Ku functions to bridge DNA ends, including the in
vitro joining of two radiolabelled DNA ends by recombinant Ku
protein and the visualization of Ku dependent DNA fragment
joining by electron and atomic force microscopy [70,71]. More
recently, utilizing an experimental system designed to visualize
DSBs introduced by restriction endonucleases in mammalian cells,
it was observed that the loss of Ku80 resulted in increased distance
between DNA ends [72]. Despite these observations, there has been
little insight into the precise mechanism of Ku end bridging. A
mutation in helix 5 of the Ku70 vWA domain, noted for its ability to
impair DSB repair in both yeast and mammalian cells, was found to
decrease multimerization of Ku proteins, leading to the speculation that this helix mediates DNA end bridging through the binding
of two Ku70 molecules [19–21].
Between the initial break recognition by Ku and the final
ligation of the break, there is considerable flexibility in the factors
involved in repair. For example, IR is known to produce a variety of
damage to the DNA, occasionally leaving non-ligatable 30 phosphate groups, 30 -phosphoglycolates, or 50 -hydroxyl groups.
Therefore, a number of kinases/phosphatases (polynucleotide
kinase/phosphatase (PNKP)), nucleases (Werner, Mre11, Artemis,
ExoI), polymerases (DNA polymerases m and l), helicases (RECQ1),
and phosphodiesterases (tyrosyl-DNA phosphodiesterase 1) may
19
Table 1
A list of proteins proposed to directly interact with Ku, classified according to
cellular process.
NHEJ and V(D)J
DNA-PKCS [228]
Polymerase
m [229]
APLF [24]
TdT [231]
Rag-1 [233]
Polymerase
l [230]
BRCA1 [124]
PARP-1 [237]
RECQ1 [239]
CAF-1 [235]
NBS1 [140]
ABH2 [130]
Mre11 [240]
Kub5-Hera [236]
INHAT [31]
MSH6 [238]
BARD1 [241]
Telomere
TRF1 [183]
TRF2 [184]
TLC1 [166]
TERT [168]
HP1a [242]
Rap1 [186]
Sir3/4 [175]
Mps3 [177]
Apoptosis
BAX [139]
p18-cyclin
E [141]
HDAC6 [142]
SIRT1 [143]
PCAF [30]
Clusterin [243]
Cell cycle
CDK9 [244]
Cyclin A/CDK1
[117]
Transcription and
replication
Oct-1 [33]
HOXC4 [33]
Dlx2 [33]
TOP2A [251]
REF1 [245]
ESE-1 [247]
Runx2 [249]
Dlx5 [249]
XRCC4/Lig4 [76]
XLF [78]
WRN [232]
DNA repair
(other)
g-H2AX [234]
CBP [30]
HIV-IN [246]
Tat [248]
HSF-1 [250]
NAA15 [249]
be required to produce compatible DNA ends [73]. In the case of
DSBs induced by Topoisomerase II (TOP2) poisons, TOP2 remains
covalently linked to via a phosphotyrosyl bond to the 50 terminus
and requires a specific end-processing enzyme, tyrosyl DNA
phosphodiesterase 2 (TDP2), that hydrolyses 50 -phosphotyrosyl
bonds at TOP2-associated DSBs [74,75]. In many cases, recruitment
of these factors to the DNA break is dependent on Ku, with some
factors directly interacting with Ku (Table 1). Interestingly, Ku
itself has been shown to have some enzymatic activity and may
also participate in the processing of DNA ends. It was identified as a
50 -dRP/AP lyase that removes abasic sites by nicking DNA 30 of the
abasic site via a mechanism involving a Schiff-base covalent
intermediate. Several lysine residues in the Ku70 vWA domain
catalyze this reaction, notably K160/164, which when mutated to
alanine residues completely inhibit Ku’s ability to form a Schiff
base [22,23].
Following end-processing, the two DNA ends are ligated back
together. Ku also has an important role in the recruitment of the
ligase complex, which is comprised of DNA ligase IV, XRCC4 and
XLF. The complex requires Ku to be recruited to the break and
forms a direct interaction with Ku, with DNA ligase IV and XLF
interacting with the whole heterodimer and XRCC4 interacting
with Ku70 specifically [54,76–79]. There is still great uncertainty
regarding the stepwise recruitment of NHEJ factors following Ku
binding, although this is not surprising given that different
processing enzymes are required in different situations. Indeed
there is also some evidence that high complexity DSBs (containing
single-strand overhangs, base oxidation and abasic sites) are
repaired with slow kinetics and are dependent on DNA-PKCS
activity, while low complexity DSBs (without surrounding DNA
junction, which is later cut to resolve the chromosome. (B) Alternative end joining/microhomology-mediated end joining (A-NHEJ/MMEJ). The break is recognized by the
MRN complex, PARP1 and XRCC1, which promote the nucleolytic degradation of DNA to reveal 5–25 bp regions of microhomology between the DNA strands. After the
alignment of complementary regions, the gaps are filled in by polymerase b, and ligated by either ligase I or ligase III. (C) Non homologous end joining (C-NHEJ). (1) The ends
are recognized by the Ku heterodimer which slides directly onto the break via its DNA binding ring. The PIKK member DNA-PKCS binds at the Ku80 C-terminus to create the
DNA-PK complex. (2) DNA-PK autophosphorylates to create the catalytically active DNA-PK complex. Artemis is recruited to DNA-PK and phosphorylated. (3) Several
enzymes, including polymerases (polymerase m or l), nucleases, kinases (polynucleotide kinase) are recruited to the break and remove damaged bases or single-strand
overhangs to create compatible ends for ligation. (4) The ligation complex, consisting of Ligase IV, XRCC4, and XLF are recruited by Ku and ligate the two DNA ends. (5) It is
unclear how the Ku dimer is removed from the ligated break. Possible mechanisms include degradation of Ku by proteases and the ubiquitin pathway, or physical nicking of
the repaired DNA by nucleases to allow Ku to slide off.
20
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
damage) do not require DNA-PKCS binding, and are efficiently
ligated with only Ku and the ligation complex [80–83].
3.2. Ku removal
One of the outstanding questions in NHEJ is how Ku is removed
from DNA following ligation of the DSB. Ku is rapidly recruited to
the break, but is only found there transiently, as laser microirradiation studies found that Ku signal steadily depletes within
the few hours following the initial damage, presumably being
removed as DNA is repaired [54,84]. Ku differs from many other
DNA binding proteins in that it binds DNA ends by encircling them
through its ring domain, which would suggest that once the two
DNA breaks are ligated, Ku is trapped on the linear repaired DNA.
Furthermore, the crystal and electron microscopy (EM) structures
do not indicate an obvious escape mechanism given that the vWA
and central ring domain conformations of Ku remain unchanged
whether or not bound to DNA [15,59]. One possible mechanism is
that the removal of Ku from DNA occurs via a protein degradation
pathway. Studies in Xenopus laevis have shown that polyubiquitination of Ku80 Lysine 48 leads to its degradation by the
Skp1-Cul1-Fbxl12 (SCF) E3 ubiquitin ligase complex [85]. Similarly
in human cells, Ku80 ubiquitination was found to be dependent on
the E3 ubiquitin ligase RING finger protein 8 (RNF8) [86]. The
depletion of RNF8 led to increased Ku80 retention at the break and
decreased NHEJ efficiency, suggesting that removal of Ku80 by
ubiquitin-mediated degradation is an important step in successful
DSB repair [86]. Another proposed mechanism for Ku removal is
the direct nicking of DNA to allow Ku escape. Evidence for this has
emerged from yeast systems, with the HR complex MRX (Mre11Rad50-Xrs2) performing an endonucleolytic incision adjacent to
the DNA end, followed by digestion of the DNA to allow Ku removal
and then restoration of the DNA by MRX and Dna2 resection [87–
89]. Overall, protein degradation and direct DNA nicking are very
different possible mechanisms for Ku removal and further work
needs to be done to resolve this discrepancy. It should be noted
that these results were obtained from different biological systems
and could indicate that there is a divergence in Ku removal
mechanisms between yeast and higher eukaryotes.
3.3. Competition between repair pathways
There is much investigation regarding how organisms regulate
the DSB repair pathway choice and increasing evidence suggests
that Ku has an inhibitory effect on the other DSB pathways. Ku is
one of the first proteins found at DSB regardless of cell cycle stage
and cells will first attempt to repair DSBs by C-NHEJ if the ends are
compatible [84,90–92]. HR is the preferred pathway in the S and
G2 phases, and the initial end binding factors of HR, for example
Mre11, antagonize Ku for DNA end binding. The binding of HR
factors initiates DNA end resection to produce single-stranded
DNA, which Ku does not have a strong affinity for, and promotes
the completion of DSB repair by HR [44]. In yeast, Ku appears to
outcompete HR factors in G1 phase, as the loss of Ku results in
increased Mre11 recruitment and Exo1 mediated resection [55,93–
96]. Overexpression of Ku is even able to reduce recruitment of
Mre11 in G2, when HR is the preferred DSB repair pathway [93].
The HR inhibitory effect appears to be specifically dependent upon
Ku’s DNA end binding function, as deletion of other NHEJ factors,
such as ligase IV, were not able to increase HR activity to the same
extent [93]. Another study has implicated, not only Ku binding, but
the kinase activity of DNA-PKCS in the DNA repair pathway choice,
as initiated of HR in G2 depended upon DNA-PKCS autophosphorylation events [97]. Ku70 also antagonizes HR via the Fanconi
Anemia (FA) and break-induced replication (BIR) repair pathways.
The FA pathway repairs DNA interstrand cross-links (ICLs) in
cooperation with the HR pathway during replication [98]. Cell lines
deficient in FA genes display increased sensitivity to DNA
damaging agents that specifically create DNA cross-links and
show a greater dependence on repair by NHEJ. The simultaneous
deletion of Ku70 however, reverses this sensitivity, and repair is
completed by HR once again [99]. Similarly, BIR is responsible for
the repair of breaks that occur in S phase following a replication
fork collapse [100]. This pathway is also dependent on HR, and
yeast cells null for Mre11 activity are sensitive to agents that
induce replication stress [101]. Several Ku70 mutants were
identified that rescue this Mre11 loss, again suggesting a
competitive interplay between Ku and Mre11 [101,102].
A-NHEJ is Ku-independent end joining, and while it is not
currently fully characterized, it is considered to be a more
mutagenic DSB pathway as it occasionally utilizes microhomologies far from the DSB which results in extended resection and
deletions at the repair site [42,43,103,104]. Similar to C-NHEJ, this
pathway is active in all phases of the cell cycle, however it was only
identified after the deletion of essential C-NHEJ components,
suggesting that this pathway is secondary to C-NHEJ
[103,105,106]. It is still unclear why A-NHEJ may be selected over
the less mutagenic C-NHEJ pathway, however A-NHEJ, similar to
HR, often begins with resection to form SSBs, which would inhibit
Ku binding and promote the binding of A-NHEJ factors, such as
PARP-1[42,43,103,104]. Ku outcompetes the DNA binding factor
PARP-1, and deletion of Ku results in increased repair by this
pathway [107–109]. Furthermore the DNA-PK complex has an
inhibitory effect on the enzymatic activity of PARP-1 [110]. Studies
have indicated that human somatic cell lines with Ku80 deletions
retain DSB repair capability, but show a shift toward the A-NHEJ
pathway [111]. However Ku80-expressing cell lines with individual deletions of the other proteins required for C-NHEJ have a
dramatic decrease in all DSB repair pathways [111]. These results
suggest that the binding of Ku, or the entire DNA-PK complex to
DNA has a dominant negative effect on the other DSB repair
pathways if C-NHEJ cannot be completed.
The DNA repair pathway choice is largely determined by cell
cycle stage, with breaks in G1 phase repaired by end joining, and
breaks in G2/S phases largely repaired by HR. Not surprisingly,
members of the cell cycle machinery, namely the cyclin-dependent
kinases (CDKs), have been implicated in regulating the activity of
several DNA repair factors [112,113]. Several studies have shown,
in both yeast and mammalian systems, that CDKs phosphorylate
the HR factors CtBP-interacting protein (CtIP) and Dna2 in order to
initiate the switch to recombination at the G1/S transition [114–
116]. There is some evidence that Ku could also be regulated by
CDK phosphorylation. Cyclin A1/CDK2 was implicated in the
regulation of NHEJ following radiation and was proposed to be a
binding partner of Ku in vertebrates [117]. Furthermore, mass
spectrometry studies revealed potential CDK phosphorylation sites
on Ku [118,119]. However, the functional significance of Ku
phosphorylation by CDKs has yet to be elucidated. While several
potential CDK1 phosphorylation sites were identified in S. cerevisae
Ku, mutation of these sites did not elicit a DNA repair defect [120].
Recent work has also identified the factors p53 binding protein
1 (53BP1) and breast cancer 1 (BRCA1) as essential regulators in
the pathway choice in mammalian cells. 53BP1, along with RAP1
interacting factor (Rif1) and Pax transactivation domain-interacting protein (PTIP), promote NHEJ and negatively regulate resection
in G1 phase [121,122]. BRCA1, along with CtBP-interacting protein
(CtiP), promotes the removal of 53BP1 during the switch to HR at
the G1/S transition [123]. It is currently unclear how Ku fits into
this mechanism, however there is evidence that Ku associates with
BRCA1 and that this interaction is important for successful NHEJ in
G1 [124–126]. It is possible that this association is essential for the
removal of Ku from breaks, with BRCA1 utilizing the exonuclease
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
activity of the MRN (Mre11-Rad50-NBS1) complex in a nicking
mechanism analogous to that proposed for yeast Ku removal.
3.4. Role of Ku in other DNA repair pathways
In addition to its role in the repair of DSBs by C-NHEJ, Ku has
been implicated in a number of other DNA repair pathways. Ku has
been suggested to function in base excision repair (BER), which
repairs DNA base damage, apurinic/apyrimidic (AP) sites and
single-strand breaks (SSBs) [127]. Cells deficient in Ku70 and Ku80
were sensitive to reactive oxygen species (ROS) and alkylation
damage producing agents that result in base lesions and SSBs
[128]. Additionally, in vitro experiments demonstrated that Ku
subunits bind AP sites and cell extracts from Ku deficient cells have
decreased BER activity [128,129]. Furthermore, Ku has been shown
to interact with ABH2, an enzyme involved DNA alkylation repair,
potentially implicating it in this pathway as well [130].
4. Ku as a DNA damage signaling molecule
The DNA damage response (DDR) pathway (Fig. 3) is initiated by
sensor proteins that form large complexes that accumulate in foci
at the site of damage (reviewed in [131]). This complex formation
promotes the activation of a phosphorylation cascade, and the
modification of surrounding chromatin to allow DNA repair factors
to access the break. These initial sensors include the MRN complex,
53BP1, BRCA1, as well as the serine/threonine (S/T) PIKK family
members Ataxia Telangiectasia Mutated (ATM), and Ataxia
telangiectasia and Rad3 related (ATR) [132]. The PIKK members
are the main regulators of the DNA damage response initiated at
the DSB and orchestrate many phosphorylation events at the site of
DNA damage. ATM/ATR phosphorylation of p53 and the checkpoint kinases (Chk1/2) initiate further signaling cascades to trigger
the activation of DNA damage cell cycle checkpoints leading to cell
cycle arrest and the possible initiation of apoptosis [133,134].
There is some evidence that Ku is involved in the modulation of
ATM activity following DNA damage. Ku appears to prevent ATMdependent ATR activation, perhaps acting as a signaling block to
the HR pathway [135]. Furthermore, Ku80 null cells were shown to
have increased S phase inhibition after DNA damage due to
increased ATM activity [136]. We identified a specific residue in
Ku70’s vWA domain, serine 155, that when mutated to alanine
results in decreased DNA damage signaling and apoptotic
activation after IR, despite having no impact on Ku’s ability to
21
function in C-NHEJ [21]. These results suggest that in addition to its
essential role in DNA repair, modification of Ku also signals to the
ATM dependent DDR cascade to regulate cell fate after DNA
damage. This is not entirely surprising as several other DNA repair
factors (ex. Mre11), have functions in both DNA repair as well as
the DNA damage signaling cascade [137]. It is possible that Ku also
has a dual role in the DNA damage response, first promoting the
repair by NHEJ, as well as relaying signals as to the completion, or
lack thereof, of DNA repair.
In addition to its impact on upstream DDR signaling, a number
of studies have implicated Ku70 as a direct inhibitor of apoptosis,
through the binding and inhibition of the pro-apoptotic factor Bax
[138,139]. This interaction is proposed to be modulated by various
mechanisms, including protein–protein interactions and posttranslational modification of Ku. Some examples of protein
interactions include the binding of Ku70 to NBS1 and a cleavage
product of cyclin E, both of which were suggested to inhibit the
binding of Ku70 to Bax and promote apoptosis [140,141]. The
acetylation of Ku70 is also suggested to negatively regulate the
Ku70-Bax interaction. Ku70 is acetylated in the C-terminal linker
domain on eight lysine residues between K539 and K556 and
alanine substitution of these residues decreased Bax-dependent
apoptotic activation [30]. This acetylation is mediated by CREBbinding protein (CBP) and P300/CBP-associated factor (PCAF)
while deacetylation is controlled by histone deacetylase (HDAC)
and sirtuin (SIRT) deacetylase enzymes [30,142,143]. There is some
evidence that Ku may have deubiquitination activity, as it has been
proposed to promote the deubuquitination of Bax, and another
anti-apoptotic regulator of Bax, Mcl-1 [138,144]. Overall, there are
several unanswered questions regarding the interaction with Ku70
and Bax. Many studies have suggested that this interaction occurs
in the absence of Ku80, despite the fact that there is little evidence
that the Ku subunits exist as monomers. Furthermore, Ku is
predominantly a nuclear protein but Bax is largely cytoplasmic in
its inactive form and then translocates to the mitochondria during
apoptotic activation [145]. The regulatory mechanisms that would
allow a subset of Ku70 monomers to remain in the cytoplasm
bound to Bax remain to be elucidated.
5. Ku at telomeres
Telomeres are the linear ends of chromosomes and therefore
have the potential to be recognized as DSBs and processed by DSB
repair pathways. Specific protein complexes, such as the shelterin
P
ATM
ATM
P
P
P
Rif1
MDC1
P
H2AX
P
P
H2AX
DNA-PK
CS
P
BRCA1
53BP1
P
P
H2AX
H2AX
P
H2AX
Rad50
Ku80 Ku70
P
p53
P
Cell cycle arrest
Chk2
Apoptosis
Fig. 3. Overview of DNA double-strand break (DSB) recognition and DNA Damage Response (DDR). After the introduction of a double-strand break (DSB), the broken DNA end
is rapidly recognized by the Ku heterodimer and the MRN complex (Mre11/Rad50/NBS1). Next is the recruitment of the PIKK family members DNA-PKcs, bound to Ku, and
ATM, which both phosphorylate the histone variant H2AX on its C-terminal residue Ser139 (to yield g-H2AX). The MRN complex promotes the activation of ATM molecules,
which in turn further amplify H2AX phosphorylation and spread the g-H2AX signal far from the initial damage site. Several other factors are phosphorylated by ATM and
recruited to the damaged site in large numbers, including MDC1, 53BP1, Rif1 and BRCA1. ATM also initiates a signaling cascade through the phosphorylation of p53 and Chk1/
2 that results in broad transcriptional changes to induce cell cycle arrest or activate apoptosis if the damage is unable to be repaired [131].
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
22
complex in mammals and the Rap1 and the CST (Cdc13–Stn1–
Ten1) complexes in yeast have evolved to bind and form protective
caps on the DNA ends to prevent access by the DNA repair
complexes (Fig. 4) [146,147]. A role for Ku in the maintenance of
telomeres in yeast has been long established. Foundational to this
was the observation that deletion of either Ku subunit in S.
cerevisae resulted in shortened telomeres and long G-tails
compared to wildtype [148–150]. The requirement of Ku for
proper telomere structure and maintenance appears to be separate
from C-NHEJ, as DNA Ligase IV deficient strains, the ligase essential
for completion of C-NHEJ, do not show any telomere defects [151].
A
Sir2/3/4
Rad50
TLC1 RNA
Mre11
Xrs2
Telomerase
Rap1
(b)
Ku70 Ku80
(a)
B
TIN1
Mre11
TPP1
Rad50
POT1
NBS1
TRF1 TRF2
Ku70 Ku80
(a)
Ku70 Ku80
(b)
Ku70
Fig. 4. General structure of telomeres. In most eukaryotes telomeric DNA consists of
extended tracts of G-rich repeat arrays. The G-rich strand forms a short 30 singlestrand protrusion called the G-overhang, which mediates the formation of a
complex secondary structure called a t-loop. The t-loop structure allows the folding
of the end of the chromosome to mask and protect the DNA end from degradation.
(A) Structure of the yeast telomere. The MRX (Mre11/Rad50/Xrs2) and Sir/Rap1
complexes directly bind the telomeric DNA [252]. Ku could potentially (a) bind the
DNA end via its DNA binding domain or (b) bind via a protein–protein interaction
with telomeric complexes (such as Sir/Rap1). Ku has a role in recruiting telomerase,
mediated by an interaction with the TLC1 RNA and Ku80. Ku interaction with the
telomere may be indirect, requiring the heterotetramerization of two Ku molecules.
(B) Structure of the mammalian telomere. The Shelterin (TRF1/TRF2/Tin1/Pot1/
TPP1) and MRN (Mre11/Rad50/Nbs1) complexes coat the telomeric DNA to protect
from degradation [253]. Again there is controversy whether (a) Ku directly binds
the telomeric DNA or (b) is retained through a protein interaction with components
of the Shelterin complex (such as TRF1/TRF2).
Analysis of telomeres in Ku-deficient mice has produced conflicting results regarding telomere length, with reports of telomere
shortening as well as lengthening [152–154]. Despite this, the
same mice exhibit increased telomere end-to-end fusions and
chromosomal aberrations [153,154]. Similarly, human cells with
deleted Ku80 show telomere loss and abnormal telomere
structure, overall indicating a clear role for Ku in proper telomere
structure maintenance in mammals [155].
Similar to its role in protecting the DNA ends of DSBs in NHEJ,
Ku protects telomere ends from recombination and degradation
events [148–150,152–154]. Paradoxically, although Ku promotes
the fusion of dysfunctional, or uncapped, telomere ends, it has an
inhibitory effect on the HR and A-NHEJ pathways and on the
recombination of normal telomeres. Mouse embryonic fibroblasts
(MEFs) deficient in Ku70 have normal telomere structure, however
they exhibit increased sister telomere exchanges mediated by HR
and chromosome fusions by the A-NHEJ pathway [156,157].
Additionally, Ku deficient S. cerevisiae display increased RAD52
dependent recombination of the subtelomeric DNA elements,
consistent with its general HR inhibitory function [158,159]. While
wildtype cells only exhibit long G-tails during late S phase, yeast
Ku deficient strains possess long G-tails throughout the cell cycle.
This defect is almost completely suppressed by the deletion of the
50 to 30 exonuclease Exo1, suggesting that Ku acts to inhibit
inappropriate nucleolytic degradation of the C strand by nucleases
[160].
Ku also positively regulates telomere length through telomere
addition by aiding in the recruitment of telomerase. In yeast, Ku
has been shown to interact with telomerase component 1 (TLC1),
the telomerase RNA subunit, via binding of TLC1’s 48nt stem-loop
region [161,162]. Deletion of the binding interface from either the
Ku80 vWA domain or the TLC1 stem-loop results in decreased
telomere length and decreased de novo telomere addition, perhaps
due to the decreased levels of TLC1 RNA or altered localization of
the telomerase holoenzyme [161,163–166]. There is no sequence
conservation between TLC1 and the human telomerase RNA
counterpart, hRT, however there is evidence that the interaction
between the RNA component of human telomerase (hRT) and Ku in
humans is conserved [167]. Another study detected an interaction
with Ku and telomerase in human cells, however this interaction
was reported to occur through the catalytic reverse transcriptase
protein subunit hTERT [168].
The phenomenon of the telomere position silencing effect (TPE),
observed in many eukaryotic species, is the process by which
organisms transcriptionally silence genes located near the
telomere (reviewed in [169,170]). In S. cerevisiae, Ku, along with
the silent information regulator (Sir) complex, are essential for TPE,
as deletion of either Ku subunit results in complete loss of
telomeric silencing [171–174]. As a consequence of this role, Ku is
also involved in the nuclear organization of telomeres. Telomeres
and TPE proteins in S. cerevisiae are found distinctly clustered in
foci around the nuclear periphery, and the loss of Ku function often
results in the random distribution of telomeres throughout the
nucleus [174–176]. Ku participates in these processes as part of
multi-protein complexes with various proteins (such as the Sir
proteins, Rap1 and Mps3) and Ku is particularly essential for their
formation in the G1 phase of the cell cycle [176–178].
It is still currently unknown how Ku is associated with the
telomere in vivo, whether it is through direct DNA binding of the
end, or is mediated through a protein–protein interaction with
another telomere bound factor. Ku has the ability to directly bind
telomere DNA in vitro, in both mammals and yeast [179,180].
Furthermore, mutations in yeast Ku’s DNA binding domain reduces
association with telomeric chromatin and renders it unable to
protect the telomere end, suggesting that this region is key for
telomere binding [181]. Despite this evidence, there is still some
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
question whether it truly slides onto the telomere end via its DNA
binding ring. Telomeres in many species form higher order loop
structures designed to conceal the end from attack by Ku and the
DSB repair pathways, so further research needs to be done to
understand how and when Ku would be allowed access to the
telomere end.
Another possibility is that Ku is retained at the telomere
through protein–protein interaction with other telomere bound
factor(s). Ku has been shown to interact with TRF1, TRF2 and Rap1
[20,182–186]. These components of the shelterin complex bind
DNA directly and then mediate the recruitment of other shelterin
factors, such as TERF1-interacting nuclear factor 2 (TIN2) to the TRF
proteins in humans and Sir3/4 to Rap1 in S. cerevisiae, and therefore
could also be involved to recruit Ku at the telomeres [187,188]. A
longstanding model for telomerase recruitment by Ku in yeast,
suggested that Ku bound to the telomere end through its DNA
binding ring and then bound the TLC1 RNA via the yeast Ku80 vWA
domain [161]. However there is evidence that Ku requires the DNA
binding domain for both DNA and RNA binding and cannot bind
both simultaneously [189]. These observations suggest that Ku
does not recruit telomerase to the telomere by binding the
telomere directly, and instead favors a model where Ku must bind
another telomere bound factor to be retained. A combination of
DNA binding and protein interaction is also possible and has been
reported for the shelterin component protection of telomeres 1
(POT1), which was shown to bind the G-strand of telomeres, as
well as form multiple protein interactions [190,191]. Given that Ku
has been shown to multimerize, it is possible that one Ku molecule
interacts with the telomere end then multimerizes with another
Ku molecule that is bound to telomerase. Although it appears that
there are similarities in Ku telomere binding between yeast and
mammalian systems, these organisms have different telomere
structure and protein complexes present, so it is possible that Ku
has different mechanisms for telomere binding and protection.
Advances in high-resolution microscopy techniques have allowed
for direct visualization of telomere structures, such as mammalian
t-loop formation by TRF2, and could help in understanding Ku’s
role at the telomere [192].
6. Ku in disease
6.1. Immune system disorders
The vertebrate immune system utilizes Ku and C-NHEJ to repair
physiological DSBs generated to create immune system genetic
diversity. Recombination of the V, D, and J segments of
immunoglobulins in lymphoid cells and class switch recombination of the T-cell receptor genes in mature B cells allows for the
recognition of a wide variety antigens, which results in an adaptive
immune response [193]. Ku70 and Ku80 knockout mice display
many immune system abnormalities including B cell developmental arrest, T cell arrest, and failed lymphocyte differentiation
following g-irradiation [50,194–196]. Humans and mice suffering
from genetic defects in V(D)J recombination present with a severe
combined immunodeficiency (SCID) phenotype resulting in severe
infections and poor prognosis. There are several characterized
human disorders with various degree of SCID phenotype that have
been linked to mutations in NHEJ factors, such as DNA Ligase IV,
Artemis, and XLF, as well as the other DSB repair pathway and DDR
factors [197–200]. Interestingly, only a few individuals have been
reported to have genetic defects in the DNA-PK complex, however,
all presented mutations occurred in the DNA-PKCS subunit, and
thus far, no disorder linked to Ku mutations has been identified
[201,202]. It is currently unknown whether mutations in Ku are
better tolerated in humans, or in the contrary, whether deleterious
Ku mutations are lethal in humans.
23
6.2. Aging
Aging is the progressive degeneration that occurs at the cellular
and organismal level. One of the observed phenotypes of Ku
deficient mice is increased aging and senescence [196,203–205].
Mice deficient in one Ku subunit have one-third the lifespan of
their wildtype counterparts, partially in due to an early onset of the
phenotypic symptoms of aging [206]. There is evidence that CNHEJ declines during the aging process, with lower efficiency
observed in aging rodents, and in Alzheimer’s patients, so the loss
of NHEJ in Ku deficient mice could be contributing to the aging
process [207–209]. Furthermore, as previously described, Ku
deficient mice and cells exhibit shortened telomeres as well as
other telomere abnormalities. Shortened telomeres have been
linked to increased aging and senescence, due to increased
genomic instability leading to cell cycle arrest, so this could also
be contributing to the aging phenotype observed in Ku-deficient
mice [210,211].
6.3. Cancer
A common characteristic amongst human tumors is genomic
instability [212]. Cells defective in DSB repair are predisposed to
chromosome translocations and gene amplifications, potentially
leading to the activation of oncogenes and tumorigenesis. Through
its role in C-NHEJ, Ku functions as a cancer caretaker gene,
promoting genomic integrity and preventing tumorigenesis. Ku
knockout mice show increased chromosomal breakage, translocations and aneuploidy [196,203,205,206]. These mice display
slightly increased cancer incidence as compared to controls,
however
this
is
mostly
restricted
to
lymphomas
[196,203,205,206]. Together with a p53 deficiency that impairs
control of proliferation, Ku80 deficient mice show early onset
tumor formation and further increased incidence of T and B cell
lymphomas [51]. This suggests that Ku mutations can be oncogenic
when accompanied by a mutation in a tumor suppressor gene.
The expression of Ku has been frequently found deregulated in
tumor samples and its expression level has been proposed as a
marker of predicting patient response to radiation therapy and
survival [213–218]. The exploitation of genetic defects in DNA
repair and DDR factors have long been employed in cancer
treatment [219]. As previously mentioned, Ku and DNA-PKCS
deficient cell lines are sensitive to radiation and chemotherapeutic
drugs, and therefore would make them potential druggable targets
in cancer treatment. There are a few examples of strategies to
target Ku specifically, including a subunit dimerization interference using a peptide, downregulation by RNAi and small molecule
inhibition [220–223]. The therapeutic potential of targeting the
Ku70/Bax interaction to activate apoptosis in cancer has been
explored through creation of Bax inhibiting peptides derived from
the Ku70 sequence and promoting Ku70 acetylation in vivo
through deacetylase inhibitors [224–226]. A more commonly
employed strategy to target the DNA-PK complex however, is to
modulate the kinase activity of DNA-PKCS with small molecular
inhibitors, as DNA-PKCS expression is frequently found upregulated in tumors [227].
7. Conclusion
Ku is a versatile, multifunctional protein that has integral roles
in DNA repair, telomere maintenance and DNA damage signaling.
The central importance of Ku to cellular homeostasis is best
demonstrated by the knockout of Ku in mouse models, which
results in a complex phenotype, highlighted by extreme immunodeficiency, susceptibility to cancer and accelerated aging. Interestingly, despite the identification of human diseases stemming
24
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
from mutation in the majority of other important DSB and DDR
factors, no Ku mutations have been observed in humans. This leads
to the question as to whether Ku function is essential for human
cell viability. Additionally, an unexplained phenotype observed in
Ku deficient mice is that they are proportional dwarfs to their
wildtype counterparts [196,205]. The cause of this phenomenon is
currently unknown, but there are several studies identifying Ku as
an interactor with proteins involved in gene transcription, DNA
replication and cell cycle regulation (Table 1). These observations
are currently not well understood but could point to even further
roles for Ku in cellular maintenance. Overall, it is of critical
importance to understand the differing roles of Ku at the cellular
level and their implications in human disease.
[21]
[22]
[23]
[24]
[25]
Acknowledgement
[26]
This work was supported by a Natural Sciences and Engineering
Research Council of Canada (NSERC) Discovery Grant to C.S.P.
[27]
References
[28]
[1] T. Mimori, M. Akizuki, H. Yamagata, S. Inada, S. Yoshida, M. Homma, Characterization of a high molecular weight acidic nuclear protein recognized by autoantibodies in sera from patients with polymyositis-scleroderma overlap, J. Clin.
Invest. 68 (1981) 611–620.
[2] T. Mimori, Clinical significance of anti-Ku autoantibodies–a serologic marker of
overlap syndrome? Intern. Med. 41 (2002) 1096–1098.
[3] H.M. Cooley, B.J. Melny, R. Gleeson, T. Greco, T.W. Kay, Clinical and serological
associations of anti-Ku antibody, J. Rheumatol. 26 (1999) 563–567.
[4] Y. Takeda, W.S. Dynan, Autoantibodies against DNA double-strand break repair
proteins, Front. Biosci.: J. Virtual Lib. 6 (2001) D1412–D1422.
[5] T. Mimori, J.A. Hardin, J.A. Steitz, Characterization of the DNA-binding protein
antigen Ku recognized by autoantibodies from patients with rheumatic disorders, J. Biol. Chem. 261 (1986) 2274–2278.
[6] E. de Vries, W. van Driel, W.G. Bergsma, A.C. Arnberg, P.C. van der Vliet, HeLa
nuclear protein recognizing DNA termini and translocating on DNA forming a
regular DNA-multimeric protein complex, J. Mol. Biol. 208 (1989) 65–78.
[7] A.J. Griffith, P.R. Blier, T. Mimori, J.A. Hardin, Ku polypeptides synthesized in vitro
assemble into complexes which recognize ends of double-stranded DNA, J. Biol.
Chem. 267 (1992) 331–338.
[8] R.C. Getts, T.D. Stamato, Absence of a Ku-like DNA end binding activity in the xrs
double-strand DNA repair-deficient mutant, J. Biol. Chem. 269 (1994) 15981–
15984.
[9] G.W. Verhaegh, W. Jongmans, B. Morolli, N.G. Jaspers, G.P. van der Schans, P.H.
Lohman, M.Z. Zdzienicka, A novel type of X-ray-sensitive Chinese hamster cell
mutant with radioresistant DNA synthesis and hampered DNA double-strand
break repair, Mutat. Res. 337 (1995) 119–129.
[10] G.E. Taccioli, A.G. Amatucci, H.J. Beamish, D. Gell, X.H. Xiang, M.I. Torres Arzayus,
A. Priestley, S.P. Jackson, A. Marshak Rothstein, P.A. Jeggo, V.L. Herrera, Targeted
disruption of the catalytic subunit of the DNA-PK gene in mice confers severe
combined immunodeficiency and radiosensitivity, Immunity 9 (1998) 355–366.
, http://dx.doi.org/10.1016/S1074-7613(00)80618-4.
[11] W.K. Rathmell, G. Chu, Involvement of the Ku autoantigen in the cellular
response to DNA double-strand breaks, Proc. Natl. Acad. Sci. U. S. A. 91
(1994) 7623–7627.
[12] W.K. Rathmell, G. Chu, A DNA end-binding factor involved in double-strand
break repair and V(D)J recombination, Mol. Cell. Biol. 14 (1994) 4741–4748.
[13] V. Smider, W.K. Rathmell, M.R. Lieber, G. Chu, Restoration of X-ray resistance and
V(D)J recombination in mutant cells by Ku cDNA, Science 266 (1994) 288–291.
[14] Y. Gu, S. Jin, Y. Gao, D.T. Weaver, F.W. Alt, Ku70-deficient embryonic stem cells
have increased ionizing radiosensitivity, defective DNA end-binding activity,
and inability to support V(D)J recombination, Proc. Natl. Acad. Sci. U. S. A. 94
(1997) 8076–8081.
[15] J.R. Walker, R.A. Corpina, J. Goldberg, Structure of the Ku heterodimer bound to
DNA and its implications for double-strand break repair, Nature 412 (2001) 607–
614. , http://dx.doi.org/10.1038/35088000.
[16] S. Paillard, F. Strauss, Analysis of the mechanism of interaction of simian Ku
protein with DNA, Nucleic Acids Res. 19 (1991) 5619–5624. , http://dx.doi.org/
10.1093/nar/19.20.5619.
[17] M. Ono, P.W. Tucker, J.D. Capra, Production and characterization of recombinant
human Ku antigen, Nucleic Acids Res. 22 (1994) 3918–3924.
[18] C.A. Whittaker, R.O. Hynes, Distribution and evolution of von Willebrand/integrin A domains: widely dispersed domains with roles in cell adhesion and
elsewhere, Mol. Biol. Cell 13 (2002) 3369–3387. , http://dx.doi.org/10.1091/
mbc.E02-05-0259.
[19] A. Ribes-Zamora, I. Mihalek, O. Lichtarge, A.A. Bertuch, Distinct faces of the Ku
heterodimer mediate DNA repair and telomeric functions, Nat. Struct. Mol. Biol.
14 (2007) 301–307. , http://dx.doi.org/10.1038/nsmb1214.
[20] A. Ribes-Zamora, S.M. Indiviglio, I. Mihalek, C.L. Williams, A.A. Bertuch, TRF2
interaction with Ku heterotetramerization interface gives insight into c-NHEJ
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
[37]
[38]
[39]
[40]
[41]
[42]
[43]
[44]
[45]
prevention at human telomeres, Cell Rep. (2013), http://dx.doi.org/10.1016/
j.celrep.2013.08.040.
V.L. Fell, C. Schild-Poulter, Ku regulates signaling to DNA damage response
pathways through the Ku70 von Willebrand A domain, Mol. Cell. Biol. 32
(2012) 76–87. , http://dx.doi.org/10.1128/MCB. 05661-11.
N. Strande, S.A. Roberts, S. Oh, E.A. Hendrickson, D.A. Ramsden, Specificity of the
dRP/AP lyase of Ku promotes nonhomologous end joining (NHEJ) fidelity at
damaged ends, J. Biol. Chem. 287 (2012) 13686–13693. , http://dx.doi.org/
10.1074/jbc.M111.329730.
S.A. Roberts, N. Strande, M.D. Burkhalter, C. Strom, J.M. Havener, P. Hasty, D.A.
Ramsden, Ku is a 50 -dRP/AP lyase that excises nucleotide damage near broken
ends, Nature 464 (2010) 1214–1217. , http://dx.doi.org/10.1038/nature08926.
P. Shirodkar, A.L. Fenton, L. Meng, C.A. Koch, Identification and functional
characterization of a Ku-binding motif in aprataxin polynucleotide kinase/
phosphatase-like factor (APLF), J. Biol. Chem. 288 (2013) 19604–19613. ,
http://dx.doi.org/10.1074/jbc.M112.440388.
D. Gell, S.P. Jackson, Mapping of protein–protein interactions within the
DNA-dependent protein kinase complex, Nucleic Acids Res. 27 (1999) 3494–
3502.
B.K. Singleton, M.I. Torres-Arzayus, S.T. Rottinghaus, G.E. Taccioli, P.A. Jeggo, The
C terminus of Ku80 activates the DNA-dependent protein kinase catalytic
subunit, Mol. Cell. Biol. 19 (1999) 3267–3277.
L. Aravind, E.V. Koonin, SAP – a putative DNA-binding motif involved in chromosomal organization, Trends Biochem. Sci. 25 (2000) 112–114. , http://
dx.doi.org/10.1016/S0968-0004(99)01537-6.
S. Hu, J.M. Pluth, F.A. Cucinotta, Putative binding modes of Ku70-SAP domain
with double strand DNA: a molecular modeling study, J. Mol. Model. 18 (2012)
2163–2174. , http://dx.doi.org/10.1007/s00894-011-1234-x.
J. Wang, X. Dong, W.H. Reeves, A model for Ku heterodimer assembly and
interaction with DNA. Implications for the function of Ku antigen, J. Biol. Chem.
273 (1998) 31068–31074.
H.Y. Cohen, S. Lavu, K.J. Bitterman, B. Hekking, T.A. Imahiyerobo, C. Miller, R. Frye,
H. Ploegh, B.M. Kessler, D.A. Sinclair, Acetylation of the C terminus of Ku70 by
CBP and PCAF controls Bax-mediated apoptosis, Mol. Cell 13 (2004) 627–638. ,
http://dx.doi.org/10.1016/S1097-2765(04)00094-2.
K.B. Kim, D.W. Kim, J.W. Park, Y.J. Jeon, D. Kim, S. Rhee, J.I. Chae, S.B. Seo,
Inhibition of Ku70 acetylation by INHAT subunit SET/TAF-Ibeta regulates Ku70mediated DNA damage response, Cell. Mol. Life Sci.: CMLS (2013), http://
dx.doi.org/10.1007/s00018-013-1525-8.
L.E. Hang, C.R. Lopez, X. Liu, J.M. Williams, I. Chung, L. Wei, A.A. Bertuch, X. Zhao,
Regulation of Ku-DNA association by Yku70 C-terminal tail and SUMO modification, J. Biol. Chem. 289 (2014) 10308–10317. , http://dx.doi.org/10.1074/
jbc.M113.526178.
C. Schild-Poulter, L. Pope, W. Giffin, J.C. Kochan, J.K. Ngsee, M. Traykova-Andonova, R.J. Hache, The binding of Ku antigen to homeodomain proteins promotes
their phosphorylation by DNA-dependent protein kinase, J. Biol. Chem. 276
(2001) 16848–16856. , http://dx.doi.org/10.1074/jbc.M100768200.
J. Bertinato, C. Schild-Poulter, R.J. Hache, Nuclear localization of Ku antigen is
promoted independently by basic motifs in the Ku70 and Ku80 subunits, J. Cell
Sci. 114 (2001) 89–99.
B.K. Singleton, A. Priestley, H. Steingrimsdottir, D. Gell, T. Blunt, S.P. Jackson, A.R.
Lehmann, P.A. Jeggo, Molecular and biochemical characterization of xrs mutants
defective in Ku80, Mol. Cell. Biol. 17 (1997) 1264–1273.
A. Errami, V. Smider, W.K. Rathmell, D.M. He, E.A. Hendrickson, M.Z. Zdzienicka,
G. Chu, Ku86 defines the genetic defect and restores X-ray resistance and V(D)J
recombination to complementation group 5 hamster cell mutants, Mol. Cell.
Biol. 16 (1996) 1519–1526.
J.A. Downs, S.P. Jackson, A means to a DNA end: the many roles of Ku, Nat. Rev.
Mol. Cell Biol. 5 (2004) 367–378. , http://dx.doi.org/10.1038/nrm1367.
A.J. Doherty, S.P. Jackson, G.R. Weller, Identification of bacterial homologues of
the Ku DNA repair proteins, FEBS Lett. 500 (2001) 186–188. , http://dx.doi.org/
10.1016/S0014-5793(01)02589-3.
L. Aravind, E.V. Koonin, Prokaryotic homologs of the eukaryotic DNA-endbinding protein Ku, novel domains in the Ku protein and prediction of a
prokaryotic double-strand break repair system, Genome Res. 11 (2001)
1365–1374. , http://dx.doi.org/10.1101/gr.181001.
R. Bowater, A.J. Doherty, Making ends meet: repairing breaks in bacterial DNA by
non-homologous end-joining, PLoS Genet. 2 (2006) e8, http://dx.doi.org/
10.1371/journal.pgen.0020008.
G.R. Weller, B. Kysela, R. Roy, L.M. Tonkin, E. Scanlan, M. Della, S.K. Devine, J.P.
Day, A. Wilkinson, F. d’Adda di Fagagna, K.M. Devine, R.P. Bowater, P.A. Jeggo, S.P.
Jackson, A.J. Doherty, Identification of a DNA nonhomologous end-joining complex in bacteria, Science 297 (2002) 1686–1689. , http://dx.doi.org/10.1126/
science.1074584.
P. Frit, N. Barboule, Y. Yuan, D. Gomez, P. Calsou, Alternative end-joining pathway(s): bricolage at DNA breaks, DNA Repair (Amst.) 17 (2014) 81–97. , http://
dx.doi.org/10.1016/j.dnarep.2014.02.007.
L. Deriano, D.B. Roth, Modernizing the nonhomologous end-joining repertoire:
alternative and classical NHEJ share the stage, Annu. Rev. Genet. 47 (2013) 433–
455. , http://dx.doi.org/10.1146/annurev-genet-110711-155540.
L. Krejci, V. Altmannova, M. Spirek, X. Zhao, Homologous recombination and its
regulation, Nucleic Acids Res. 40 (2012) 5795–5818. , http://dx.doi.org/10.1093/
nar/gks270.
G.J. Williams, M. Hammel, S.K. Radhakrishnan, D. Ramsden, S.P. Lees-Miller, J.A.
Tainer, Structural insights into NHEJ: building up an integrated picture of the
dynamic DSB repair super complex, one component and interaction at a time,
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
[46]
[47]
[48]
[49]
[50]
[51]
[52]
[53]
[54]
[55]
[56]
[57]
[58]
[59]
[60]
[61]
[62]
[63]
[64]
[65]
[66]
[67]
[68]
DNA Repair (Amst.) 17 (2014) 110–120. , http://dx.doi.org/10.1016/j.dnarep.
2014.02.009.
T. Ochi, Q. Wu, T.L. Blundell, The spatial organization of non-homologous end
joining: from bridging to end joining, DNA Repair (Amst.) 17 (2014) 98–109. ,
http://dx.doi.org/10.1016/j.dnarep.2014.02.010.
G.J. Grundy, H.A. Moulding, K.W. Caldecott, S.L. Rulten, One ring to bring them all
– the role of Ku in mammalian non-homologous end joining, DNA Repair (Amst.)
17 (2014) 30–38. , http://dx.doi.org/10.1016/j.dnarep.2014.02.019.
S.J. Boulton, S.P. Jackson, Saccharomyces cerevisiae Ku70 potentiates illegitimate
DNA double-strand break repair and serves as a barrier to error-prone DNA
repair pathways, EMBO J. 15 (1996) 5093–5103.
S. Chen, K.V. Inamdar, P. Pfeiffer, E. Feldmann, M.F. Hannah, Y. Yu, J.W. Lee, T.
Zhou, S.P. Lees-Miller, L.F. Povirk, Accurate in vitro end joining of a DNA double
strand break with partially cohesive 30 -overhangs and 30 -phosphoglycolate
termini: effect of Ku on repair fidelity, J. Biol. Chem. 276 (2001) 24323–
24330. , http://dx.doi.org/10.1074/jbc.M010544200.
A. Nussenzweig, K. Sokol, P. Burgman, L. Li, G.C. Li, Hypersensitivity of Ku80deficient cell lines and mice to DNA damage: the effects of ionizing radiation on
growth, survival, and development, Proc. Natl. Acad. Sci. U. S. A. 94 (1997)
13588–13593.
M.J. Difilippantonio, J. Zhu, H.T. Chen, E. Meffre, M.C. Nussenzweig, E.E. Max, T.
Ried, A. Nussenzweig, DNA repair protein Ku80 suppresses chromosomal aberrations and malignant transformation, Nature 404 (2000) 510–514. , http://
dx.doi.org/10.1038/35006670.
D.O. Ferguson, J.M. Sekiguchi, S. Chang, K.M. Frank, Y. Gao, R.A. DePinho, F.W. Alt,
The nonhomologous end-joining pathway of DNA repair is required for genomic
stability and the suppression of translocations, Proc. Natl. Acad. Sci. U. S. A. 97
(2000) 6630–6633. , http://dx.doi.org/10.1073/pnas.110152897.
P.R. Blier, A.J. Griffith, J. Craft, J.A. Hardin, Binding of Ku protein to DNA.
Measurement of affinity for ends and demonstration of binding to nicks, J. Biol.
Chem. 268 (1993) 7594–7601.
P.O. Mari, B.I. Florea, S.P. Persengiev, N.S. Verkaik, H.T. Bruggenwirth, M. Modesti,
G. Giglia-Mari, K. Bezstarosti, J.A. Demmers, T.M. Luider, A.B. Houtsmuller, D.C.
van Gent, Dynamic assembly of end-joining complexes requires interaction
between Ku70/80 and XRCC4, Proc. Natl. Acad. Sci. U. S. A. 103 (2006)
18597–18602. , http://dx.doi.org/10.1073/pnas.0609061103.
E.P. Mimitou, L.S. Symington, Ku prevents Exo1 and Sgs1-dependent resection of
DNA ends in the absence of a functional MRX complex or Sae2, EMBO J. 29 (2010)
3358–3369. , http://dx.doi.org/10.1038/emboj.2010.193.
S.A. Roberts, D.A. Ramsden, Loading of the nonhomologous end joining factor,
Ku, on protein-occluded DNA ends, J. Biol. Chem. 282 (2007) 10605–10613. ,
http://dx.doi.org/10.1074/jbc.M611125200.
S. Britton, J. Coates, S.P. Jackson, A new method for high-resolution imaging of Ku
foci to decipher mechanisms of DNA double-strand break repair, J. Cell Biol. 202
(2013) 579–595. , http://dx.doi.org/10.1083/jcb.201303073.
C.A. Lovejoy, D. Cortez, Common mechanisms of PIKK regulation, DNA Repair
(Amst.) 8 (2009) 1004–1008. , http://dx.doi.org/10.1016/j.dnarep.2009.04.006.
A. Rivera-Calzada, L. Spagnolo, L.H. Pearl, O. Llorca, Structural model of fulllength human Ku70-Ku80 heterodimer and its recognition of DNA and DNAPKcs, EMBO Rep. 8 (2007) 56–62. , http://dx.doi.org/10.1038/sj.embor.7400847.
M. Hammel, Y. Yu, B.L. Mahaney, B. Cai, R. Ye, B.M. Phipps, R.P. Rambo, G.L. Hura,
M. Pelikan, S. So, R.M. Abolfath, D.J. Chen, S.P. Lees-Miller, J.A. Tainer, Ku and
DNA-dependent protein kinase dynamic conformations and assembly regulate
DNA binding and the initial non-homologous end joining complex, J. Biol. Chem.
285 (2010) 1414–1423. , http://dx.doi.org/10.1074/jbc.M109.065615.
P. Douglas, G.P. Sapkota, N. Morrice, Y. Yu, A.A. Goodarzi, D. Merkle, K. Meek, D.R.
Alessi, S.P. Lees-Miller, Identification of in vitro and in vivo phosphorylation sites
in the catalytic subunit of the DNA-dependent protein kinase, Biochem. J. 368
(2002) 243–251. , http://dx.doi.org/10.1042/BJ20020973.
D.W. Chan, B.P. Chen, S. Prithivirajsingh, A. Kurimasa, M.D. Story, J. Qin, D.J. Chen,
Autophosphorylation of the DNA-dependent protein kinase catalytic subunit is
required for rejoining of DNA double-strand breaks, Genes Dev. 16 (2002) 2333–
2338. , http://dx.doi.org/10.1101/gad.1015202.
Y. Yu, W. Wang, Q. Ding, R. Ye, D. Chen, D. Merkle, D. Schriemer, K. Meek, S.P.
Lees-Miller, DNA-PK phosphorylation sites in XRCC4 are not required for survival after radiation or for V(D)J recombination, DNA Repair (Amst.) 2 (2003)
1239–1252.
B.P. Chen, D.W. Chan, J. Kobayashi, S. Burma, A. Asaithamby, K. Morotomi-Yano,
E. Botvinick, J. Qin, D.J. Chen, Cell cycle dependence of DNA-dependent protein
kinase phosphorylation in response to DNA double strand breaks, J. Biol. Chem.
280 (2005) 14709–14715. , http://dx.doi.org/10.1074/jbc.M408827200.
P. Douglas, X. Cui, W.D. Block, Y. Yu, S. Gupta, Q. Ding, R. Ye, N. Morrice, S.P. LeesMiller, K. Meek, The DNA-dependent protein kinase catalytic subunit is phosphorylated in vivo on threonine 3950, a highly conserved amino acid in the
protein kinase domain, Mol. Cell. Biol. 27 (2007) 1581–1591. , http://dx.doi.org/
10.1128/MCB. 01962-06.
P. Douglas, S. Gupta, N. Morrice, K. Meek, S.P. Lees-Miller, DNA-PK-dependent
phosphorylation of Ku70/80 is not required for non-homologous end joining,
DNA Repair (Amst.) 4 (2005) 1006–1018. , http://dx.doi.org/10.1016/j.dnarep.
2005.05.003.
Y. Yu, B.L. Mahaney, K. Yano, R. Ye, S. Fang, P. Douglas, D.J. Chen, S.P. Lees-Miller,
DNA-PK and ATM phosphorylation sites in XLF/Cernunnos are not required for
repair of DNA double strand breaks, DNA Repair (Amst.) 7 (2008) 1680–1692. ,
http://dx.doi.org/10.1016/j.dnarep.2008.06.015.
A.A. Goodarzi, Y. Yu, E. Riballo, P. Douglas, S.A. Walker, R. Ye, C. Harer, C.
Marchetti, N. Morrice, P.A. Jeggo, S.P. Lees-Miller, DNA-PK autophosphorylation
[69]
[70]
[71]
[72]
[73]
[74]
[75]
[76]
[77]
[78]
[79]
[80]
[81]
[82]
[83]
[84]
[85]
[86]
[87]
[88]
[89]
[90]
[91]
[92]
[93]
25
facilitates Artemis endonuclease activity, EMBO J. 25 (2006) 3880–3889. , http://
dx.doi.org/10.1038/sj.emboj.7601255.
Y.G. Wang, C. Nnakwe, W.S. Lane, M. Modesti, K.M. Frank, Phosphorylation and
regulation of DNA ligase IV stability by DNA-dependent protein kinase, J. Biol.
Chem. 279 (2004) 37282–37290. , http://dx.doi.org/10.1074/jbc.M401217200.
D.A. Ramsden, M. Gellert, Ku protein stimulates DNA end joining by mammalian
DNA ligases: a direct role for Ku in repair of DNA double-strand breaks, EMBO J.
17 (1998) 609–614. , http://dx.doi.org/10.1093/emboj/17.2.609.
R.B. Cary, S.R. Peterson, J. Wang, D.G. Bear, E.M. Bradbury, D.J. Chen, DNA looping
by Ku and the DNA-dependent protein kinase, Proc. Natl. Acad. Sci. U. S. A. 94
(1997) 4267–4272.
E. Soutoglou, J.F. Dorn, K. Sengupta, M. Jasin, A. Nussenzweig, T. Ried, G. Danuser,
T. Misteli, Positional stability of single double-strand breaks in mammalian cells,
Nat. Cell Biol. 9 (2007) 675–682. , http://dx.doi.org/10.1038/ncb1591.
B.L. Mahaney, K. Meek, S.P. Lees-Miller, Repair of ionizing radiation-induced
DNA double-strand breaks by non-homologous end-joining, Biochem. J. 417
(2009) 639–650. , http://dx.doi.org/10.1042/BJ20080413.
Z. Zeng, F. Cortes-Ledesma, S.F. El Khamisy, K.W. Caldecott, TDP2/TTRAP is the
major 50 -tyrosyl DNA phosphodiesterase activity in vertebrate cells and is
critical for cellular resistance to topoisomerase II-induced DNA damage, J. Biol.
Chem. 286 (2011) 403–409. , http://dx.doi.org/10.1074/jbc.M110.181016.
F. Gomez-Herreros, R. Romero-Granados, Z. Zeng, A. Alvarez-Quilon, C. Quintero,
L. Ju, L. Umans, L. Vermeire, D. Huylebroeck, K.W. Caldecott, F. Cortes-Ledesma,
TDP2-dependent non-homologous end-joining protects against topoisomerase
II-induced DNA breaks and genome instability in cells and in vivo, PLoS Genet. 9
(2013) e1003226, http://dx.doi.org/10.1371/journal.pgen.1003226.
S. Costantini, L. Woodbine, L. Andreoli, P.A. Jeggo, A. Vindigni, Interaction of the
Ku heterodimer with the DNA ligase IV/Xrcc4 complex and its regulation by
DNA-PK, DNA Repair (Amst.) 6 (2007) 712–722. , http://dx.doi.org/10.1016/
j.dnarep.2006.12.007.
H.L. Hsu, S.M. Yannone, D.J. Chen, Defining interactions between DNA-PK and
ligase IV/XRCC4, DNA Repair (Amst.) 1 (2002) 225–235. , http://dx.doi.org/
10.1016/S1568-7864(01)00018-0.
K. Yano, K. Morotomi-Yano, S.Y. Wang, N. Uematsu, K.J. Lee, A. Asaithamby, E.
Weterings, D.J. Chen, Ku recruits XLF to DNA double-strand breaks, EMBO Rep. 9
(2008) 91–96. , http://dx.doi.org/10.1038/sj.embor.7401137.
K. Yano, K. Morotomi-Yano, K.J. Lee, D.J. Chen, Functional significance of the
interaction with Ku in DNA double-strand break recognition of XLF, FEBS Lett.
585 (2011) 841–846. , http://dx.doi.org/10.1016/j.febslet.2011.02.020.
P. Reynolds, J.A. Anderson, J.V. Harper, M.A. Hill, S.W. Botchway, A.W. Parker, P.
O’Neill, The dynamics of Ku70/80 and DNA-PKcs at DSBs induced by ionizing
radiation is dependent on the complexity of damage, Nucleic Acids Res. 40
(2012) 10821–10831. , http://dx.doi.org/10.1093/nar/gks879.
K. Datta, P. Jaruga, M. Dizdaroglu, R.D. Neumann, T.A. Winters, Molecular
analysis of base damage clustering associated with a site-specific radiationinduced DNA double-strand break, Radiat. Res. 166 (2006) 767–781. , http://
dx.doi.org/10.1667/RR0628.1.
K. Datta, R.D. Neumann, T.A. Winters, Characterization of complex apurinic/
apyrimidinic-site clustering associated with an authentic site-specific radiationinduced DNA double-strand break, Proc. Natl. Acad. Sci. U. S. A. 102 (2005)
10569–10574. , http://dx.doi.org/10.1073/pnas.0503975102.
K. Datta, R.D. Neumann, T.A. Winters, Characterization of a complex 125Iinduced DNA double-strand break: implications for repair, Int. J. Radiat. Biol.
81 (2005) 13–21.
J.S. Kim, T.B. Krasieva, H. Kurumizaka, D.J. Chen, A.M. Taylor, K. Yokomori,
Independent and sequential recruitment of NHEJ and HR factors to DNA damage
sites in mammalian cells, J. Cell Biol. 170 (2005) 341–347. , http://dx.doi.org/
10.1083/jcb.200411083.
L. Postow, C. Ghenoiu, E.M. Woo, A.N. Krutchinsky, B.T. Chait, H. Funabiki, Ku80
removal from DNA through double strand break-induced ubiquitylation, J. Cell
Biol. 182 (2008) 467–479. , http://dx.doi.org/10.1083/jcb.200802146.
L. Feng, J. Chen, The E3 ligase RNF8 regulates KU80 removal and NHEJ repair, Nat.
Struct. Mol. Biol. 19 (2012) 201–206. , http://dx.doi.org/10.1038/nsmb.2211.
P. Langerak, E. Mejia-Ramirez, O. Limbo, P. Russell, Release of Ku and MRN from
DNA ends by Mre11 nuclease activity and Ctp1 is required for homologous
recombination repair of double-strand breaks, PLoS Genet. 7 (2011) e1002271,
http://dx.doi.org/10.1371/journal.pgen.1002271.
M.J. Neale, J. Pan, S. Keeney, Endonucleolytic processing of covalent proteinlinked DNA double-strand breaks, Nature 436 (2005) 1053–1057. , http://
dx.doi.org/10.1038/nature03872.
D. Wu, L.M. Topper, T.E. Wilson, Recruitment and dissociation of nonhomologous
end joining proteins at a DNA double-strand break in Saccharomyces cerevisiae,
Genetics 178 (2008) 1237–1249. , http://dx.doi.org/10.1534/genetics.107.
083535.
A.J. Pierce, P. Hu, M. Han, N. Ellis, M. Jasin, Ku DNA end-binding protein
modulates homologous repair of double-strand breaks in mammalian cells,
Genes Dev. 15 (2001) 3237–3242. , http://dx.doi.org/10.1101/gad.946401.
M. Frank-Vaillant, S. Marcand, Transient stability of DNA ends allows nonhomologous end joining to precede homologous recombination, Mol. Cell 10
(2002) 1189–1199. , http://dx.doi.org/10.1016/S1097-2765(02)00705-0.
C. Allen, A. Kurimasa, M.A. Brenneman, D.J. Chen, J.A. Nickoloff, DNA-dependent
protein kinase suppresses double-strand break-induced and spontaneous homologous recombination, Proc. Natl. Acad. Sci. U. S. A. 99 (2002) 3758–3763. ,
http://dx.doi.org/10.1073/pnas.052545899.
M. Clerici, D. Mantiero, I. Guerini, G. Lucchini, M.P. Longhese, The Yku70-Yku80
complex contributes to regulate double-strand break processing and checkpoint
26
[94]
[95]
[96]
[97]
[98]
[99]
[100]
[101]
[102]
[103]
[104]
[105]
[106]
[107]
[108]
[109]
[110]
[111]
[112]
[113]
[114]
[115]
[116]
[117]
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
activation during the cell cycle, EMBO Rep. 9 (2008) 810–818. , http://dx.doi.org/
10.1038/embor.2008.121.
J.H. Barlow, M. Lisby, R. Rothstein, Differential regulation of the cellular response
to DNA double-strand breaks in G1, Mol. Cell 30 (2008) 73–85. , http://
dx.doi.org/10.1016/j.molcel.2008.01.016.
K. Tomita, A. Matsuura, T. Caspari, A.M. Carr, Y. Akamatsu, H. Iwasaki, K. Mizuno,
K. Ohta, M. Uritani, T. Ushimaru, K. Yoshinaga, M. Ueno, Competition between
the Rad50 complex and the Ku heterodimer reveals a role for Exo1 in processing
double-strand breaks but not telomeres, Mol. Cell. Biol. 23 (2003) 5186–5197. ,
http://dx.doi.org/10.1128/MCB.23.15.5186-5197.2003.
E.Y. Shim, W.H. Chung, M.L. Nicolette, Y. Zhang, M. Davis, Z. Zhu, T.T. Paull, G. Ira,
S.E. Lee, Saccharomyces cerevisiae Mre11/Rad50/Xrs2 and Ku proteins regulate
association of Exo1 and Dna2 with DNA breaks, EMBO J. 29 (2010) 3370–3380. ,
http://dx.doi.org/10.1038/emboj.2010.219.
A. Shibata, S. Conrad, J. Birraux, V. Geuting, O. Barton, A. Ismail, A. Kakarougkas, K.
Meek, G. Taucher-Scholz, M. Lobrich, P.A. Jeggo, Factors determining DNA
double-strand break repair pathway choice in G2 phase, EMBO J. 30 (2011)
1079–1092. , http://dx.doi.org/10.1038/emboj.2011.27.
H. Walden, A.J. Deans, The Fanconi anemia DNA repair pathway: structural and
functional insights into a complex disorder, Annu. Rev. Biophys. 43 (2014) 257–
278. , http://dx.doi.org/10.1146/annurev-biophys-051013-022737.
P. Pace, G. Mosedale, M.R. Hodskinson, I.V. Rosado, M. Sivasubramaniam, K.J.
Patel, Ku70 corrupts DNA repair in the absence of the Fanconi anemia pathway,
Science 329 (2010) 219–223. , http://dx.doi.org/10.1126/science.1192277.
A.M. Carr, S. Lambert, Replication stress-induced genome instability: the dark
side of replication maintenance by homologous recombination, J. Mol. Biol. 425
(2013) 4733–4744. , http://dx.doi.org/10.1016/j.jmb.2013.04.023.
S.S. Foster, A. Balestrini, J.H. Petrini, Functional interplay of the Mre11 nuclease
and Ku in the response to replication-associated DNA damage, Mol. Cell. Biol. 31
(2011) 4379–4389. , http://dx.doi.org/10.1128/MCB. 05854-11.
A. Balestrini, D. Ristic, I. Dionne, X.Z. Liu, C. Wyman, R.J. Wellinger, J.H. Petrini,
The Ku heterodimer and the metabolism of single-ended DNA double-strand
breaks, Cell Rep. 3 (2013) 2033–2045. , http://dx.doi.org/10.1016/j.celrep.
2013.05.026.
H. Wang, A.R. Perrault, Y. Takeda, W. Qin, H. Wang, G. Iliakis, Biochemical
evidence for Ku-independent backup pathways of NHEJ, Nucleic Acids Res.
31 (2003) 5377–5388.
M. Betermier, P. Bertrand, B.S. Lopez, Is non-homologous end-joining really an
inherently error-prone process? PLoS Genet. 10 (2014) e1004086, http://
dx.doi.org/10.1371/journal.pgen.1004086.
J. Guirouilh-Barbat, S. Huck, B.S. Lopez, S-phase progression stimulates both the
mutagenic KU-independent pathway and mutagenic processing of KU-dependent intermediates, for nonhomologous end joining, Oncogene 27 (2008) 1726–
1736. , http://dx.doi.org/10.1038/sj.onc.1210807.
K. Rothkamm, I. Kruger, L.H. Thompson, M. Lobrich, Pathways of DNA doublestrand break repair during the mammalian cell cycle, Mol. Cell. Biol. 23 (2003)
5706–5715.
M. Wang, W. Wu, B. Rosidi, L. Zhang, H. Wang, G. Iliakis, PARP-1 and Ku compete
for repair of DNA double strand breaks by distinct NHEJ pathways, Nucleic Acids
Res. 34 (2006) 6170–6182. , http://dx.doi.org/10.1093/nar/gkl840.
W.Y. Mansour, K. Borgmann, C. Petersen, E. Dikomey, J. Dahm-Daphi, The
absence of Ku but not defects in classical non-homologous end-joining is
required to trigger PARP1-dependent end-joining, DNA Repair (Amst.) 12
(2013) 1134–1142. , http://dx.doi.org/10.1016/j.dnarep.2013.10.005.
Q. Cheng, N. Barboule, P. Frit, D. Gomez, O. Bombarde, B. Couderc, G.S. Ren, B.
Salles, P. Calsou, Ku counteracts mobilization of PARP1 and MRN in chromatin
damaged with DNA double-strand breaks, Nucleic Acids Res. 39 (2011) 9605–
9619. , http://dx.doi.org/10.1093/nar/gkr656.
Y. Ariumi, M. Masutani, T.D. Copeland, T. Mimori, T. Sugimura, K. Shimotohno, K.
Ueda, M. Hatanaka, M. Noda, Suppression of the poly(ADP-ribose) polymerase
activity by DNA-dependent protein kinase in vitro, Oncogene 18 (1999) 4616–
4625. , http://dx.doi.org/10.1038/sj.onc.1202823.
F. Fattah, E.H. Lee, N. Weisensel, Y. Wang, N. Lichter, E.A. Hendrickson, Ku
regulates the non-homologous end joining pathway choice of DNA doublestrand break repair in human somatic cells, PLoS Genet. 6 (2010) e1000855,
http://dx.doi.org/10.1371/journal.pgen.1000855.
J.M. Enserink, R.D. Kolodner, An overview of Cdk1-controlled targets and processes, Cell Div. 5 (2010) 11, http://dx.doi.org/10.1186/1747-1028-5-11.
C. Trovesi, N. Manfrini, M. Falcettoni, M.P. Longhese, Regulation of the DNA
damage response by cyclin-dependent kinases, J. Mol. Biol. 425 (2013) 4756–
4766. , http://dx.doi.org/10.1016/j.jmb.2013.04.013.
M.H. Yun, K. Hiom, CtIP-BRCA1 modulates the choice of DNA double-strandbreak repair pathway throughout the cell cycle, Nature 459 (2009) 460–463. ,
http://dx.doi.org/10.1038/nature07955.
P. Huertas, S.P. Jackson, Human CtIP mediates cell cycle control of DNA end
resection and double strand break repair, J. Biol. Chem. 284 (2009) 9558–9565. ,
http://dx.doi.org/10.1074/jbc.M808906200.
X. Chen, H. Niu, W.H. Chung, Z. Zhu, A. Papusha, E.Y. Shim, S.E. Lee, P. Sung, G. Ira,
Cell cycle regulation of DNA double-strand break end resection by Cdk1-dependent Dna2 phosphorylation, Nat. Struct. Mol. Biol. 18 (2011) 1015–1019. ,
http://dx.doi.org/10.1038/nsmb.2105.
C. Muller-Tidow, P. Ji, S. Diederichs, J. Potratz, N. Baumer, G. Kohler, T. Cauvet, C.
Choudary, T. van der Meer, W.Y. Chan, C. Nieduszynski, W.H. Colledge, M.
Carrington, H.P. Koeffler, A. Restle, L. Wiesmuller, J. Sobczak-Thepot, W.E. Berdel,
H. Serve, The cyclin A1-CDK2 complex regulates DNA double-strand break
[118]
[119]
[120]
[121]
[122]
[123]
[124]
[125]
[126]
[127]
[128]
[129]
[130]
[131]
[132]
[133]
[134]
[135]
[136]
[137]
[138]
[139]
[140]
[141]
[142]
repair, Mol. Cell. Biol. 24 (2004) 8917–8928. , http://dx.doi.org/10.1128/
MCB.24.20.8917-8928.2004.
Y. Chi, M. Welcker, A.A. Hizli, J.J. Posakony, R. Aebersold, B.E. Clurman, Identification of CDK2 substrates in human cell lysates, Genome Biol. 9 (2008) R149,
http://dx.doi.org/10.1186/gb-2008-9-10-r149.
J.V. Olsen, M. Vermeulen, A. Santamaria, C. Kumar, M.L. Miller, L.J. Jensen, F.
Gnad, J. Cox, T.S. Jensen, E.A. Nigg, S. Brunak, M. Mann, Quantitative phosphoproteomics reveals widespread full phosphorylation site occupancy during
mitosis, Sci. Signal 3 (2010), http://dx.doi.org/10.1126/scisignal.2000475ra3.
Y. Zhang, E.Y. Shim, M. Davis, S.E. Lee, Regulation of repair choice: Cdk1
suppresses recruitment of end joining factors at DNA breaks, DNA Repair (Amst.)
8 (2009) 1235–1241. , http://dx.doi.org/10.1016/j.dnarep.2009.07.007.
A. Bothmer, D.F. Robbiani, N. Feldhahn, A. Gazumyan, A. Nussenzweig, M.C.
Nussenzweig, 53BP1 regulates DNA resection and the choice between classical
and alternative end joining during class switch recombination, J. Exp. Med. 207
(2010) 855–865. , http://dx.doi.org/10.1084/jem.20100244.
S.F. Bunting, E. Callen, N. Wong, H.T. Chen, F. Polato, A. Gunn, A. Bothmer, N.
Feldhahn, O. Fernandez-Capetillo, L. Cao, X. Xu, C.X. Deng, T. Finkel, M. Nussenzweig, J.M. Stark, A. Nussenzweig, 53BP1 inhibits homologous recombination in Brca1-deficient cells by blocking resection of DNA breaks, Cell 141 (2010)
243–254. , http://dx.doi.org/10.1016/j.cell.2010.03.012.
C. Escribano-Diaz, A. Orthwein, A. Fradet-Turcotte, M. Xing, J.T. Young, J. Tkac,
M.A. Cook, A.P. Rosebrock, M. Munro, M.D. Canny, D. Xu, D. Durocher, A cell
cycle-dependent regulatory circuit composed of 53BP1-RIF1 and BRCA1-CtIP
controls DNA repair pathway choice, Mol. Cell 49 (2013) 872–883. , http://
dx.doi.org/10.1016/j.molcel.2013.01.001.
L. Wei, L. Lan, Z. Hong, A. Yasui, C. Ishioka, N. Chiba, Rapid recruitment of BRCA1
to DNA double-strand breaks is dependent on its association with Ku80,
Mol. Cell. Biol. 28 (2008) 7380–7393. , http://dx.doi.org/10.1128/MCB.
01075-08.
G. Jiang, I. Plo, T. Wang, M. Rahman, J.H. Cho, E. Yang, B.S. Lopez, F. Xia, BRCA1Ku80 protein interaction enhances end-joining fidelity of chromosomal doublestrand breaks in the G1 phase of the cell cycle, J. Biol. Chem. 288 (2013) 8966–
8976. , http://dx.doi.org/10.1074/jbc.M112.412650.
W.Y. Lin, J.H. Wilson, Y. Lin, Repair of chromosomal double-strand breaks by
precise ligation in human cells, DNA Repair (Amst.) 12 (2013) 480–487. , http://
dx.doi.org/10.1016/j.dnarep.2013.04.024.
H.E. Krokan, M. Bjoras, Base excision repair, Cold Spring Harb. Perspect. Biol. 5
(2013), http://dx.doi.org/10.1101/cshperspect.a012583a012583.
H. Li, T. Marple, P. Hasty, Ku80-deleted cells are defective at base excision repair,
Mutat. Res. 745–746 (2013) 16–25. , http://dx.doi.org/10.1016/j.mrfmmm.
2013.03.010.
Y.J. Choi, H. Li, M.Y. Son, X.H. Wang, J.L. Fornsaglio, R.W. Sobol, M. Lee, J. Vijg, S.
Imholz, M.E. Dolle, H. van Steeg, E. Reiling, P. Hasty, Deletion of individual Ku
subunits in mice causes an NHEJ-independent phenotype potentially by altering
apurinic/apyrimidinic site repair, PLOS ONE 9 (2014) e86358, http://dx.doi.org/
10.1371/journal.pone.0086358.
P. Li, S. Gao, L. Wang, F. Yu, J. Li, C. Wang, J. Li, J. Wong, ABH2 couples regulation of
ribosomal DNA transcription with DNA alkylation repair, Cell Rep. 4 (2013) 817–
829. , http://dx.doi.org/10.1016/j.celrep.2013.07.027.
G. Giglia-Mari, A. Zotter, W. Vermeulen, DNA damage response, Cold Spring
Harb. Perspect. Biol. 3 (2011), http://dx.doi.org/10.1101/cshperspect.a000745,
a000745.
A. Marechal, L. Zou, DNA damage sensing by the ATM and ATR kinases, Cold
Spring Harb. Perspect. Biol. 5 (2013), http://dx.doi.org/10.1101/cshperspect.a012716.
Y. Shiloh, Y. Ziv, The ATM protein kinase: regulating the cellular response to
genotoxic stress, and more, Nature reviews, Mol. Cell Biol. 14 (2013) 197–210.
E.A. Nam, D. Cortez, ATR signalling: more than meeting at the fork, Biochem. J.
436 (2011) 527–536. , http://dx.doi.org/10.1042/BJ20102162.
N. Tomimatsu, C.G. Tahimic, A. Otsuki, S. Burma, A. Fukuhara, K. Sato, G. Shiota,
M. Oshimura, D.J. Chen, A. Kurimasa, Ku70/80 modulates ATM and ATR signaling
pathways in response to DNA double strand breaks, J. Biol. Chem. 282 (2007)
10138–10145. , http://dx.doi.org/10.1074/jbc.M611880200.
X.Y. Zhou, X. Wang, H. Wang, D.J. Chen, G.C. Li, G. Iliakis, Y. Wang, Ku affects the
ATM-dependent S phase checkpoint following ionizing radiation, Oncogene 21
(2002) 6377–6381. , http://dx.doi.org/10.1038/sj.onc.1205782.
T.H. Stracker, J.H. Petrini, The MRE11 complex: starting from the ends, Nat. Rev.
Mol. Cell Biol. 12 (2011) 90–103. , http://dx.doi.org/10.1038/nrm3047.
A.D. Amsel, M. Rathaus, N. Kronman, H.Y. Cohen, Regulation of the proapoptotic
factor Bax by Ku70-dependent deubiquitylation, Proc. Natl. Acad. Sci. U. S. A. 105
(2008) 5117–5122. , http://dx.doi.org/10.1073/pnas.0706700105.
M. Sawada, W. Sun, P. Hayes, K. Leskov, D.A. Boothman, S. Matsuyama, Ku70
suppresses the apoptotic translocation of Bax to mitochondria, Nat. Cell Biol. 5
(2003) 320–329. , http://dx.doi.org/10.1038/ncb950.
K. Iijima, C. Muranaka, J. Kobayashi, S. Sakamoto, K. Komatsu, S. Matsuura, N.
Kubota, H. Tauchi, NBS1 regulates a novel apoptotic pathway through Bax
activation, DNA Repair (Amst.) 7 (2008) 1705–1716. , http://dx.doi.org/
10.1016/j.dnarep.2008.06.013.
S. Mazumder, D. Plesca, M. Kinter, A. Almasan, Interaction of a cyclin E fragment
with Ku70 regulates Bax-mediated apoptosis, Mol. Cell. Biol. 27 (2007) 3511–
3520. , http://dx.doi.org/10.1128/MCB. 01448-06.
C. Subramanian, J.A. Jarzembowski, A.W. Opipari Jr., V.P. Castle, R.P. Kwok,
HDAC6 deacetylates Ku70 and regulates Ku70-Bax binding in neuroblastoma,
Neoplasia 13 (2011) 726–734.
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
[143] Z. Yuan, E. Seto, A functional link between SIRT1 deacetylase and NBS1 in DNA
damage response, Cell Cycle 6 (2007) 2869–2871. , http://dx.doi.org/10.4161/
cc.6.23.5026.
[144] B. Wang, M. Xie, R. Li, T.K. Owonikoko, S.S. Ramalingam, F.R. Khuri, W.J.
Curran, Y. Wang, X. Deng, Role of Ku70 in deubiquitination of Mcl-1 and
suppression of apoptosis, Cell Death Differ. (2014), http://dx.doi.org/10.1038/
cdd.2014.42.
[145] J.C. Martinou, R.J. Youle, Mitochondria in apoptosis: Bcl-2 family members and
mitochondrial dynamics, Dev. Cell 21 (2011) 92–101. , http://dx.doi.org/
10.1016/j.devcel.2011.06.017.
[146] M.P. Longhese, S. Anbalagan, M. Martina, D. Bonetti, The role of shelterin in
maintaining telomere integrity, Front. Biosci. 17 (2012) 1715–1728.
[147] C.M. Price, K.A. Boltz, M.F. Chaiken, J.A. Stewart, M.A. Beilstein, D.E. Shippen,
Evolution of CST function in telomere maintenance, Cell Cycle 9 (2010) 3157–
3165. , http://dx.doi.org/10.4161/cc.9.16.12547.
[148] S.E. Porter, P.W. Greenwell, K.B. Ritchie, T.D. Petes, The DNA-binding protein
Hdf1p (a putative Ku homologue) is required for maintaining normal telomere
length in Saccharomyces cerevisiae, Nucleic Acids Res. 24 (1996) 582–585. ,
http://dx.doi.org/10.1093/nar/24.4.582.
[149] S.J. Boulton, S.P. Jackson, Identification of a Saccharomyces cerevisiae Ku80
homologue: roles in DNA double strand break rejoining and in telomeric
maintenance, Nucleic Acids Res. 24 (1996) 4639–4648. , http://dx.doi.org/
10.1093/nar/24.23.4639.
[150] S. Gravel, M. Larrivee, P. Labrecque, R.J. Wellinger, Yeast, Ku as a regulator of
chromosomal DNA end structure, Science 280 (1998) 741–744. , http://
dx.doi.org/10.1126/science.280.5364.741.
[151] S.H. Teo, S.P. Jackson, Identification of Saccharomyces cerevisiae DNA ligase IV:
involvement in DNA double-strand break repair, EMBO J. 16 (1997) 4788–4795. ,
http://dx.doi.org/10.1093/emboj/16.15.4788.
[152] F. d’Adda di Fagagna, M.P. Hande, W.M. Tong, D. Roth, P.M. Lansdorp, Z.Q. Wang,
S.P. Jackson, Effects of DNA nonhomologous end-joining factors on telomere
length and chromosomal stability in mammalian cells, Curr. Biol. 11 (2001)
1192–1196. , http://dx.doi.org/10.1016/S0960-9822(01)00328-1.
[153] E. Samper, F.A. Goytisolo, P. Slijepcevic, P.P. van Buul, M.A. Blasco, Mammalian
Ku86 protein prevents telomeric fusions independently of the length of TTAGGG
repeats and the G-strand overhang, EMBO Rep. 1 (2000) 244–252. , http://
dx.doi.org/10.1093/embo-reports/kvd051.
[154] S. Espejel, S. Franco, S. Rodriguez-Perales, S.D. Bouffler, J.C. Cigudosa, M.A. Blasco,
Mammalian Ku86 mediates chromosomal fusions and apoptosis caused by
critically short telomeres, EMBO J. 21 (2002) 2207–2219. , http://dx.doi.org/
10.1093/emboj/21.9.2207.
[155] Y. Wang, G. Ghosh, E.A. Hendrickson, Ku86 represses lethal telomere deletion
events in human somatic cells, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 12430–
12435. , http://dx.doi.org/10.1073/pnas.0903362106.
[156] G.B. Celli, E.L. Denchi, T. de Lange, Ku70 stimulates fusion of dysfunctional
telomeres yet protects chromosome ends from homologous recombination, Nat.
Cell Biol. 8 (2006) 885–890. , http://dx.doi.org/10.1038/ncb1444.
[157] A. Sfeir, T. de Lange, Removal of shelterin reveals the telomere end-protection
problem, Science 336 (2012) 593–597. , http://dx.doi.org/10.1126/science.1218498.
[158] B. Fellerhoff, F. Eckardt-Schupp, A.A. Friedl, Subtelomeric repeat amplification is
associated with growth at elevated temperature in yku70 mutants of Saccharomyces cerevisiae, Genetics 154 (2000) 1039–1051.
[159] V. Lundblad, E.H. Blackburn, An alternative pathway for yeast telomere maintenance rescues est1-senescence, Cell 73 (1993) 347–360, pii:00928674(93)90234-H.
[160] L. Maringele, D. Lydall, EXO1-dependent single-stranded DNA at telomeres
activates subsets of DNA damage and spindle checkpoint pathways in budding
yeast yku70Delta mutants, Genes Dev. 16 (2002) 1919–1933. , http://dx.doi.org/
10.1101/gad.225102.
[161] A.E. Stellwagen, Z.W. Haimberger, J.R. Veatch, D.E. Gottschling, Ku interacts with
telomerase RNA to promote telomere addition at native and broken chromosome ends, Genes Dev. 17 (2003) 2384–2395. , http://dx.doi.org/10.1101/
gad.1125903.
[162] T.S. Fisher, A.K. Taggart, V.A. Zakian, Cell cycle-dependent regulation of yeast
telomerase by Ku, Nat. Struct. Mol. Biol. 11 (2004) 1198–1205. , http://
dx.doi.org/10.1038/nsmb854.
[163] S.E. Peterson, A.E. Stellwagen, S.J. Diede, M.S. Singer, Z.W. Haimberger, C.O.
Johnson, M. Tzoneva, D.E. Gottschling, The function of a stem-loop in telomerase
RNA is linked to the DNA repair protein Ku, Nat. Genet. 27 (2001) 64–67. , http://
dx.doi.org/10.1038/83778.
[164] A.D. Mozdy, E.R. Podell, T.R. Cech, Multiple yeast genes, including Paf1 complex
genes, affect telomere length via telomerase RNA abundance, Mol. Cell. Biol. 28
(2008) 4152–4161. , http://dx.doi.org/10.1128/MCB. 00512-08.
[165] D.C. Zappulla, K.J. Goodrich, J.R. Arthur, L.A. Gurski, E.M. Denham, A.E. Stellwagen, T.R. Cech, Ku can contribute to telomere lengthening in yeast at multiple
positions in the telomerase RNP, RNA 17 (2011) 298–311. , http://dx.doi.org/
10.1261/rna.2483611.
[166] F. Gallardo, C. Olivier, A.T. Dandjinou, R.J. Wellinger, P. Chartrand, TLC1 RNA
nucleo-cytoplasmic trafficking links telomerase biogenesis to its recruitment to
telomeres, EMBO J. 27 (2008) 748–757. , http://dx.doi.org/10.1038/
emboj.2008.21.
[167] N.S. Ting, Y. Yu, B. Pohorelic, S.P. Lees-Miller, T.L. Beattie, Human Ku70/80
interacts directly with hTR, the RNA component of human telomerase, Nucleic
Acids Res. 33 (2005) 2090–2098. , http://dx.doi.org/10.1093/nar/gki342.
27
[168] W. Chai, L.P. Ford, L. Lenertz, W.E. Wright, J.W. Shay, Human Ku70/80 associates
physically with telomerase through interaction with hTERT, J. Biol. Chem. 277
(2002) 47242–47247. , http://dx.doi.org/10.1074/jbc.M208542200.
[169] K. Mekhail, D. Moazed, The nuclear envelope in genome organization, expression and stability, Nat. Rev. Mol. Cell Biol. 11 (2010) 317–328. , http://dx.doi.org/
10.1038/nrm2894.
[170] S. Kueng, M. Oppikofer, S.M. Gasser, SIR proteins and the assembly of silent
chromatin in budding yeast, Annu. Rev. Genet. 47 (2013) 275–306. , http://
dx.doi.org/10.1146/annurev-genet-021313-173730.
[171] O.M. Aparicio, B.L. Billington, D.E. Gottschling, Modifiers of position effect are
shared between telomeric and silent mating-type loci in S. cerevisiae, Cell 66
(1991) 1279–1287.
[172] S.J. Boulton, S.P. Jackson, Components of the Ku-dependent non-homologous
end-joining pathway are involved in telomeric length maintenance and telomeric silencing, EMBO J. 17 (1998) 1819–1828. , http://dx.doi.org/10.1093/
emboj/17.6.1819.
[173] C.I. Nugent, G. Bosco, L.O. Ross, S.K. Evans, A.P. Salinger, J.K. Moore, J.E. Haber, V.
Lundblad, Telomere maintenance is dependent on activities required for end
repair of double-strand breaks, Curr. Biol. 8 (1998) 657–660.
[174] T. Laroche, S.G. Martin, M. Gotta, H.C. Gorham, F.E. Pryde, E.J. Louis, S.M. Gasser,
Mutation of yeast Ku genes disrupts the subnuclear organization of telomeres,
Curr. Biol. 8 (1998) 653–656.
[175] M. Gotta, T. Laroche, A. Formenton, L. Maillet, H. Scherthan, S.M. Gasser, The
clustering of telomeres and colocalization with Rap1, Sir3, and Sir4 proteins in
wild-type Saccharomyces cerevisiae, J. Cell Biol. 134 (1996) 1349–1363.
[176] A. Taddei, F. Hediger, F.R. Neumann, C. Bauer, S.M. Gasser, Separation of silencing
from perinuclear anchoring functions in yeast Ku80, Sir4 and Esc1 proteins,
EMBO J. 23 (2004) 1301–1312. , http://dx.doi.org/10.1038/sj.emboj.7600144.
[177] H. Schober, H. Ferreira, V. Kalck, L.R. Gehlen, S.M. Gasser, Yeast telomerase and
the SUN domain protein Mps3 anchor telomeres and repress subtelomeric
recombination, Genes Dev. 23 (2009) 928–938. , http://dx.doi.org/10.1101/
gad.1787509.
[178] F. Hediger, F.R. Neumann, G. Van Houwe, K. Dubrana, S.M. Gasser, Live imaging
of telomeres: yKu and Sir proteins define redundant telomere-anchoring pathways in yeast, Curr. Biol. 12 (2002) 2076–2089.
[179] H.L. Hsu, D. Gilley, E.H. Blackburn, D.J. Chen, Ku is associated with the telomere
in mammals, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 12454–12458. , http://
dx.doi.org/10.1073/pnas.96.22.12454.
[180] T.J. Wu, Y.H. Chiang, Y.C. Lin, C.R. Tsai, T.Y. Yu, M.T. Sung, Y.H. Lee, J.J. Lin,
Sequential loading of Saccharomyces cerevisiae Ku and Cdc13p to telomeres, J.
Biol. Chem. 284 (2009) 12801–12808. , http://dx.doi.org/10.1074/
jbc.M809131200.
[181] C.R. Lopez, A. Ribes-Zamora, S.M. Indiviglio, C.L. Williams, S. Haricharan, A.A.
Bertuch, Ku must load directly onto the chromosome end in order to mediate its
telomeric functions, PLoS Genet. 7 (2011) e1002233, http://dx.doi.org/10.1371/
journal.pgen.1002233.
[182] A. Bianchi, T. de Lange, Ku binds telomeric DNA in vitro, J. Biol. Chem. 274 (1999)
21223–21227.
[183] H.L. Hsu, D. Gilley, S.A. Galande, M.P. Hande, B. Allen, S.H. Kim, G.C. Li, J. Campisi,
T. Kohwi-Shigematsu, D.J. Chen, Ku acts in a unique way at the mammalian
telomere to prevent end joining, Genes Dev. 14 (2000) 2807–2812. , http://
dx.doi.org/10.1101/gad.844000.
[184] K. Song, D. Jung, Y. Jung, S.G. Lee, I. Lee, Interaction of human Ku70 with TRF2,
FEBS Lett. 481 (2000) 81–85. , http://dx.doi.org/10.1016/S0014-5793(00)
01958-X.
[185] L.S. Fink, C.A. Lerner, P.F. Torres, C. Sell, Ku80 facilitates chromatin binding of the
telomere binding protein, TRF2, Cell Cycle 9 (2010) 3798–3806. , http://
dx.doi.org/10.4161/cc.9.18.13129.
[186] M.S. O’Connor, A. Safari, D. Liu, J. Qin, Z. Songyang, The human Rap1 protein
complex and modulation of telomere length, J. Biol. Chem. 279 (2004) 28585–
28591. , http://dx.doi.org/10.1074/jbc.M312913200.
[187] J.Z. Ye, J.R. Donigian, M. van Overbeek, D. Loayza, Y. Luo, A.N. Krutchinsky, B.T.
Chait, T. de Lange, TIN2 binds TRF1 and TRF2 simultaneously and stabilizes the
TRF2 complex on telomeres, J. Biol. Chem. 279 (2004) 47264–47271. , http://
dx.doi.org/10.1074/jbc.M409047200.
[188] M. Cockell, F. Palladino, T. Laroche, G. Kyrion, C. Liu, A.J. Lustig, S.M. Gasser, The
carboxy termini of Sir4 and Rap1 affect Sir3 localization: evidence for a multicomponent complex required for yeast telomeric silencing, J. Cell Biol. 129
(1995) 909–924.
[189] J.S. Pfingsten, K.J. Goodrich, C. Taabazuing, F. Ouenzar, P. Chartrand, T.R. Cech,
Mutually exclusive binding of telomerase RNA and DNA by Ku alters telomerase
recruitment model, Cell 148 (2012) 922–932. , http://dx.doi.org/10.1016/
j.cell.2012.01.033.
[190] M. Lei, E.R. Podell, T.R. Cech, Structure of human POT1 bound to telomeric singlestranded DNA provides a model for chromosome end-protection, Nat. Struct.
Mol. Biol. 11 (2004) 1223–1229. , http://dx.doi.org/10.1038/nsmb867.
[191] D. Liu, A. Safari, M.S. O’Connor, D.W. Chan, A. Laegeler, J. Qin, Z. Songyang, PTOP
interacts with POT1 and regulates its localization to telomeres, Nat. Cell Biol. 6
(2004) 673–680. , http://dx.doi.org/10.1038/ncb1142.
[192] Y. Doksani, J.Y. Wu, T. de Lange, X. Zhuang, Super-resolution fluorescence
imaging of telomeres reveals TRF2-dependent T-loop formation, Cell 155
(2013) 345–356. , http://dx.doi.org/10.1016/j.cell.2013.09.048.
[193] S. Malu, V. Malshetty, D. Francis, P. Cortes, Role of non-homologous end joining
in V(D)J recombination, Immunol. Res. 54 (2012) 233–246. , http://dx.doi.org/
10.1007/s12026-012-8329-z.
28
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
[194] H. Ouyang, A. Nussenzweig, A. Kurimasa, V.C. Soares, X. Li, C. Cordon-Cardo, W.
Li, N. Cheong, M. Nussenzweig, G. Iliakis, D.J. Chen, G.C. Li, Ku70 is required for
DNA repair but not for T cell antigen receptor gene recombination in vivo, J. Exp.
Med. 186 (1997) 921–929. , http://dx.doi.org/10.1084/jem.186.6.921.
[195] J.P. Manis, Y. Gu, R. Lansford, E. Sonoda, R. Ferrini, L. Davidson, K. Rajewsky, F.W.
Alt, Ku70 is required for late B cell development and immunoglobulin heavy
chain class switching, J. Exp. Med. 187 (1998) 2081–2089. , http://dx.doi.org/
10.1084/jem.187.12.2081.
[196] Y. Gu, K.J. Seidl, G.A. Rathbun, C. Zhu, J.P. Manis, N. van der Stoep, L. Davidson,
H.L. Cheng, J.M. Sekiguchi, K. Frank, P. Stanhope-Baker, M.S. Schlissel, D.B. Roth,
F.W. Alt, Growth retardation and leaky SCID phenotype of Ku70-deficient mice,
Immunity 7 (1997) 653–665. , http://dx.doi.org/10.1016/S1074-7613(00)
80386-6.
[197] M.A. Slatter, A.R. Gennery, Primary immunodeficiency syndromes, Adv. Exp.
Med. Biol. 685 (2010) 146–165.
[198] D. Moshous, I. Callebaut, R. de Chasseval, B. Corneo, M. Cavazzana-Calvo, F. Le
Deist, I. Tezcan, O. Sanal, Y. Bertrand, N. Philippe, A. Fischer, J.P. de Villartay,
Artemis, a novel DNA double-strand break repair/V(D)J recombination protein,
is mutated in human severe combined immune deficiency, Cell 105 (2001) 177–
186. , http://dx.doi.org/10.1016/S0092-8674(01)00309-9.
[199] M. O’Driscoll, K.M. Cerosaletti, P.M. Girard, Y. Dai, M. Stumm, B. Kysela, B. Hirsch,
A. Gennery, S.E. Palmer, J. Seidel, R.A. Gatti, R. Varon, M.A. Oettinger, H. Neitzel,
P.A. Jeggo, P. Concannon, DNA ligase IV mutations identified in patients exhibiting developmental delay and immunodeficiency, Mol. Cell 8 (2001) 1175–1185. ,
http://dx.doi.org/10.1016/S1097-2765(01)00408-7.
[200] D. Buck, L. Malivert, R. de Chasseval, A. Barraud, M.C. Fondaneche, O. Sanal, A.
Plebani, J.L. Stephan, M. Hufnagel, F. le Deist, A. Fischer, A. Durandy, J.P. de
Villartay, P. Revy, Cernunnos, a novel nonhomologous end-joining factor, is
mutated in human immunodeficiency with microcephaly, Cell 124 (2006) 287–
299. , http://dx.doi.org/10.1016/j.cell.2005.12.030.
[201] L. Woodbine, J.A. Neal, N.K. Sasi, M. Shimada, K. Deem, H. Coleman, W.B. Dobyns,
T. Ogi, K. Meek, E.G. Davies, P.A. Jeggo, PRKDC mutations in a SCID patient with
profound neurological abnormalities, J. Clin. Invest. 123 (2013) 2969–2980. ,
http://dx.doi.org/10.1172/JCI67349.
[202] M. van der Burg, J.J. van Dongen, D.C. van Gent, DNA-PKcs deficiency in human:
long predicted, finally found, Curr. Opin. Allergy Clin. Immunol. 9 (2009) 503–
509. , http://dx.doi.org/10.1097/ACI.0b013e3283327e41.
[203] G.C. Li, H. Ouyang, X. Li, H. Nagasawa, J.B. Little, D.J. Chen, C.C. Ling, Z. Fuks, C.
Cordon-Cardo, Ku70: a candidate tumor suppressor gene for murine T cell
lymphoma, Mol. Cell 2 (1998) 1–8. , http://dx.doi.org/10.1016/S10972765(00)80108-2.
[204] H. Vogel, D.S. Lim, G. Karsenty, M. Finegold, P. Hasty, Deletion of Ku86 causes
early onset of senescence in mice, Proc. Natl. Acad. Sci. U. S. A. 96 (1999) 10770–
10775. , http://dx.doi.org/10.1073/pnas.96.19.10770.
[205] A. Nussenzweig, C. Chen, V. da Costa Soares, M. Sanchez, K. Sokol, M.C. Nussenzweig, G.C. Li, Requirement for Ku80 in growth and immunoglobulin V(D)J
recombination, Nature 382 (1996) 551–555. , http://dx.doi.org/10.1038/
382551a0.
[206] H. Li, H. Vogel, V.B. Holcomb, Y. Gu, P. Hasty, Deletion of Ku70, Ku80, or both
causes early aging without substantially increased cancer, Mol. Cell. Biol. 27
(2007) 8205–8214. , http://dx.doi.org/10.1128/MCB. 00785-07.
[207] K. Ren, S. Pena de Ortiz, Non-homologous DNA end joining in the mature rat
brain, J. Neurochem. 80 (2002) 949–959. , http://dx.doi.org/10.1046/j.00223042.2002.00776.x.
[208] V.N. Vyjayanti, K.S. Rao, DNA double strand break repair in brain: reduced NHEJ
activity in aging rat neurons, Neurosci. Lett. 393 (2006) 18–22. , http://
dx.doi.org/10.1016/j.neulet.2005.09.053.
[209] D.A. Shackelford, DNA end joining activity is reduced in Alzheimer’s disease,
Neurobiol. Aging 27 (2006) 596–605. , http://dx.doi.org/10.1016/j.neurobiolaging.2005.03.009.
[210] P.M. Reaper, F. di Fagagna, S.P. Jackson, Activation of the DNA damage response
by telomere attrition: a passage to cellular senescence, Cell Cycle 3 (2004) 543–
546. , http://dx.doi.org/10.4161/cc.3.5.835.
[211] M.A. Shammas, Telomeres, lifestyle, cancer, and aging, Curr. Opin. Clin. Nutr.
Metab. Care 14 (2011) 28–34. , http://dx.doi.org/10.1097/MCO.0b013e
32834121b1.
[212] D. Hanahan, R.A. Weinberg, Hallmarks of cancer: the next generation, Cell 144
(2011) 646–674. , http://dx.doi.org/10.1016/j.cell.2011.02.013.
[213] J.M. Luk, Y.C. Su, S.C. Lam, C.K. Lee, M.Y. Hu, Q.Y. He, G.K. Lau, F.W. Wong, S.T. Fan,
Proteomic identification of Ku70/Ku80 autoantigen recognized by monoclonal
antibody against hepatocellular carcinoma, Proteomics 5 (2005) 1980–1986. ,
http://dx.doi.org/10.1002/pmic.200401084.
[214] M. Korabiowska, J. Voltmann, J.F. Honig, P. Bortkiewicz, F. Konig, C. CordonCardo, F. Jenckel, P. Ambrosch, G. Fischer, Altered expression of DNA doublestrand repair genes Ku70 and Ku80 in carcinomas of the oral cavity, Anticancer
Res. 26 (2006) 2101–2105.
[215] P. Mazzarelli, P. Parrella, D. Seripa, E. Signori, G. Perrone, C. Rabitti, D. Borzomati,
A. Gabbrielli, M.G. Matera, C. Gravina, M. Caricato, M.L. Poeta, M. Rinaldi, S.
Valeri, R. Coppola, V.M. Fazio, DNA end binding activity and Ku70/80 heterodimer expression in human colorectal tumor, World J. Gastroenterol.: WJG 11
(2005) 6694–6700.
[216] P. Parrella, P. Mazzarelli, E. Signori, G. Perrone, G.F. Marangi, C. Rabitti, M.
Delfino, M. Prencipe, A.P. Gallo, M. Rinaldi, G. Fabbrocini, S. Delfino, P. Persichetti, V.M. Fazio, Expression and heterodimer-binding activity of Ku70 and
Ku80 in human non-melanoma skin cancer, J. Clin. Pathol. 59 (2006) 1181–1185.
, http://dx.doi.org/10.1136/jcp.2005.031088.
[217] K. Abdelbaqi, D. Di Paola, E. Rampakakis, M. Zannis-Hadjopoulos, Ku protein
levels, localization and association to replication origins in different stages of
breast tumor progression, J. Cancer 4 (2013) 358–370. , http://dx.doi.org/
10.7150/jca.6289.
[218] Y. Komuro, T. Watanabe, Y. Hosoi, Y. Matsumoto, K. Nakagawa, N. Tsuno, S.
Kazama, J. Kitayama, N. Suzuki, H. Nagawa, The expression pattern of Ku
correlates with tumor radiosensitivity and disease free survival in patients with
rectal carcinoma, Cancer 95 (2002) 1199–1205. , http://dx.doi.org/10.1002/
cncr.10807.
[219] C.J. Lord, A. Ashworth, The DNA damage response and cancer therapy, Nature
481 (2012) 287–294. , http://dx.doi.org/10.1038/nature10760.
[220] C.H. Kim, S.J. Park, S.H. Lee, A targeted inhibition of DNA-dependent
protein kinase sensitizes breast cancer cells following ionizing radiation, J.
Pharmacol. Exp. Ther. 303 (2002) 753–759. , http://dx.doi.org/10.1124/
jpet.102.038505.
[221] L.R. Bertolini, M. Bertolini, G.B. Anderson, E.A. Maga, K.R. Madden, J.D. Murray,
Transient depletion of Ku70 and Xrcc4 by RNAi as a means to manipulate the
non-homologous end-joining pathway, J. Biotechnol. 128 (2007) 246–257. ,
http://dx.doi.org/10.1016/j.jbiotec.2006.10.003.
[222] I.S. Ayene, L.P. Ford, C.J. Koch, Ku protein targeting by Ku70 small interfering RNA
enhances human cancer cell response to topoisomerase II inhibitor and gamma
radiation, Mol Cancer Ther 4 (2005) 529–536. , http://dx.doi.org/10.1158/15357163.MCT-04-0130.
[223] P. Zhu, D. Zhang, D. Chowdhury, D. Martinvalet, D. Keefe, L. Shi, J. Lieberman,
Granzyme A, which causes single-stranded DNA damage, targets the doublestrand break repair protein Ku70, EMBO Rep. 7 (2006) 431–437. , http://
dx.doi.org/10.1038/sj.embor.7400622.
[224] T. Yoshida, I. Tomioka, T. Nagahara, T. Holyst, M. Sawada, P. Hayes, V. Gama, M.
Okuno, Y. Chen, Y. Abe, T. Kanouchi, H. Sasada, D. Wang, T. Yokota, E. Sato, S.
Matsuyama, Bax-inhibiting peptide derived from mouse and rat Ku70, Biochem.
Biophys. Res. Commun. 321 (2004) 961–966. , http://dx.doi.org/10.1016/j.bbrc.
2004.07.054.
[225] M. Sawada, P. Hayes, S. Matsuyama, Cytoprotective membrane-permeable
peptides designed from the Bax-binding domain of Ku70, Nat. Cell Biol. 5
(2003) 352–357. , http://dx.doi.org/10.1038/ncb955.
[226] C. Subramanian, A.W. Opipari Jr., X. Bian, V.P. Castle, R.P. Kwok, Ku70 acetylation
mediates neuroblastoma cell death induced by histone deacetylase inhibitors,
Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 4842–4847. , http://dx.doi.org/10.1073/
pnas.0408351102.
[227] F.M. Hsu, S. Zhang, B.P. Chen, Role of DNA-dependent protein kinase catalytic
subunit in cancer development and treatment, Transl. Cancer Res. 1 (2012) 22–
34. , http://dx.doi.org/10.3978/j.issn.2218-676X.0104.
[228] R.B. West, M. Yaneva, M.R. Lieber, Productive and nonproductive complexes of
Ku and DNA-dependent protein kinase at DNA termini, Mol. Cell. Biol. 18 (1998)
5908–5920.
[229] K.N. Mahajan, S.A. Nick McElhinny, B.S. Mitchell, D.A. Ramsden, Association of
DNA polymerase mu (pol mu) with Ku and ligase IV: role for pol mu in endjoining double-strand break repair, Mol. Cell. Biol. 22 (2002) 5194–5202. , http://
dx.doi.org/10.1128/MCB.22.14.5194-5202.2002.
[230] Y. Ma, H. Lu, B. Tippin, M.F. Goodman, N. Shimazaki, O. Koiwai, C.L. Hsieh, K.
Schwarz, M.R. Lieber, A biochemically defined system for mammalian nonhomologous DNA end joining, Mol. Cell 16 (2004) 701–713. , http://dx.doi.org/
10.1016/j.molcel.2004.11.017.
[231] M.M. Purugganan, S. Shah, J.F. Kearney, D.B. Roth, Ku80 is required for addition of
N nucleotides to V(D)J recombination junctions by terminal deoxynucleotidyl
transferase, Nucleic Acids Res. 29 (2001) 1638–1646. , http://dx.doi.org/
10.1093/nar/29.7.1638.
[232] P. Karmakar, C.M. Snowden, D.A. Ramsden, V.A. Bohr, Ku heterodimer binds to
both ends of the Werner protein and functional interaction occurs at the Werner
N-terminus, Nucleic Acids Res. 30 (2002) 3583–3591.
[233] P. Raval, A.N. Kriatchko, S. Kumar, P.C. Swanson, Evidence for Ku70/Ku80
association with full-length RAG1, Nucleic Acids Res. 36 (2008) 2060–2072. ,
http://dx.doi.org/10.1093/nar/gkn049.
[234] A. Kinner, W. Wu, C. Staudt, G. Iliakis, Gamma-H2AX in recognition and signaling
of DNA double-strand breaks in the context of chromatin, Nucleic Acids Res. 36
(2008) 5678–5694. , http://dx.doi.org/10.1093/nar/gkn550.
[235] M. Hoek, M.P. Myers, B. Stillman, An analysis of CAF-1-interacting proteins
reveals dynamic and direct interactions with the KU complex and 14-3-3
proteins, J. Biol. Chem. 286 (2011) 10876–10887. , http://dx.doi.org/10.1074/
jbc.M110.217075.
[236] J.C. Morales, P. Richard, A. Rommel, F.J. Fattah, E.A. Motea, P.L. Patidar, L. Xiao, K.
Leskov, S.Y. Wu, W.N. Hittelman, C.M. Chiang, J.L. Manley, D.A. Boothman, Kub5Hera, the human Rtt103 homolog, plays dual functional roles in transcription
termination and DNA repair, Nucleic Acids Res. (2014), http://dx.doi.org/
10.1093/nar/gku160.
[237] S. Galande, T. Kohwi-Shigematsu, Poly(ADP-ribose) polymerase and Ku autoantigen form a complex and synergistically bind to matrix attachment
sequences, J. Biol. Chem. 274 (1999) 20521–20528.
[238] A. Shahi, J.H. Lee, Y. Kang, S.H. Lee, J.W. Hyun, I.Y. Chang, J.Y. Jun, H.J. You,
Mismatch-repair protein MSH6 is associated with Ku70 and regulates DNA
double-strand break repair, Nucleic Acids Res. 39 (2011) 2130–2143. , http://
dx.doi.org/10.1093/nar/gkq1095.
[239] S. Parvathaneni, A. Stortchevoi, J.A. Sommers, R.M. Brosh Jr., S. Sharma, Human
RECQ1 interacts with Ku70/80 and modulates DNA end-joining of double-strand
breaks, PLOS ONE 8 (2013) e62481, http://dx.doi.org/10.1371/journal.
pone.0062481.
V.L. Fell, C. Schild-Poulter / Mutation Research 763 (2015) 15–29
[240] W. Goedecke, M. Eijpe, H.H. Offenberg, M. van Aalderen, C. Heyting, Mre11 and
Ku70 interact in somatic cells, but are differentially expressed in early meiosis,
Nat. Genet. 23 (1999) 194–198. , http://dx.doi.org/10.1038/13821.
[241] A. Feki, C.E. Jefford, P. Berardi, J.Y. Wu, L. Cartier, K.H. Krause, I. Irminger-Finger,
BARD1 induces apoptosis by catalysing phosphorylation of p53 by DNA-damage
response kinase, Oncogene 24 (2005) 3726–3736. , http://dx.doi.org/10.1038/
sj.onc.1208491.
[242] K. Song, Y. Jung, D. Jung, I. Lee, Human Ku70 interacts with heterochromatin
protein 1alpha, J. Biol. Chem. 276 (2001) 8321–8327. , http://dx.doi.org/
10.1074/jbc.M008779200.
[243] K.S. Leskov, D.Y. Klokov, J. Li, T.J. Kinsella, D.A. Boothman, Synthesis and
functional analyses of nuclear clusterin, a cell death protein, J. Biol. Chem.
278 (2003) 11590–11600. , http://dx.doi.org/10.1074/jbc.M209233200.
[244] H. Liu, C.H. Herrmann, K. Chiang, T.L. Sung, S.H. Moon, L.A. Donehower, A.P. Rice,
55K isoform of CDK9 associates with Ku70 and is involved in DNA repair,
Biochem. Biophys. Res. Commun. 397 (2010) 245–250. , http://dx.doi.org/
10.1016/j.bbrc.2010.05.092.
[245] U. Chung, T. Igarashi, T. Nishishita, H. Iwanari, A. Iwamatsu, A. Suwa, T. Mimori,
K. Hata, S. Ebisu, E. Ogata, T. Fujita, T. Okazaki, The interaction between Ku
antigen and REF1 protein mediates negative gene regulation by extracellular
calcium, J. Biol. Chem. 271 (1996) 8593–8598.
[246] Y. Zheng, Z. Ao, B. Wang, K.D. Jayappa, X. Yao, Host protein Ku70 binds and
protects HIV-1 integrase from proteasomal degradation and is required for HIV
replication, J. Biol. Chem. 286 (2011) 17722–17735. , http://dx.doi.org/10.1074/
jbc.M110.184739.
29
[247] H. Wang, R. Fang, J.Y. Cho, T.A. Libermann, P. Oettgen, Positive and negative
modulation of the transcriptional activity of the ETS factor ESE-1 through
interaction with p300, CREB-binding protein, and Ku 70/86, J. Biol. Chem.
279 (2004) 25241–25250. , http://dx.doi.org/10.1074/jbc.M401356200.
[248] W. Kaczmarski, S.A. Khan, Lupus autoantigen Ku protein binds HIV-1 TAR RNA in
vitro, Biochem. Biophys. Res. Commun. 196 (1993) 935–942. , http://dx.doi.org/
10.1006/bbrc.1993.2339.
[249] D.M. Willis, A.P. Loewy, N. Charlton-Kachigian, J.S. Shao, D.M. Ornitz, D.A.
Towler, Regulation of osteocalcin gene expression by a novel Ku antigen
transcription factor complex, J. Biol. Chem. 277 (2002) 37280–37291. , http://
dx.doi.org/10.1074/jbc.M206482200.
[250] J. Huang, A. Nueda, S. Yoo, W.S. Dynan, Heat shock transcription factor 1 binds
selectively in vitro to Ku protein and the catalytic subunit of the DNA-dependent
protein kinase, J. Biol. Chem. 272 (1997) 26009–26016.
[251] D. Matheos, M.T. Ruiz, G.B. Price, M. Zannis-Hadjopoulos, Ku antigen, an originspecific binding protein that associates with replication proteins, is required for
mammalian DNA replication, Biochim. Biophys. Acta 1578 (2002) 59–72. ,
http://dx.doi.org/10.1016/S0167-4781(02)00497-9.
[252] R.J. Wellinger, V.A. Zakian, Everything you ever wanted to know about Saccharomyces cerevisiae telomeres: beginning to end, Genetics 191 (2012) 1073–1105.
, http://dx.doi.org/10.1534/genetics.111.137851.
[253] R.J. O’Sullivan, J. Karlseder, Telomeres: protecting chromosomes against genome
instability, Nat. Rev. Mol. Cell Biol. 11 (2010) 171–181. , http://dx.doi.org/
10.1038/nrm2848.