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Current Molecular Pharmacology, 2012, 5, 88-101
Base Excision Repair, the Redox Environment and Therapeutic Implications
S.J. Storr, C.M. Woolston and S.G. Martin*
Academic Oncology, University of Nottingham, School of Molecular Medical Sciences, Nottingham University Hospitals
NHS Trust, City Hospital Campus, Nottingham, NG5 1PB, UK
Abstract: Control of redox homeostasis is crucial for a number of cellular processes with deregulation leading to a
number of serious consequences including oxidative damage such induction of DNA base lesions. The DNA lesions
caused by oxidative damage are principally repaired by the base excision repair (BER) pathway. Pharmacological
inhibition of BER is becoming an increasingly active area of research with the emergence of PARP inhibitors in cancer
therapy. The redox status of the cell is modulated by a number of systems, including a large number of anti-oxidant
enzymes who function in the control of superoxide and hydrogen peroxide, and ultimately in the release of the damaging
hydroxyl radical. Here we provide an overview of reactive oxygen species (ROS) production and its modulation by antioxidant enzymes. The review also discusses the effect of ROS on the BER pathway, particularly in relation to cancer.
Finally, as the modulation of the redox environment is of interest in cancer therapy, with certain agents having the
potential to reverse chemo- and radiotherapy resistance or treat therapy related toxicity, we discuss redox modulating
agents currently under development.
Keywords: Base excision repair, cancer, homeostasis, redox.
THE REDOX ENVIRONMENT
The term ‘redox’ is used to define the transfer of
electrons through reduction and oxidation and the control of
redox homeostasis is extremely important for normal cell
function and survival. The redox biochemistry that living
cells experience is dominated by oxygen. However cells
require a reducing environment to function and therefore
oxygen and its intermediates, collectively known as reactive
oxygen species (ROS), are a constant threat. Cells have
adapted to this problem through the development of a
complex mechanism of redox buffering systems that aid in
the control of ROS levels. These include the thioredoxin and
glutathione systems, antioxidants enzymes such as catalase
and superoxide dismutase and non-enzymatic antioxidants
such as ascorbic acid, -tocopherol and carotenoids. Such
control mechanisms, involved in the maintenance of redox
homeostasis, have been reviewed by Dröge, 2002 [1]. ROS
can be broadly categorised as free radicals such as
superoxide (O2•-) and hydroxyl radicals (•OH) or non radicals
such as hydrogen peroxide (H2O2). Free radicals are
molecules containing one or more unpaired electrons in
atomic or molecular orbitals. ROS are hazardous for living
organisms and can damage most major cellular constituents.
The accumulation of ROS within the cell can lead to
oxidative DNA damage, altered cytoplasmic and nuclear
signalling and can change the activity and/or expression of
proteins that respond to stress. There are a number of human
diseases where oxidative damage plays an influential role,
such as carcinogenesis, pre-eclampsia, stroke and chronic
heart failure, but also in the natural process of ageing [2].
*Address correspondence to this author at the Academic Oncology,
University of Nottingham, School of Molecular Medical Sciences,
Nottingham University Hospitals NHS Trust, City Hospital Campus,
Nottingham, NG5 1PB, UK; Tel: +44 (0)115 823 1846; Fax: +44 (0)115
823 1849; E-mail: [email protected]
1874-4672/12 $58.00+.00
Normal cellular metabolism is the major source of ROS
in aerobic cells. Mitochondria use NADH to funnel electrons
through the respiratory chain. During this process superoxide
can leak from the chain, principally through complexes I and
III. Other sources of ROS include membrane bound NADPH
oxidase (Nox), which is able to produce ROS involved in
cellular signalling triggered by receptor binding. In addition,
ROS are produced during pathological processes such as
inflammation by neutrophils, eosinophils and macrophages
[3]. Exogenous sources of ROS include irradiation by UV
light and X-rays and atmospheric pollution. Metal induced
oxidative stress is reviewed by Valko et al. (2006) [4].
The fate of the ‘leaked’ superoxide can be either nonenzymatic or enzymatic dismutation to H2O2. The enzymes
involved in this reaction are superoxide dismutases (SOD)
and superoxide reductases. Superoxide that escapes
dismutation can participate in the Harber-Weiss reaction
with H2O2 to produce hydroxyl radicals, however, in-vivo
these radicals are principally formed through the Fenton
reaction when Fe2+ contacts H2O2. In addition, through
peroxynitrate formation, ‘leaked’ superoxide can also
interact with nitric oxide to form ONOO- which is a
powerful oxidant comparable to hydroxyl radicals [5]. H2O2
can specifically and reversibly modify proteins, principally
via thiol groups.
In the context of this review, the hydroxyl radical is
perhaps the most important ROS due to its highly reactive
nature. The hydroxyl radical is able to react with proteins
and DNA, and has no specific biological partner or agent to
modify its actions. Importantly there is no enzyme that
functions to supervise its removal from the cellular
environment. Therefore its precursor, H2O2, is tightly
regulated and it is when these control mechanisms fail that
damage can occur. However, the hydroxyl radical has a short
half life of less that 1 nano second [6]. In-vivo, peroxynitrite
may be one of the principal agents of oxidative damage as it
has a longer half life than the hydroxyl radical. In addition, it
© 2012 Bentham Science Publishers
Base Excision Repair, the Redox Environment
does not require iron chelation for production, purely nitric
oxide and superoxide; and nitric oxide is readily available,
produced by oxidising argenine [7, 8]. Furthermore the
reaction between the radicals superoxide and nitric oxide, in
the production of peroxynitrite have a rate constant 3.5 times
higher than that for SOD catalysed dismutation of
superoxide. The chemical biology of peroxynitrite is
reviewed by Ferrer-Sueta and Radi (2009) [9] and Burney et
al. (1999) [10].
Oxidative stress can trigger two major events, the
oxidation of cysteine containing proteins and the formation
of reactive sulphur species. These two events can propagate
pro- or anti-apoptotic pathways. Thiol oxidation can cause
the formation of disulphide bonds which can activate or
deactivate the activity of a protein, often by causing
conformational changes in protein structure [11]. The redox
sensitive sulfhydryl switches are central to maintaining a
balanced redox environment. In normal cells, oxidative
stress can cause a range of responses from a transient growth
arrest and adaptation, increase in cellular proliferation,
permanent growth arrest or senescence, apoptosis or necrosis
[12]. In cancer, abnormal cells are continuously under
increased oxidative stress due to accelerated cellular
proliferation, prolonged stimulation of growth promoting
signalling pathways and alterations in metabolic activity.
Cellular ROS are able to cause oxidative DNA damage in
addition to influencing the numerous aberrantly altered
processes. ROS can transmit further cellular signals that are
able to influence tumour progression, metabolic pathways
and alter transcription by modulating redox controlled
transcription factors including p53, activator protein 1 (AP1), and NF-B. In addition, ROS have been implicated in
epithelial-mesenchymal transition (EMT), angiogenesis and
cell migration and in the mechanism of action of therapeutic
interventions including chemotherapy and radiotherapy [3,
13]. ROS mediated molecular and biochemical changes
including DNA lesions and mutations, contribute to the
heterogenous nature of a cancer and are likely to aid in the
generation of therapy resistant subpopulations. The role of
ROS in oncogenic transformation has been reviewed by
Behrend et al. (2003) [14].
OXIDATIVE DAMAGE TO DNA
A high proportion of DNA damage can be attributed to
ROS, in particular the hydroxyl radical, and may partly
explain the induction of certain spontaneous cancers [15].
The generation of ROS, in particular hydrogen peroxide and
the hydroxyl radical, has been shown to be increased in
cancer [16]. The hydroxyl radical is able to cause DNA
damage by a variety of means including its addition to
double bonds of DNA bases and removal of hydrogen from
thymine and each of the C-H bonds of 2’-deoxyribose [4].
Each ROS acts via different mechanisms to damage DNA,
with superoxide and hydrogen peroxide not directly
responsible for oxidative damage, but involved through the
accumulation of the hydroxyl radical. The hydroxyl radical
can damage all bases, although preferentially acts to oxidise
guanine due to its low redox potential [17, 18].
Oxidative DNA damage can be repaired by a number of
mechanisms but principally by the base excision repair
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
89
(BER) pathway, which removes single lesions [19].
Interestingly there are a number of enzymes within the BER
pathway that can themselves be modulated by the redox
environment. The removal of larger base lesions containing
oligonucleotides is conducted by nucleotide excision repair
(NER). The process of oxidative damage repair is reviewed
by Barzilai and Yamamoto (2004) [20]. The BER process is
implicated in a number of pathologies such as cancer, ageing
and neurodegeneration [21]. All of these conditions result in
the accumulation of DNA damage, often through defective
BER enzymes. The pathway can be subdivided into either
short patch (SP) or long patch (LP), which differ in the
amount of DNA synthesised during repair, being one to two
bases or three to eight bases respectively. BER consists of a
number of steps starting with the identification and excision
of the lesion, and repair of the DNA. There are a large
number of enzymes that cooperate in the BER pathway,
through protein-protein interactions and post translational
modification, with a degree of overlap, and are reviewed by
Fan and Wilson III (2005) [22].
Oxidative damage to DNA can cause a plethora of base
lesions, the most frequently described of which is an
oxidised guanine, 8-oxo-2’deoxyguanosine (8-oxoG) and
there are numerous repair mechanisms to remove them. 8oxoG mutations can cause A:T to C:C and G:C to T:A
transversions [17]. In addition lesions can include thymine
glycol, 3-methyladenine and double strand breaks. Repair
intermediates such as apurinic/apyrimidinic (AP) sites are
also damaging to the cell. The repair of oxidised base lesions
is initiated by DNA glycosylases such as endonuclease III
homologue NTH1 and 8-oxoG DNA glycosylases (OGG1
and OGG2) which generate strand breaks by -elimination
(Fig. 1). As well as OGG1 and OGG2 there are a number of
other glycosylases that are involved in the process, often
recognising different lesions with different affinities such as
NEIL1 [23]. In addition, some degree of cooperativity exists
between the glycosylases, such as increased turnover of
OGG1 by NEIL1 [24]. This process results in 3’-end
structures which block DNA polymerase activity. In addition
these 3’ blocking structures can also be directly caused by
ROS. One of the most influential proteins in the BER
pathway is human AP endonuclease, APE1 (APEX, APE,
HAP1 and Ref-1), which cleaves phosphodiester bonds 5’
and adjacent to an AP site in addition to 3’ phosphoesterase
activity which is utilised to remove the 3’-blocking damage
as a result of ROS. Research has shown that APE1 is rate
limiting in the repair of 3’-blocking damage [25]. The
accumulation of AP sites causes increased mutagenic
potential and can result in DNA strand breaks and apoptosis
[26]. Interestingly NEIL1 does not require APE1 as it can
perform -elimination of the AP site, it may also act as a
backup for APE1 in the repair of 8-oxoG with OGG1 [24]. It
is outside the scope of the current review to examine the
entire BER pathway but there are a large number of enzymes
that facilitate the repair of oxidative DNA damage [22] and
those with redox regulation will be discussed later.
It should be noted at this juncture that although the
Xeroderma pigmentosum C (XPC) protein is widely
recognised as a component of the NER pathway there is also
some degree of cooperativity with BER. XPC has been
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Storr et al.
Prdx Family
GPx Family
Catalase
SOD
O2•-
H2O2
H2O + O2
Fenton reaction
•OH
OGG1
Repair
Fig. (1). Hydrogen peroxide can result in production of the hydroxyl radical via the fenton reaction. The hydroxyl radical can cause oxidative
DNA damage, one of the most common lesions is 8-oxoG. DNA can be repaired through the removal of 8-oxoG by the BER pathway
initiated by DNA glycosylases, in this example OGG1.
shown to act as a cofactor for the efficient cleavage of 8oxoG by OGG1 [27].
THE ROLE OF ANTIOXIDANT PROTEINS IN DNA
DAMAGE
The production of the hydroxyl radical and ONOO- and
therefore the prevention of DNA damage, are effectively
maintained in normal cells by the control of H2O2. This
control is managed by a number of redox enzymes including
the peroxiredoxins (Prdx). Prdx are ubiquitously expressed
enzymes that regulate whether H2O2 acts as a signal inducer
or becomes a harmful oxidant. The enzymes catalyse the
reaction by oxidation of the peroxidatic cysteine which can
then form a disulphide bond with the resolving cysteine
residue; this is mainly reduced by thioredoxin (Fig. 2).
During those times when H2O2 is acting as a signalling
mediator, the peroxidatic cysteine can become over oxidised
to sulfinic acid and reduced back to thiol by a sulfinic acid
reductase, sulfiredoxin (Srx) in an ATP dependent
mechanism [28, 29]. However over oxidation to sulfonic
acid is also a common occurrence which is irreversible and
leads to protein degradation [30] or to the formation of
oligomeric peroxidase-inactive chaperones [31]. In addition
to overoxidation, Prdx activity can also be regulated by
phosphorylation and proteolysis [32, 33].
Abnormal expression of Prdx has been found in many
kinds of cancers [34-36] and further induction of oxidative
stress such as that caused by ionizing radiation can induce
their expression [37]. Mice lacking Prdx1 show decreased
lifespan and increased tumour incidence and an increased
level of 8-oxo-2’deoxyguanosine (8-oxoG) formation [38].
Similar observations have been made for Prdx5 whereby
siRNA mediated knockdown results in increased 8-oxoG
formation [39]. The aberrant expression of the Prdx family is
reviewed by Zhang et al. (2009) [37].
The thioredoxin system has been shown to be both over
and under expressed in a number of different cancers [4044]. The activity of thioredoxin can be inhibited by
thioredoxin interacting protein (TxNIP), and recycled by the
selenoenzyme thioredoxin reductase. Thioredoxin has been
implicated in a number of cellular processes including
maintaining a reducing cellular environment (through
interaction with downstream Prdx), regulating cell growth
(via ribonucleotide reductase and promotion of growth factor
expression), transcription factor activation (including NFkB, p53, Hypoxia inducing factor HIF-1) and also
apoptosis (ASK-1) [45-47]. Thioredoxin translocates to the
nucleus in response to oxidative stress induced by hydrogen
peroxide, exposure to UV, ionising radiation and platinum
based treatments [48, 49]. Debatably one of the most
important functions of thioredoxin within the nucleus,
excluding its action in controlling hydrogen peroxide levels,
is its role in regulating DNA repair through interaction with
factors such as p53, which will be discussed later in terms of
APE1 redox regulation.
In addition to Prdx, both the glutathione peroxidase
(GPx) family and catalase function to control H2O2 (Fig. 3).
The GPx family consists of 8 members, 5 of which are
selenoproteins, GPx -1, 2, 4, 5 and 6, and respond via
different mechanisms to selenium deficiency. The family
members have been implicated, to varying degrees, in
carcinogenesis, with selenium supplementation indicated in
some reports [50]. Studies have shown that GPx1 null mice
that are heterozygous for manganese superoxide dismutase
(MnSOD) show increased oxidative damage, through
detection of 8-oxoG, and increased incidence of tumours
Base Excision Repair, the Redox Environment
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
O2•-
H2O2
H2O + O2
SOD
Thioredoxin
interacting
protein (TxNIP)
91
Prdx Family
Thioredoxin
Thioredoxin s-s
Thioredoxin reductase
or
Thiol oxidation of target protein
Fig. (2). The peroxiredoxin (Prdx) family involvement in the control of H2O2 levels within the cell. The thioredoxin system plays an
important role in regulating its turnover.
O2•-
H2O2
H2O + O2
SOD
GPx Family
GSH
GSSG
Glutathione reductase
NADPH
NADP+
Glucose 6-phosphate dehydrogenase
Fig. (3). The glutathione redox system and its role in the control of H2O2 levels within the cell by oxidising the tripeptide glutathione (GSH)
to glutathione disulphide (GSSG).
[51]. Expressing SOD 1 or 2 in retinal pigmented epithelial
cells, causes greater oxidative damage, which can be
nullified by increased expression of GPx4, and GPx1 to a
lesser extent [52]. Glutathione itself also acts as a scavenger
molecule against ROS.
Current research shows that it is possible to use routine
immunohistochemistry of the redox proteins regulating H2O2
and markers of oxidative DNA damage to predict clinical
outcome in some cancers [44, 53-57]. The current authors
have also demonstrated that members of the thioredoxin and
glutathione families can predict response to ROS generating
therapies in breast and ovarian cancer [58-61].
DIRECT MODULATION OF DNA REPAIR BY THE
REDOX ENVIRONMENT
An interesting aspect of the redox environment is its
ability to directly affect DNA repair, in addition to causing
DNA damage. There are limited examples of this modulation
of activity, with a number of them associated with the BER
pathway. The direct effect of ROS and the redox
environment on the short patch BER pathway is summarised
in Fig. (4).
As mentioned previously one of the most frequent, and
most studied, base lesions caused by oxidative damage is 8oxoG. One of the first observations that redox status had
effects on damage repair was conducted using HeLa cervical
cancer cells in that cadmium increased the appearance of the
ROS-mediated 8-oxoG by the creation of an oxidising
environment, but importantly the activity of DNA repair was
also impaired leading to increased carcinogenic potential
[62]. 8-oxoG is eliminated through the BER pathway
through excision by OGG1 and cadmium has since been
shown to inactivate OGG1 through cysteine modification
[63]. The activity of OGG1 can be directly modulated by the
redox environment where a polymorphism switching serine
to cysteine at position 326 of the protein is detrimental to
DNA repair. Significantly the Ser326Cys polymorphism is
92 Current Molecular Pharmacology, 2012, Vol. 5, No. 1
Storr et al.
ROS Modulation
Bifunctional glycosylase
Decreased activity (with
Ser326Cys mutation)
OGG1
Increased activity through
interaction with YB-1
NEILs
NEILs
OGG1
OGG / NTH1
P
P
PUA
Redox control of
transcription factors
NEILs
PNK
APE1
APE1
OH
P
Short patch
Increased activity with
Pol β via thiol switch
XRCC1
P
Long patch
XRCC1
Pol β
LigIII and I
FEN-1
PCNA
PARP
Fig. (4). ROS modulation of the BER pathway on an exemplary 8-oxoG lesion as a result of oxidative damage by bifunctional DNA
glycosylases. OGG and NTH1 act via elimination with the NEILs acting via elimination. PNK: polynucleotide kinase; LigIII and I:
DNA ligase III and I; FEN-1: flap structure-specific endonuclease 1; PCNA: proliferating cell nuclear antigen. The NEIL route of repair may
function during active transcription and replication due to its increased affinity for DNA bubbles.
associated with an increased risk of developing cancer [64]
due to the altered repair capabilities in an oxidising
environment. Bravard and colleagues (2006, 2009) showed
that oxidation of the mutant cysteine 326 of OGG1 forms a
disulphide bond with other cysteines within the protein. It is
postulated that the reduction in enzymatic activity is a result
of the conformational change induced but can be reversed in
the presence of reducing agents such as DTT [63, 65].
Human ribosomal protein S3 plays a role in influencing the
recognition of 8-oxoG sites by blocking recognition by
OGG1. Interestingly 8-oxoG binding of hS3 can be
abrogated by a single amino acid change, which stimulates
the repair of 8-oxoG by OGG1 [66, 67]. hS3 has also been
shown to influence NF-kB mediated transcription [68] and
can interact with p53 and MDM2 with interactions
increasing upon exposure to oxidative stress [69].
OGG1 is not the only BER pathway member to be
directly modulated by the redox environment. APE1 is a
multifunctional protein that in addition to being an A/P
endonuclease has redox activity. Human APE1 was first
described and cloned as a DNA repair enzyme in 1991 [70,
71] and the redox activity of APE1 was described shortly
after in 1992 [72]. These two functional aspects of the
protein reside in different, distinct areas of the protein, with
the redox domain located in the N-terminal amino region
[72, 73]. The redox domain of APE1 can function to
modulate DNA damage in an indirect manor through its
interaction with transcription factors and is discussed later.
There is also evidence that the redox state of APE1 is
implicated directly in its DNA repair activity [74-76].
Immediately adjacent to the crucial histidine in the DNA
repair active site of APE1 is a cysteine that can be redox
regulated [74], potentially interfering with the function of the
active site. Ramana et al. (1998), [75] were able to
demonstrate that APE1 is activated by non-toxic levels of
ROS, but not UV light or alkylating agents, and promotes
translocation into the nucleus. The translocation of APE1
into the nucleus in response to ROS has been investigated
further. Extracellular ATP stimulates the purinergic receptors
(P2) through Ca2+ mobilisation and the production of ROS
and is responsible for the localisation of APE1 [77]. In
addition phosphorylation by protein kinase C (PKC)
following an oxidative challenge has been shown to increase
the activity of the APE1 redox domain [78].
NEILs (Nie-like-1 and 2) are other DNA glycosylases
involved in the BER pathway. They function by a different
mechanism to OGG1 and NTH1 (homologue of E. coli
endonuclease III) but all can act on duplex DNA. NEILs,
however, preferentially act upon DNA bubbles or single
stranded DNA, implying the activity of these glycosylases
during active transcription and replication [79]. Oxidative
stress causes the translocation of the Y-box binding protein-1
(YB-1) to the nucleus and its stable interaction with NEIL2,
increasing NEIL2 activity in the BER pathway. The
interaction between NEIL2 and YB-1 is physical and
increases the excision activity of NEIL2 seven fold [80].
In addition to OGG1 and APE1, the interaction between
XRCC1 (X-ray repair, cross-complementing defective, in
Chinese hamster, 1) and DNA polymerase (Pol ) binding
Base Excision Repair, the Redox Environment
is influenced by the redox environment. Recently, Cuneo and
London (2010), [81] examined the crystal structure of the
oxidised and reduced N-terminal domain of XRCC1 in
complex with Pol and showed that oxidised XRCC1
demonstrated altered folding topology through the formation
of a disulphide bond. Most of the structural changes occurred
in an area of the protein that was not directly responsible for
Pol binding but the oxidative changes were able to enhance
affinity. The authors offer an interesting hypothesis whereby
APE1, a known binding partner of XRCC1, may play a role
in the activation or deactivation of XRCC1s disulfide switch
through its redox domain [81].
It is not only the BER pathway that can be modulated
directly by oxidative stress. As the role of therapeutic
ionising radiation is to induce oxidative stress and create
double strand breaks within the cell an element of redox
control in the double strand DNA repair pathway may seem
paradoxical. However the redox environment seems to play
an important role within this pathway. In the nonhomologous end joining (NHEJ) double strand DNA break
repair pathway Ku is responsible for binding DNA. An
oxidising environment results in lower DNA binding of the
protein which is reversible upon reduction. It is unclear how
oxidisation affects the ability of the protein to bind DNA as
disulphide bonds are not observed but it is possible that
cysteine sulfenic acids are formed [82]. The influence
exerted by the redox environment functions to increase or
decrease the time Ku is bound to the DNA which has an
impact upon the likelihood of recruitment of the DNA-PK
catalytic subunit (DNA-PKcs) to form the DNA-PK complex
[83]. A further redox effect within this process is glucose-6phosphate dehydrogenase (G6PD). G6PD is important in the
redox pathway for its role in the oxidative pentose phosphate
cycle regulating the NADPH/NADP+ ratio. Genetic defects
in G6PD are relatively common, and can lead to an increased
susceptibility to oxidative stress. Ayene and colleagues
(2002), showed that in G6PD null mutant Chinese hamster
ovary cells that Ku binding is inactivated during induced
oxidative stress [84].
In another aspect of the double strand DNA pathway,
ataxia-telangiectasia mutated (ATM) protein kinase is
activated following DNA damage to sense double strand
breaks which starts a signalling cascade. Recent research has
identified a role of ROS regulation of ATM, whereby
elevated ROS activate ATM, to activate the tumour sclerosis
complex 2 (TSC2) tumour suppressor in the cytoplasm to
repress mTORC1 and induce autophagy [85]. This shows an
interesting influence exerted by a redox environment on a
DNA damage sensing protein pushing the system to
autophagy in response to ROS.
Interestingly the redox regulation of a number of other
DNA binding proteins has also been described. Human
replication protein A (RPA) is a DNA binding protein
implicated in DNA replication, repair and recombination.
Analysis of RPA using mass spectrometry reveals that in
oxidative conditions the cysteines in the zinc-finger motif of
the p70 subunit can form disulphide bonds that impair DNA
binding [86]. Although these latter examples are proteins not
involved in the DNA BER pathway it is interesting to note
that oxidative stress plays a role in modulating a number of
other DNA repair pathways.
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
93
INDIRECT EFFECT OF THE REDOX ENVIRONMENT ON DNA REPAIR
In addition to the direct modulation mentioned above the
redox environment can also influence DNA repair indirectly
through other interactions. For example, the expression of
NEIL1 is increased by ROS through the activation of
CREB/c-Jun transcription factors [87].
APE1 not only acts directly in BER but also has its own
distinct redox domain. This redox domain is itself able to
influence DNA repair, primarily through the binding to
various transcription factors such as AP-1, HIF-1 and NFB
[72, 88, 89]. In the example of HIF-1 activation, expression
of both APE1 and the dithiol reducing enzyme thioredoxin
potentiate its activation by redox dependent stabilisation of
the HIF-1 alpha subunit [88]. Interestingly HIF-1 is able to
attenuate APE1 expression in endothelial cells [90]. In
addition to the indirect effect upon DNA repair APE1 is also
implicated in angiogenesis following oxidative damage.
APE1 and HIF-1 are implicated in the formation of the
transcriptional complex of the hypoxic response element
(HRE) of the VEGF gene, which is increased through
selective modification of nucleotides within the HRE by
oxidative damage; importantly these hypoxia induced base
modifications are associated with transcriptionally active
nucleosomes [91-93].
APE1 and thioredoxin can act independently or in
concert on various transcription factors in an interaction
influenced by ROS [94-96]. Redox dependent transcriptional
activation is reviewed by Liu et al. (2005) [97]. APE1 is
thought to act in a redox cycle with thioredoxin whereby
APE1 is able to maintain transcription factors in their
reduced state, and the redox state of APE1 is maintained by
thioredoxin, which itself can have a direct action on
transcription. The redox regulation of APE1 by thioredoxin
is required for the activation of p53 and AP-1 [94, 95]. p53
has been well characterised in respect to it’s acting as a
gatekeeper for DNA damage, inducing G1 arrest, to provide
time for, and inducing enzymes involved in, DNA repair [98,
99]. An interesting hypothesis has been proposed by
Seemann and Hainut (2005) whereby thioredoxin, p53, and
APE1 function as an important switch in the BER pathway
[100]. In a reduced environment, where basal levels of DNA
damage are observed or when thioredoxin recycling is high,
APE1 functions principally in the BER pathway. This allows
p53 to stimulate the actions of the glycosylases [101] and
stabilise interactions between Pol and abasic DNA [102]. In
a highly oxidising environment, where excessive DNA
damage occurs, APE1 is thought to function through p53 to
suppress growth or initiate apoptosis. p53 is subject to redox
modulation by reduction of cysteine 277 in its C-terminal
part of its DNA binding domain [103]. Thioredoxin itself can
also directly enhance the specific DNA binding of p53 [95].
The anti-oxidant functions of p53 are reviewed by
Olovnikov et al. (2009) [104].
Further evidence for the importance of the redox
environment in DNA repair is highlighted in current research
which demonstrates that BRCA1, a breast cancer
susceptibility gene, plays an important role in regulating the
BER pathway. BRCA1 encodes a tumour suppressor protein
94 Current Molecular Pharmacology, 2012, Vol. 5, No. 1
and mutations within the gene account for 40-50% of
hereditary breast cancer. Saha and colleagues (2010),
demonstrated that challenging T47D breast cancer cells with
H2O2 is able to cause an increase in BRCA1 expression and
three BER enzymes; OGG1, NTH1, and APE1[105]. The
authors investigated the importance of BRCA1 in
influencing the BER pathway and suggest that it acts as a coregulator of the octomer-binding transcription factor OCT1.
Expression of BRCA1 is increased in response to H2O2 and
can increase 8-oxoG excision by stimulating the expression
of OGG1, NTH1 and APE1, and the incising ability of
NTH1 [105].
The importance of the oxidative environment on poly
(ADP-ribose) polymerase 1 (PARP-1) is also becoming
apparent. PARP-1 is a nuclear enzyme involved in DNA
repair and cell death. PARP-1 participates in the BER
pathway through its interaction with XRCC1 [106]. It is
believed that XRCC1 is recruited through the addition of
poly (ADP-ribose) to PARP, which at the same time reduces
the affinity of PARP for DNA via the large addition of
negative charge [107]. The activity of PARP-1 is increased
following ROS challenge, through the increase in oxidative
DNA damage that signal activation of the enzyme [108]. In
addition, PARP-1 has been shown to play a role in chromatin
repair. Histones, the main protein in chromatin structure, act
as a defence against oxidative DNA damage as they
themselves can be oxidised and the level of oxidation
determines their fate. Histones that have undergone oxidative
damage can cross link with DNA impairing transcription and
replication. The PARP-1 mediated poly (ADP-ribosyl)ation
of undamaged histones, in addition to the (ADP-ribosyl)ation
activation of the proteasome, mediates the degradation of
oxidatively damaged histones [109]. H2O2 stimulates the
activity of PARP-1 through DNA damage and the
mechanism of apoptosis changes with the stimulation
intensity. A continuous level of H2O2, rather than a H2O2
bolus, results in different mechanisms of cell death. A H2O2
bolus results in caspase-dependent apoptosis, whereas
continuous stimulus results in caspase-independent apoptosis
through apoptosis independent factor (AIF) [110, 111]. In
fact, arsenic trioxide (As2O3), approved for treatment of
acute promyelocytic leukaemia (APL) in patients who have
relapsed or are refractory to first line intervention using
retinoid and anthracycline chemotherapy, acts in a
prooxidative mechanism and can influence PARP-1
mediated apoptosis. As2O3 acts to induce the mitochondrial
pathway of apoptosis and appears to inactivate a number of
anti-oxidant enzymes [112]. In solid cancers, such as ovarian
cancer, As2O3 mediates PARP-1 activation to induce AIF
release from mitochondria initiating caspase-independent
cell death [113].
There is increasing evidence for a direct role of the redox
environment in PARP-1 activation in a mechanism that
initially seems counter intuitive. PARP-1 can be inactivated
by the oxidation of thiols within its DNA binding zinc finger
motif causing expulsion of the zinc ion. Both ONOO and
N2O3, formed through superoxide reaction with NO, are
capable of inactivating PARP through oxidation or
nitrosylation [114]. Further work demonstrated that
nitrosylation of PARP-1 impairs the ability to bind to the
inducible nitric oxide synthase (iNOS) promoter by negative
Storr et al.
feedback regulation [115]. This proves an interesting
finding, as the oxidative environment can cause inhibition of
the DNA binding component of the enzyme, however its
activity is activated by oxidative DNA damage.
THE REDOX PARADIGM
ROS induced DNA damage can result in induction of
signal transduction pathways, induction or arrest of
transcription, replication errors and therefore genomic
instability, all of which are associated with carcinogenesis
[116]. It may be logical to assume that, in the cancer setting,
reduction of ROS would be beneficial. However, the
majority of non-surgical therapies for cancer utilise
treatments that function through the production of ROS such
as radiotherapy, photodynamic therapy and certain
chemotherapy agents. The logic for this strategy is that
cancer cells are under increased intrinsic ROS stress and
therefore further insult from ROS generating therapies could
exhaust the enhanced antioxidant capacity of the cancer cells
and lead to apoptosis. Therefore the question still remains ‘is
it beneficial or detrimental to reduce ROS?’
Redox buffering systems in cells such as the thioredoxin
and glutathione systems and antioxidant enzymes, such as
catalase, superoxide dismutases and the peroxidases, are
often deregulated in cancer cells and can interfere with the
effectiveness of ROS generated by radiotherapy [116-119].
Ionising radiation acts directly via the production of ROS
from intracellular H2O to hydrogen peroxide causing
oxidative DNA and protein damage and indirectly through
cellular signalling. A number of studies have shown, in vitro,
that modulation of redox homeostasis can alter the response
of cancer cells to low LET radiations such as X-rays and rays i.e. those used in conventional radiotherapy [117, 120122].
Certain chemotherapeutic agents can also produce ROS
through a number of mechanisms, including superoxide
generation by anthracyclines through the redox quinone
cycle, and nucleophilic substitution reactions by platinum
complexes. In 1988, Kramer and colleagues demonstrated
that the glutathione redox cycle played a role in
chemotherapy resistance [123]. A more reducing cellular
environment can result in chemotherapy resistance, in part
through actions on the multidrug resistance (MDR)
transporter P-glycoprotein (P-gp) [123, 124]. This indicates
the potential therapeutic significance of increasing cellular
ROS levels. It should also be borne in mind however that
certain antioxidants, for example thioredoxin reductase [125]
and glutathione [126, 127] can become prooxidants at high
levels. Thioredoxin has been shown to increase the redox
cycling of daunomycin, enhancing apoptosis, demonstrating
both novel prooxidant and proapoptotic roles [128].
Significantly, ROS inhibit the function of protein tyrosine
phosphatase (PTP) which allows increased growth factor
signalling resulting in proliferation, and increases growth
factors and matrix metalloproteins (MMP) involved in
angiogenesis. However ROS also act to initiate receptor- and
mitochondria- mediated apoptosis. The Fas ligand (FL) can
cause an increase in ROS which can result in receptormediated apoptosis via Fas receptor binding and signalling
through the death receptor pathway; and during
Base Excision Repair, the Redox Environment
mitochondria-mediated apoptosis ROS modulate the
permeability of the transition pore complex to influence the
process. The ROS paradigm has been reviewed previously
[129].
As part of the debate of the benefits of reducing or
increasing ROS in cancer, dietary antioxidant supplements
have often been thought to reduce the risk of cancer. Studies
have shown some efficacy and the effect on cellular DNA
damage and BER activity has been assessed. Caple and
colleagues (2009) demonstrated that DNA damage following
H2O2 challenge and BER levels differ amongst healthy
individuals and that supplementation with an antioxidant
supplement containing selenium and vitamins A, C and E, in
the group with highest level of DNA damage could mediate
a protective effect [130]. However, there is increasing
evidence showing that there may be a link between dietary
antioxidants and the risk of cancer. For example recent
studies have shown that intake of carotene and carotene
infers an increased risk of ER and PR negative breast cancer
in smokers [131] The action of vitamins however is not
always antioxidant, the hormonal form of vitamin D
(1,25(OH)2D,3), acts as a pro-oxidant which is able to
reverse treatment resistance [132]. The therapeutic advantage
of increasing ROS through various mechanisms including
inhibition of anti-oxidant enzymes in the treatment of cancer,
either as a stand alone agent or in combination with other
therapies such as chemotherapy and radiotherapy has been
investigated and the main targets discussed in the next
section.
The differences in the redox environment between
normal and malignant tissues and within malignant tissues
themselves i.e. hypoxic regions, can alter the response of a
tumour to treatment modalities that utilise changes in
oxidative stress as a main or by-product of their action.
Therefore techniques to non-invasively distinguish these
differences between normal and malignant cells could have
clinical implications and imaging techniques are currently
being developed. Hyodo et al. (2008) review the work
conducted so far in this area and demonstrate that cellpermeable nitroxides, coupled with magnetic resonance
imaging (MRI) can non-invasively examine the differences
between the redox status of tissues [133]. Paramagnetic
nitroxide radicals undergo reduction to the corresponding
diamagnetic hydroxylamine which can revert to nitroxide in
the presence of oxidants or if the cellular oxygen status
permits [134]. Hypoxic conditions, tissue redox status and
oxidative stress will enhance the conversion of the
paramagnetic nitroxide radicals to its corresponding
diamagnetic products distinguishing tumour from normal
tissue [135].
THE REDOX SYSTEM AS A THERAPEUTIC TARGET
The redox system is becoming an increasingly interesting
target for cancer therapies, but as indicated above questions
remain as to whether increasing or decreasing ROS is
beneficial and the most appropriate therapeutic option. By
reducing ROS, oxidative DNA damage is reduced. By
increasing ROS, mutations through oxidative damage are
more likely to occur, however the cell is more likely to
initiate an apoptotic pathway.
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
95
Various small molecule inhibitors are available for the
inhibition of anti-oxidant enzymes involved in the control of
cellular H2O2 levels and it is thought that these inhibitors
may be of therapeutic benefit when given in combination
with traditional cancer therapies which act through the
generation of ROS. There are a large number of proteins that
could be targeted by this mechanism to aid in ROS
production to push the cell to an apoptotic response. This
also however, makes this strategy more challenging due to
the redundancy of these pathways. In addition to anti-oxidant
enzyme modulation, ROS can be generated directly within
the cell, such as with arsenicals, organic endoperoxidases
and redox cyclers such as motexafin gadolinium. One of the
first redox modulating drugs, procarbazine, works by the
production of hydrogen peroxidase via oxidation and is used
in the treatment of Hodgkins lymphoma [119]. Motexafin
gadolinium (Xcytrin), utilizes thioredoxin reductase with
NADPH, and other cellular metabolites, increasing oxidative
stress in a process known as futile redox cycling [136, 137]
and has successfully completed a phase III international
study [138]. Arsenic trioxide (As2O3) is able to inhibit GPx,
resulting in an increase in H2O2, as well as affecting the
mitochondrial respiratory chain thus causing generation of
superoxide [139]. However the actions of arsenic compounds
have been implicated in a number of ROS generating
pathways [140]. Low concentrations of arsenic trioxide have
been shown to induce a high rate of clinical remission in
acute promyelocytic leukemia (APL) patients [141, 142].
Redox directed cancer therapeutics are comprehensively
reviewed by Wondrak (2009) [112]. Pennington et al. (2005)
outlines a set of criteria for consideration when selecting a
redox target for therapeutic gain and reviews some of the
proteins that fulfil some or all of them: 1) be over-expressed
or be constitutively active in tumour cells, 2) enhance
tumour proliferation, 3) exhibit pro-survival response, 4)
enhance resistance to therapeutic modalities i.e. radiotherapy
and chemotherapy [118]. The main redox systems already
discussed in the context of DNA damage and repair, that
have been targeted therapeutically are outlined below.
Although impressive in vitro and in vivo effectiveness is
often shown translation into clinically efficacy has often
been disappointing, for this reason emphasis is given to those
agents that have reached clinical evaluation.
The Superoxide Dismutase System
The SOD system can be targeted using two methods; the
inhibitors, TETA, ATN-224 and 2-methoxyestradiol and the
mimetics,
M40403,
mangafodipir,
cis-FeMPy2p2p,
MnTBAP and TEMPO. The SOD inhibitors act as copper
chelators (TETA and ATN-224) or oestrogen derivatives (2methoxyestradiol) to inhibit the dismutation of superoxide to
H2O2, causing an increase in superoxide that can act to
induce apoptosis. The SOD mimics can be nitroxide free
radicals or metal based agents, such as chelates of
manganese (II). The mimics act principally through the
turnover of superoxide, and have important clinical effects in
conditions other than cancer, such as cardioprotective
intervention or to limit radiation induced side effects [143]
and M40403 and mangafodipir have been trialled as
palliative management agents [112]. The inhibitors ATN-
96 Current Molecular Pharmacology, 2012, Vol. 5, No. 1
224 and 2-methoxyestradiol have been the focus of phase II
clinical trials for advanced melanoma, prostate cancer,
multiple myeloma, recurrent or advanced breast cancer and
in numerous cancer types respectively [112]. Results from
the phase II trial of 2-methoxyestradiol in taxane-refractory
castrate-resistant prostate cancer reveal the study was
terminated following futility analysis [144]. In another study
of hormone-refractory prostate cancer 2-methoxyestradiol
showed a dose response effects on PSA velocity [145].
Storr et al.
downstream targets of thioredoxin. There are however, a
number of agents that can act upon thioredoxin reductase-1
in a non-specific manner such as motexafin gadolinium as
mentioned previously. Chaetocin and gliotoxin are also able
to inhibit thioredoxin reductase-1 by acting as competitive
substrates for the enzyme however no clinical trials, to our
knowledge, are currently underway with these agents. There
are a number of other agents that have effects on the
thioredoxin system, although often not specific inhibitors;
these agents are discussed by Gromer et al. (2004) [47].
The Glutathione System
The glutathione (GSH) system can be targeted by the
modulation of the pathway using agents including NOV-002,
Imexon, L-buthionine R sulfoximine (BSO), and PABA/NO.
NOV-002 is a combination of glutathione disulphide
(GSSG) and cis-platinum and acts by causing a decreased
GSH:GSSG ratio and accumulation of H2O2. Imexon, a thiol
reactive electrophile, creates spontaneous thiol adducts with
glutathione amongst other actions. BSO depletes glutathione
and induces oxidative stress. PABA/NO acts as a
glutathione-S-transferase prodrug that is metabolised to
nitric oxide. These agents all act to increase oxidative stress
by impairing H2O2 turnover. The inhibitor NOV-002 has
been the focus of a number of clinical trials for various
cancer types including current phase III trials in lung cancer
in combination with chemotherapy, and the safety of Imexon
is being evaluated. BSO is currently in phase I clinical trials
in neuroblastoma and melanoma [112].
The Peroxiredoxin (Prdx) Family
There are fewer examples of inhibitors or modulators of
enzymatic activity of the Prdx family, the antioxidant
enzymes involved in hydrogen peroxide dismutation. An
example of an agent that has inhibitory effects on Prdx is
conoidin A, able to inhibit Prdx1 and Prdx2 [146];
Ladostigil, an inhibitor of cholinesterase monoamine oxidase
inhibitor, has been shown to increase the expression of Prdx1
amongst other anti-oxidant enzymes [147]. The impact of
Prdx inhibition has been demonstrated in-vitro by Wang and
colleagues (2005) [148]. Their experiments have indicated
that specific siRNA knockdown of Prdx1 can cause
radiosensitisation in MCF7 breast cancer cells, indicating the
possible therapeutic impact of controlling the redox response
to radiotherapy. The Prdx are reviewed, in relation to
radiotherapy, by Zhang and colleagues (2009) [37].
The Thioredoxin System
The thioredoxin system is also a target for inhibition,
through the actions of drugs such as PX-12 (IV-2), PX-916,
PMX464 and PMX290. PX-12 and PMX464 inactivate
thioredoxin through disulphide exchange of cysteine 73 and
oxidative formation of a disulphide bridge between cysteine
32 and 35 respectively [149-151]. PX-12 has been examined
in clinical trials in advanced metastatic cancers [112],
however recent results suggest no anti-tumour effect as a
single agent in advanced pancreatic cancer previously treated
with gemcitabine [152]. As with PMX464, PMX290
(AJM290) also serves as a specific thioredoxin inhibitor.
PX-916 is able to inhibit thioredoxin reductase-1 agonising
Catalase
Catalase is one of the enzymes responsible for the control
of cellular hydrogen peroxide. In 1941, Greenstein et al.
reported that liver catalase activity was reduced in rats with
subcutaneously implanted hepatic tumours [153]. 3-amino1:2:4-triazole is able to inhibit catalase irreversibly in the
presence of a continuous supply of hydrogen peroxide [154].
The inhibitor has been shown to promote thyroid tumour
formation in rats when given in combination with carcinogen
[155]. As with some of the redox proteins mentioned above
we are unaware of any clinical trials examining catalase
inhibitory effects. Catalase mimics are reviewed by Day
(2009) [156].
Altering DNA Repair Through the Redox Environment
Due to the redox domain of APE1 this enzyme is the
current target of two approaches to modulate its activity for
cancer therapy. The first is the inhibition of its DNA repair
facility and the second is to inhibit its redox ability. Both
approaches are the subject of other articles in the current
publication. The inhibitors for APE1 DNA repair activity
have been previously reviewed by Fishel and Kelley (2007)
[157]. Examples of inhibitors of the redox domain of APE1
include E3330 and analogues benzoquinone and
napthoquinone which are able to inhibit the redox function
of APE-1 and the growth of an ovarian cancer cell line [158].
MSH2 is a gene that encodes components of the DNA
mismatch repair pathway. Treatment with methotrexate
causes the accumulation of 8-oxoG in cells lacking
functional MSH2. MSH2 is a relatively common mutation in
hereditary non-polyposis colon cancer (HNPCC). Martin and
colleagues (2009) demonstrated that although 8-oxoG
accumulated in both MSH2 deficient and proficient cells,
accumulation only occurred in deficient cells with rapid
clearance in proficient cells. The group investigated this
interaction further by specific knockdown of dihydrofolate
reductase and demonstrated that methotrexate seemed to
modulate folate synthesis via inhibition of dihydrofolate
reductase to explain MSH2 involvement [159].
Antioxidant Supplement
Due to the importance of redox homeostasis and the
implications of oxidative DNA damage the dietary
supplementation by antioxidants is of interest. To date the
information surrounding dietary supplementation and cancer
risk has shown inconclusive results. Antioxidant
supplementation and cancer risk is reviewed by Loft et al.
(2008) [160].
Base Excision Repair, the Redox Environment
CONCLUSIONS AND FUTURE DIRECTIONS
Redox homeostasis is critical for the function of the cell.
ROS play important cellular roles in mediating signal
pathways; however aberration of such normal pathways can
lead to oxidative DNA damage. Oxidative DNA damage is
repaired principally through the BER pathway allowing
excision of lesions such as 8-oxoG. The accumulation of
oxidative DNA lesions can result in genomic instability, and
increased risk of cancer. In the cancer setting, cells have
aberrant ROS production; this increased production of ROS
can lead to further DNA mutations, in addition to increased
cellular signalling. Perhaps surprisingly the generation of
ROS is the mechanism of action of most cancer therapeutics
such as radiotherapy and chemotherapy, and the current
interest is to try and circumvent resistance to these therapies.
In a healthy cell there are a number of anti-oxidant enzymes
that serve to modulate the redox environment by controlling
the production of superoxide and hydrogen peroxide to
mitigate the production of the damaging hydroxyl radical.
These enzyme pathways are altered during carcinogenesis,
and current interest is to modulate their actions to allow the
accumulation of ROS in the case of therapy resistance, or
reducing ROS to manage treatment toxicity. There are a
number of agents that inhibit enzymes involved in redox
homeostasis, or involve the direct modulation of the redox
state of the cell.
The development of inhibitors of the BER pathway is an
area of intense focus with the aim of mirroring the successes
observed with the current generation of PARP inhibitors,
especially in BRCA related disease. It is hoped that a similar
strategy may be possible for enzymes such as APE1 [161].
The effects of altered BER pathways in patients remain to be
elucidated, such as the accumulation of AP sites in terms of
APE1 inhibitors. However, due to the potential mutagenic
effects of modulating these pathways secondary cancers may
become apparent. Interestingly, many of the enzymes
involved in the BER pathway are subject to some degree of
regulation by ROS, often activated in the case of elevated
stress levels. This may suggest that ROS generating
therapies, in addition to anti-oxidant enzyme modulators and
inhibitors of the BER pathway may be of interest.
Further research is required to understand the full impact
of redox homeostasis on the cell, its involvement in
modulating the BER response in addition to causing the
initial oxidative DNA damage.
ABREVIATIONS
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
97
EMT
=
Epithelial-mesenchymal transition
FL
=
Fas ligand
G6PD
=
Glucose-6-phosphate dehydrogenase
GPx
=
Glutathione peroxidase
GSH
=
Glutathione
GSSG
=
Glutathione disulphide
HNPCC
=
Hereditary
cancer
HRE
=
Hypoxic response element
non-polyposis
iNOS
=
Inducible nitric oxide synthase
LP
=
Long patch
MDR
=
Multidrug resistance
MMP
=
Matrix metalloproteins
colon
MnSOD
=
Manganese superoxide dismutase
MRI
=
Magnetic resonance imaging
NEILs
=
Nie-like-1 and 2
NER
=
Nucleotide excision repair
NF-B
=
Nuclear factor-B
NHEJ
=
Non-homologous end joining
Nox
=
NADPH oxidase
OGG1 and OGG2 =
8-oxoG DNA glycosylases
8-oxoG
=
8-oxo-2’deoxyguanosine
P2
=
Purinergic receptors
PARP-1
=
Poly (ADP-ribose) polymerase 1
P-gp
=
MDR transporter P-glycoprotein
PKC
=
Protein kinase C
Prdx
=
Peroxiredoxins
PTP
=
Protein tyrosine phosphatase
ROS
=
Reactive oxygen species
RPA
=
Replication protein A (human),
SOD
=
Superoxide dismutases
SP
=
Short patch
Srx
=
Sulfiredoxin
TSC2
=
Tumour sclerosis complex 2
TxNIP
=
Thioredoxin interacting protein
XPC
=
Xeroderma pigmentosum C
XRCC1
=
X-ray repair, cross-complementing
defective, in Chinese hamster, 1
AIF
=
Apoptosis independent factor
AP
=
Apurinic/apyrimidinic
AP-1
=
Activator protein 1
APE1
=
AP endonuclease-1 (human)
[1]
APL
=
Acute promyelocytic leukaemia
[2]
As2O3
=
Arsenic trioxide
ATM
=
Ataxia-telangiectasia mutated
[3]
BER
=
Base excision repair
[4]
BSO
=
L-buthionine R sulfoximine
REFERENCES
Dröge, W. Free radicals in the physiological control of cell
function. Physiol. Rev., 2002, 82, 47-95.
D’Errico, M.; Parlanti, E.; Dogliotti, E. Mechanism of oxidative
DNA damage repair and relevance to human pathology. Mutat.
Res., 2008, 659, 4-14.
Wu, W-S. The signalling mechanism of ROS in tumour
progression. Cancer Metastasis Rev., 2006, 25, 695-705.
Valko, M.; Rhodes, C.J.; Moncol, J.; Izakovic, M.; Mazur, M. Free
radicals, metals and antioxidants in oxidative stress-induced cancer.
Chem. Biol. Interact., 2006, 160, 1-40.
98 Current Molecular Pharmacology, 2012, Vol. 5, No. 1
[5]
[6]
[7]
[8]
[9]
[10]
[11]
[12]
[13]
[14]
[15]
[16]
[17]
[18]
[19]
[20]
[21]
[22]
[23]
[24]
[25]
[26]
[27]
Karihtala, P.; Soini, Y. Reactive oxygen species and antioxidant
mechanisms in human tissues and their relationship to
malignancies. APMIS, 2007, 115, 81-103.
Pastor, N.; Weinstein, H.; Jamison, E.; Brenowitz, M. A detailed
interpretation of OH radical footprints in a TBP-DNA complex
reveals the role of dynamics in the mechanism of sequence specific
binding. J. Mol. Biol., 2000, 304, 55-68.
Beckman, J.A.; Beckman, T.W.; Chen, J.; Marshall, P.A. Apparent
hydroxyl radical production by peroxynitrite: implications for
endothelial injury from nitric oxide and superoxide. Proc. Natl.
Acad. Sci. USA, 1990, 87, 1620-1624.
Stamler, J.S.; Singel, D.J.; Loscalzo, J. Biochemistry of nitric oxide
and its redox-activated forms. Science, 1992, 258(5090), 18981902.
Ferrer-Sueta, G.; Radi, R. Chemical biology of peroxynitrite:
kinetics, diffusion, and radicals. ACS Chem. Biol., 2009, 4, 161-177
Burney, S.; Caulfield, J.L.; Niles, J.C.; Wishnok, J.S.;
Tannenbaum, S.R. The chemistry of DNA damage from nitric
oxide and peroxynitrite. Mutat. Res., 1999, 424, 37-49.
Jacob, C.; Knight, I.; Winyard, P.G. Aspects of the biological redox
chemistry of cysteine: from simple redox responses to sophisticated
signalling pathways. Biol. Chem., 2006, 387, 1385-1397.
Davies, K.J. The broad spectrum of responses to oxidants in
proliferating cells: a new paradigm for oxidative stress. IUBMB
Life, 1999, 48, 41-47.
Brown, N.S.; Bicknell, R. hypoxia and oxidative stress in breast
cancer. Oxidative stress: its effects on growth, metastatic potential
and response to therapy in breast cancer. Breast Cancer Res., 2001,
3, 323-327.
Behrend, L.; Henderson, G.; Zwacka, R.M. Reactive oxygen
species in oncogenic transformation. Biochem. Soc. Trans., 2003,
31, 1441-1444.
Totter, J.R. Spontaneous cancer and its possible relationship to
oxygen metabolism. Proc. Natl. Acad. Sci. USA, 1980, 77, 17631767.
Szatrowski, T.P.; Nathan, C.F. Production of large amounts of
hydrogen peroxide by human tumour cells. Cancer Res., 1991, 51,
794-798.
Cooke, M.S.; Evans, M.D.; Dizdaroglu, M.; Lunec, J. Oxidative
DNA damage: mechanisms, mutation, and disease. FASEB, 2003,
17, 1195-1214.
Wiseman, H.; Halliwell, B. Damage to DNA by reactive oxygen
and nitrogen species: role in inflammatory disease and progression
to cancer. Biochem. J., 1996, 313, 17-29.
Demple, B.; Harrison, L. Repair of oxidative damage to DNA:
enzymology and biology. Annu. Rev. Biochem., 1994, 63, 915-948.
Barzilai, A.; Yamamoto, K-I. DNA damage responses to oxidative
stress. DNA Repair, 2004, 3, 1109-1115.
Wilson III, D.M.; Bohr, V.A. The mechanisms of base excision
repair and its relationship to ageing and disease. DNA Repair
(Amst.), 2007, 6(4), 544-559.
Fan, J.; Wilson III, D.M. Protein-protein interactions and
posttranslational modifications in mammalian base excision repair.
Free Radic. Biol. Med., 2005, 38(9), 1121-1138.
Dizdaroglu, M.; Karahalil, B.; Sentürker, S.; Buckley, T.J.; RoldánArjona, T. Excision of products of oxidative DNA base damage by
human NTH1 protein. Biochemistry, 1999, 38, 243-246.
Mokkapati, S.K.; Wiederhold, L.; Hazra, T.K.; Mitra, S.
Stimulation of DNA glycosylase activity of OGG1 by NEIL1:
functional collaboration between two human DNA glycosylases.
Biochemistry, 2004, 43, 11596-11604.
Izumi, T.; Hazra, T.K.; Boldogh, I.; Tomkinson, A.E.; Park, M.S.;
Ikeda, S.; Mitra, S. Requirement for human AP endonuclease 1 for
repair of 3’- blocking damage at DNA single-strand breaks induced
by reactive oxygen species. Carcinogenesis, 2000, 21(7), 13291334.
Loeb, L.A.; Preston, B.D. Mutagenesis by apurinic/apyrimidinic
sites. Ann. Rev. Genet., 1986, 20, 201-230.
D’Errico, M.; Parlanti, E.; Teson, M.; Bernardes de Jesus, B.M.;
Degan, P.; Calcagnile, A.; Jaruga, P.; Bjørås, M.; Crescenzi, M.;
Pedrini, A.M.; Egly, J-M.; Zambruno, G.; Stefanini, M.;
Dizdaroglu, M.; Dogliotti, E. New functions of XPC in the
protection of human skin cells from oxidative damage. EMBO J.,
2006, 25, 4305-4315
Storr et al.
[28]
[29]
[30]
[31]
[32]
[33]
[34]
[35]
[36]
[37]
[38]
[39]
[40]
[41]
[42]
[43]
[44]
[45]
[46]
[47]
[48]
[49]
Wood, Z.A.; Poole, L.B.; Karplus, A. Peroxiredoxin evolution and
the regulation of hydrogen peroxide signalling. Science, 2003, 300,
650-653.
Biteau, B.; Labarre, J.; Toledano, M.B. ATP-dependent reduction
of cysteine-sulphinic acid by S.cerevisiae sulphiredoxin. Nature,
2003, 425, 980-984.
Neumann, C.A.; Fang, Q. Are peroxiredoxins tumor suppressors?
Curr. Opin. Pharmacol., 2007, 7, 375-380.
Neumann, C.A.; Cao, J.; Manevich, Y. Peroxiredoxin 1 and its role
in cell signaling. Cell Cycle, 2009, 8, 4072-4078.
Koo, K.H.; Lee, S.; Jeong, S.Y.; Kim, E.T.; Kim, K.; Song, K.;
Chae, H.Z. Regulation of thioredoxin peroxidase activity by Cterminal truncation. Arch. Biochem. Biophys., 2002, 397, 312-318.
Chang, T.S.; Jeong, W.; Choi, S.Y.; Yu, S.; Kang, S.W.; Rhee,
S.G. Regulation of peroxiredoxin I activity by Cdc2-mediated
phosphorylation. J. Biol. Chem., 2002, 277, 25370-25376.
Kinnula, V.; Lehtonen, S.; Sormunen, R.; Kaarteenaho-Wiik, R.;
Kang, S.; Rhee, S.; Soini, Y. Overexpression of peroxiredoxins I,
II, III, V and VI in malignant mesothelioma. J. Pathol., 2002, 196,
316-323.
Karihtala, P.; Mantyniemi, A.; Kang, S.; Kinnula, V.; Soini, Y.
Peroxiredoxins in breast carcinoma. Clin. Cancer Res., 2003, 9,
3418-3424.
Choi, J.; Kim, T.; Kim, S.; Baek, S.; Kim, J.; Lee, S.; Kim, J.
Overexpression of mitochondrial thioredoxin reductase and
peroxiredoxin III in hepatocellular carcinomas. Anticancer Res.,
2002, 22, 3331-3335.
Zhang, B.; Wang, Y.; Su, Y. Peroxiredoxins, a novel target in
cancer radiotherapy. Cancer Lett., 2009, 286, 154-160.
Neumann, C.A.; Krause, D.S.; Carman, C.V.; Das, S.; Dubey, D.P.;
Abraham, J.L.; Bronson, R.T.; Fujiwara, Y.; Orkin, S.H.; Van
Etten, R.A.Essential role for the peroxiredoxin Prdx1 in erythrocyte
antioxidant defence and tumour suppression. Nature, 2003, 424,
561-565.
Kropotov, A.; Serikov, V.; Suh, J.; Smirnova, A.; Bashkirov, V.;
Zhivotovsky, B.; Tomilin, N. Constitutive expression of the human
peroxiredoxin V gene contributes to protection of the genome from
oxidative DNA lesions and to suppression of transcription of noncoding DNA. FEBS J., 2006, 273, 2607-2617.
Miyazaki, K.; Noda, N.; Okada, S.; Hagiwara, Y.; Miyata, M.;
Sakurabayashi, I.; Yamaguchi, N.; Sugimura, T.; Terada, M.;
Wakasugi, H. Elevated serum level of thioredoxin in patients with
hepatocellular carcinoma. Biotherapy, 1998, 11, 277-288.
Lichtenfels, R.; Kellner, R.; Atkins, D.; Bukur, J.; Ackermann, A.;
Beck, J.; Brenner, W.; Melchior, S.; Seliger, B. Identification of
metabolic enzymes in renal cell carcinoma utilizing PROTEOMEX
analyses. Biochim. Biophys. Acta., 2003, 1646, 21-31.
Lincoln, D.; Ali Emadi, E.; Tonissen, K.; Clarke, F. The
thioredoxin-thioredoxin reductase system: over-expression in
human cancer. Anticancer Res., 2003, 23, 2425-2433.
Raffel, J.; Bhattacharyya, A.K.; Gallegos, A.; Cui, H.; Einspahr,
J.G.; Alberts, D.S.; Powis, G. Increased expression of thioredoxin1 in human colorectal cancer is associated with decreased patient
survival. J. Lab. Clin. Med., 2003, 142, 46-51.
Soini, Y.; Kahlos, K.; Näpänkangas, U.; Kaarteenaho-Wiik, R.;
Säily, M.; Koistinen, P.; Pääakkö, P.; Holmgren, A.; Kinnula, V.L.
Widespread expression of thioredoxin and thioredoxin reductase in
non-small cell lung carcinoma. Clin. Cancer Res., 2001, 7, 17501757.
Powis, G.; Montfort, W. Properties and biological activities of
thioredoxins. Annu. Rev. Biophys. Biomol. Struct., 2001, 30, 421455.
Biaglow, J.; Miller, R. The thioredoxin reductase/thioredoxin
system. Cancer Biology & Therapy, 2005, 4, 6-13.
Gromer, S.; Urig, S.; Becker K. The thioredoxin system – from
science to clinic. Med. Res. Rev., 2004, 24, 40-89.
Ueno, M.; Matsutani, Y.; Nakamura, H.; Masutani, H.; Yagi, M.;
Yamashiro, H.; Kato, H.; Inamoto, T.; Yamauchi, A.; Takahashi,
R.; Yamaoka, Y.; Yodoi, J. Possible association of thioredoxin and
p53 in breast cancer. Immunol. Lett., 2000, 75, 15-20.
Naito, S.; Koga, H.; Yokomizo, A.; Sakamoto, N.; Kotoh, S.;
Nakashima, M.; Kiue, A.; Kuwano, M. Molecular analysis of
mechanisms regulating drug sensitivity and the development of
new chemotherapy strategies for genitourinary carcinomas. World
J. Surg., 2000, 24, 1183-1186.
Base Excision Repair, the Redox Environment
[50]
[51]
[52]
[53]
[54]
[55]
[56]
[57]
[58]
[59]
[60]
[61]
[62]
[63]
[64]
[65]
[66]
[67]
[68]
Brigelius-Flohé, R.; Kipp, A. Glutathione peroxidases in different
stages of carcinogenesis. Biochim. Biophys. Acta, 2009, 1790(11),
1555-1568.
Zhang, Y.; Ikeno, Y.; Qi, W.; Chaudhuri, A.; Li, Y. ; Bokov, A. ;
Thorpe, S.R. ; Baynes, J.W. ; Epstein, C. ; Richardson, A.; Van
Remmen, H. Mice deficient in both Mn superoxide dismutase and
glutathione peroxidase-1 have increased oxidative damage and a
greater incidence of pathology but no reduction in longevity. J.
Gerontol. A. Biol. Sci. Med. Sci., 2009, 64, 1212-1220.
Lu, L.; Oveson, B.C.; Jo, Y-J.; Lauer, T.W. ; Usui, S. ; Komeima,
K. ; Xie, B.; Campochiaro, P.A. increased expression of
glutathione peroxidase 4 strongly protects retina from oxidative
damage. Antioxid. Redox Signal., 2009, 11, 715-724.
Bai, F.; Nakanishi, Y.; Kawasaki, M.; Takayama, K.; Yatsunami,
J.; Pei, X.H.; Tsuruta, N.; Wakamatsu, K.; Hara, N.
Immunohistochemical expression of glutathione S-transferase-Pi
can predict chemotherapy response in patients with non small cell
lung carcinoma. Cancer, 1996, 78, 416-421.
Curtis,
C.D.;
Thorngren,
D.L.;
Nardulli,
A.M.
Immunohistochemical analysis of oxidative stress and DNA repair
proteins in normal mammary and breast cancer tissues. BMC
Cancer, 2010, 10, 9.
Fernandes, A.P.; Capitanio, A.; Selenius, M.; Brodin, O.; Rundlöf,
A-K.; Björnstedt, M. Expression profiles of thioredoxin family
proteins in human lung cancer tissue: correlation with proliferation
and differentiation. Histopathology, 2009, 54, 313-320.
Grogan, T.M.; Fenoglio-Prieser, C.; Zeheb, R.; Bellamy, W.;
Frutiger, Y.; Vela, E.; Stemmerman, G.; Macdonald, J.; Richter, L.;
Gallegos, A.; Powis, G. Thioredoxin, a putative oncogene product,
is overexpressed in gastric carcinoma and associated with increased
proliferation and increased cell survival. Hum. Pathol., 2000, 31,
475-448.
Kahlos, K.; Pääkkö, P.; Kurttila, E.; Soini, Y.; Kinnula, V.L.
Manganese superoxide dismutase as a diagnostic marker for
malignant pleural mesothelioma. B. J. Cancer., 2000, 82, 10221029.
Woolston, C.M.; Zhang, L.; Evans, H.; Al-Attar, A.; Shehata, M.;
Balls, G., Chan, S.Y.; Martin, S.G. Thioredoxin and related redox
systems as targets in breast cancer. Breast Cancer Res., 2010,
12(Suppl. 1), P29.
Woolston, C.M.; Al-Attar, A.; Storr, S.J.; Ellis, I.O.; Morgan, D.A.;
Martin, S.G. Redox protein expression predicts radiotherapeutic
response in early stage invasive breast cancer patients. Int. J.
Radiat. Oncol. Biol. Phys., 2011, 79, 1532-1540.
Woolston, C.M.; Deen, S.; Al-Attar, A.; Shehata, M.; Chan, S.Y.;
Martin, S.G. Redox protein expression predicts progression-free
and overall survival in ovarian cancer patients treated with
platinum based chemotherapy. Free Radic. Biol. Med., 2010, 49,
1263-1272.
Woolston, C.M.; Storr, S.J.; Ellis, I.O.; Morgan, D.A.; Martin, S.G.
Expression of thioredoxin system and related peroxiredoxin
proteins is associated with clinical outcome in radiotherapy in
breast cancer. Radiother. Oncol., 2011, 100(2), 308-313.
Dally, H.; Hartwig, A. Induction and repair inhibition of oxidative
DNA damage by nickel(II) and cadmium(II) in mammalian cells.
Carcinogenesis, 1997, 18, 1021-1026.
Bravard, A.; Vacher, M.; Gouget, B.; Coutant, A.; Hillairet de
Boisferon, F. ; Radicella, J.P. Redox regulation of human OGG1
activity in response to cellular oxidative stress. Mol. Cell. Biol.,
2006, 26, 7430-7436.
Li, H.; Hao, X.; Zhang, W.; Wei, Q.; Chen, K. The hOGG1
Ser326Cys polymorphism and lung cancer risk: a meta-analysis.
Cancer Epidemiol. Biomarkers Prev., 2008, 17, 1739-1745.
Bravard, A.; Vacher, M.; Moritz, E.; Vaslin, L.; Hall, J.; Epe, B.;
Radicella, J.P. Oxidation status of human OGG1-S326C
polymorphic variant determines cellular DNA repair capacity.
Cancer Res., 2009, 69, 3642-3649.
Hegde, V.; Wang, M.; Deutsch, W.A. Human ribosomal protein S3
interacts with DNA base excision repair proteins hAPE/Ref-1 and
hOGG1. Biochemistry, 2004, 43, 14211-14217.
Hegde, V.; Wang, M.; Mian, I.S.; Spyres, L.; Deutsch, W.A. The
high binding affinity of human ribosomal protein S3 to 7,8dihydrp-8-oxoguanine is abrogated by a single amino acid change.
DNA Repair, 2006, 5(7), 810-815.
Wan, F.; Anderson, D.E.; Barnitz, R.A.; Snow, A.; Bidere, N.;
Zheng, L.; Hegde, V.; Lam, L.T.; Staudt, L.M.; Levens, D.;
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
[69]
[70]
[71]
[72]
[73]
[74]
[75]
[76]
[77]
[78]
[79]
[80]
[81]
[82]
[83]
[84]
[85]
[86]
[87]
[88]
99
Deutsch, W.A.; Lenardo, M.J. Ribosomal protein S3: A KH
domain subunit in NF-KB complexes that mediates selective gene
regulation. Cell, 2007, 131, 927-939.
Yadavilli, S.; Mayo, L.D.; Higgins, M.; Lain, S.; Hegde, V.;
Deutsch, W.A. Ribosomal protein S3: A multi-functional protein
that interacts with both p53 and MDM2 through its KH domain.
DNA Repair, 2009, 8, 1215-1224.
Demple, B.; Herman, T.; Chen, T.S. Cloning and expression of
APE1, the cDNA encoding the major human apurinic
endonuclease: Definition of a family of DNA repair enzymes.
Proc. Natl. Acad. Sci. USA, 1991, 88, 11450-11454.
Robson, C.N.; Hickson, I.D. Isolation of cDNA clones encoding a
human apurinic/apyrimidinic endonuclease that corrects DNA
repair and mutagenesis defects in E.coli xth (exonuclease III)
mutants. Nucleic Acids Res., 1991, 19, 5519-5523.
Xanthoudakis, S.; Curran, T. Identification and characterisation of
Ref-1, a nuclear protein that facilitates AP-1 DNA binding activity.
EMBO J., 1992, 11, 653-665.
Xanthoudakis, S.; Miao, G.G.; Curran, T. The redox and DNArepair activities of Ref-1 are encoded by nonoverlapping regions.
Proc. Natl. Acad. Sci. USA, 1994, 91, 23-27.
Kelley, M.R.; Parsons, S.H. Redox regulation of the DNA function
of the human AP endonuclease APE1/Ref1. Antioxid. Redox
Signal., 2001, 3, 671-683.
Ramana, C.C.; Boldogh, I.; Izumi, T.; Mitra, S. Activation of
apurinic/apyrimidinic endonuclease in human cells by reactive
oxygen species and its correlation with their adaptive response to
genotoxicity of free radicals. Proc. Natl. Acad. Sci. USA, 1998, 95,
5061-5066.
Yang, S.; Misner, B.J.; Chiu, R.J.; Meyskens, F.L. Redox effector
factor 1, combined with reactive oxygen species, plays an
important role in the transformation of JB6 cells. Carcinogenesis,
2007, 28, 2382-2390.
Pines, A.; Perrone, L.; Bivi, N.; Romanello, M.; Damante, G.;
Gulisano, M.; Kelley, M.R.; Quadrifoglio, F.; Tell, G. Activation of
APE1/Ref-1 is dependent upon reactive oxygen species generated
after purinergic receptor stimulation by ATP. Nucleic Acids Res.,
2005, 33, 4379-4394.
Hsieh, M.M.; Hegde, V.; Kelley, M.R.; Deutsch, W.A. activation
of APE1/Ref-1 redox activity is mediated by reactive oxygen
species and PKC phosphorylation. Nucleic Acids Res., 2001, 29,
3116-3122.
Dou, H.; Mitra, S.; Hazra, T.K. Repair of oxidised bases in DNA
bubble structures by human DNA glycosylases NEIL1 and NEIL2.
J. Biol. Chem., 2003, 278, 49679-49684.
Das, S.; Chattopadhyay, R.; Bhakat, K.K.; Boldogh, I.; Kohno, K.;
Prasad, R.; Wilson, S.H.; Hazra, T.K. Stimulation of NEIL2mediated oxidative base excision repair via YB-1 interaction during
oxidative stress. J. Biol. Chem., 2007, 282, 28474-28484.
Cuneo, M.J.; London, R.E. Oxidation state of the XRCC1 Nterminal domain regulates DNA polymerase binding affinity.
Proc. Natl. Acad. Sci. USA, 2010, 107, 6805-6810.
Bennett, S.M.; Neher, T.M.; Shatilla, A.; Turchi, J.J. Molecular
analysis of Ku redox regulation. BMC Mol. Biol., 2009, 10, 86.
Andrews, B.J.; Lehman, J.A.; Turchi, J.J. Kinetic analysis of the
Ku-DNA binding activity reveals a redox-dependent alteration in
protein structure that stimulates dissociation of the Ku-DNA
complex. J. Biol. Chem., 2006, 281, 13596-13603.
Ayene, I.S.; Stamato, T.D.; Mauldin, S.K.; Biaglow, J.E.; Tuttle,
S.W.; Jenkins, S.F.; Koch, C.J. Mutation in the glucose-6phosphate dehydrogenase gene leads to inactivation of Ku DNA
end binding during oxidative stress. J. Biol. Chem., 2002, 277,
9929-9935.
Alexander, A.; Cai, S-L.; Kim, J.; Nanez, A. ; Sahin, M. ;
MacLean, K.H. ; Inoki, K. ; Guan, K-L. ; Shen, J.; Person, M.D.;
Kusewitt, D.; Mills, G.B.; Kastan, M.B.; Walker, C.L. ATM
signals to TSC2 in the cytoplasm to regulate mTORC1 in response
to ROS. Proc. Natl. Acad. Sci. USA, 2010, 107, 4153-4158.
Men, L.; Roginskaya, M.; Zou, Y.; Wang, Y. Redox-dependent
formation of disulphide bonds in human replication protein A.
Rapid Commun. Mass Spectrom., 2007, 21, 2743-2749.
Das, A.; Hazra, T.K.; Boldogh, I.; Mitra, S.; Bhakat, K.K.
Induction of the human oxidised base specific DNA glycosylase by
reactive oxygen species. J. Biol. Chem., 2005, 280, 35272-35280.
Huang, L.E.; Arany, Z.; Livingston, D.M.; Bunn, H.F. Activation
of hypoxia inducible transcription factor depends primarily on the
100 Current Molecular Pharmacology, 2012, Vol. 5, No. 1
[89]
[90]
[91]
[92]
[93]
[94]
[95]
[96]
[97]
[98]
[99]
[100]
[101]
[102]
[103]
[104]
[105]
[106]
[107]
[108]
[109]
redox-sensitive stabilisation of its alpha subunit. J. Biol. Chem.,
1996, 271, 32253-32259.
Nishi, T.; Shimizu, N.; Hiramoto, M.; Sato, I.; Yamaguchi, Y.;
Hasegawa, M.; Aizawa, S.; Tanaka, H.; Kataoka, K.; Watanabe,
H.; Handa, H. Spatial redox regulation of a critical cysteine residue
of NFB in-vivo. J. Biol. Chem., 2002, 277, 44548-44556.
Loboda, A.; Stachurska, A.; Dorosz, J.; Wegrzyn, J.; Kozakowska,
M; Jozkowicz, A.; Dulak, J. HIF-1 attenuates REF-1 expression in
endothelial cells: reversal by siRNA and inhibition of
geranylgeranylation. Vascul. Pharmacol., 2010, 51, 133-139.
Ziel, K.A.; Grishko, V.; Campbell, C.C.; Breit, J.F.; Wilson, G.L.;
Gillespie, M.N. Oxidants in signal transduction: impact on DNA
integrity and gene expression. FASEB J., 2005, 19, 387-394.
Pastukh, V.; Ruchko, M.; Gorodnya, O.; Wilson, G.L.; Gillespie,
M.N. Sequence-specific oxidative base modifications in hypoxiainducible genes. Free Radic. Biol. Med., 2007, 43, 1616-1626.
Ruchko, M.V.; Gorodnya, O.M.; Pastukh, V.M.; Swiger, B.M.;
Middleton, N.S.; Wilson, G.L.; Gillespie, M.N. Hypoxia-induced
oxidative base modifications in the VEGF hypoxia-response
element are associated with transcriptionally active nucleosomes.
Free Radic. Biol. Med., 2009, 46, 352-359.
Hirota, K.; Matsui, M.; Iwata, S.; Nishiyama, A; Mori, K.; Yodoi,
J. AP-1 transcriptional activity is regulated by a direct association
between thioredoxin and Ref-1. Proc. Natl. Acad. Sci. USA, 1997,
94, 3633-3638.
Ueno, M.; Masutani, H.; Yamauchi, A.; Jun Arai, R.; Hirota, K. ;
Sakai, T.; Inamoto, T.; Yamaoka, Y.; Yodo, J.; Nikaido, T.
Thioredoxin-dependent Redox Regulation of p53-mediated p21
Activation. J. Biol. Chem., 1999, 274, 35809-35815.
Wei, S.J.; Botero, A.; Hirota, K.; Bradbury, C.M.; Markovina, S.;
Laszlo, A.; Spitz, D.R.; Goswami, P.C.; Yodoi, J.; Gius, D.
Thioredoxin Nuclear Translocation and Interaction with Redox
Factor-1 Activates the Activator Protein-1 Transcription Factor in
Response to Ionizing Radiation. Cancer Res., 2000, 60, 6688-6695.
Liu, H.; Colavitti, R.; Rovira, I.I.; Finkel, T. Redox dependent
transcriptional regulation. Circ. Res., 2005, 97, 967-974.
Levine, A.J. p53, the cellular gatekeeper for growth and division.
Cell, 1997, 88, 323-331.
Lotum, J.; Peled-Kamar, M.; Groner, Y.; Sachs, L. Cellular
oxidative stress and the control of apoptosis by wild-type p53,
cytotoxic compounds and cytokines. Proc. Natl. Acad. Sci. USA,
1996, 93, 9166-9171.
Seemann, S.; Hainaut, P. Roles of thioredoxin reductase 1 and
APE/Red-1 in the control of basal p53 stability and activity.
Oncogene, 2005, 24, 3853-3863.
Achanta, G.; Huang, P. Role of p53 in sensing oxidative DNA
damage in response to reactive oxygen species generating agents.
Cancer Res., 2004, 64, 6233-6239.
Zhou, J.; Ahn, J.; Wilson, S.H.; Prives, C. A role for p53 in base
excision repair. EMBO J., 2001, 20, 914-923.
Buzek, J.; Latonen, L.; Kurki, S.; Peltonen, K.; Laiho, M. Redox
state of tumour suppressor p53 regulates its sequence specific DNA
binding in DNA damaged cells by cysteine 277. Nucleic Acids
Res., 2002, 30, 2340-2348.
Olovnikov, I.A.; Kravchenko, J.E.; Chumakov, P.M. Homeostatic
functions of the p53 tumour suppressor: regulation of energy
metabolism and antioxidant defence. Semin. Cancer Biol., 2009,
19, 32-41.
Saha, T.; Rih, J.K.; Roy, R.; Ballal, R.; Rosen, E.M.
Transcriptional regulation of the base excision repair pathway by
BRCA1. J. Biol. Chem., 2010, 285, 19092-19105.
Masson, M.; Niedergang, C.; Schreiber, V.; Muller, S.; Menissierde Murcia, J.; de Murcia, G. XRCC1 is specifically associated with
poly(ADP-ribose) polymerase and negatively regulates its activity
following DNA damage. Mol. Cell. Biol., 1998, 18, 3563-3571.
Rouleau, M.; Patel, A.; Hendzel, M.J.; Kaufmann, S.H.; Poirier,
G.G. PARP inhibition: PARP1 and beyond. Nat. Rev. Cancer,
2010, 10, 293-301.
Pacher, P.; Szabo, C. Role of the peroxynitrite-poly(ADP-ribose)
polymerase pathway in human disease. Am. J. Pathol., 2008, 173,
2-13.
Catalgol, B.; Wendt, B.; Grimm, S.; Breusing, N.; zer, N.K.;
Grune, T. chromatin repair after oxidative stress: role of PARP
mediated proteasome activation. Free Radic. Biol. Med., 2010, 48,
673-680.
Storr et al.
[110]
[111]
[112]
[113]
[114]
[115]
[116]
[117]
[118]
[119]
[120]
[121]
[122]
[123]
[124]
[125]
[126]
[127]
[128]
[129]
[130]
Yu, S-W.; Wang, H.; Poitras, M.F.; Coombs, C.; Bowers, W.J.;
Federoff, H.J.; Poirier, G.G; Dawson, T.M.; Dawson, V.L.
Mediation of poly (ADP-ribose) polymerase-1 dependent cell death
by apoptosis inducing factor. Science, 2002, 297, 259-263.
Son, Y-O.; Kook, S-H.; Jang, Y-S.; Shi, X. ; Lee, J-C. Critical role
of poly(ADP-ribose) polymerase-1 in modulating the mode of cell
death caused by continuous oxidative stress. J. Cell Biochem.,
2009, 108, 989-997.
Wondrak, G.G. Redox-directed cancer therapeutics: molecular
mechanisms and opportunities Antioxid. Redox Signal., 2009, 11,
3013-3069.
Kang, Y-H.; Yi, M-J.; Kim, M-J.; Park, M-T.; Bae S.; Kang, C-M.;
Cho, C-K.; Park, I-C.; Park, M-J.; Rhee, C.H.; Hong, S-I.; Cheung,
H.Y.; Lee, Y-S.; Lee, S-J. Caspase-independent cell death by
arsenic trioxide in human cervical cancer cells: reactive oxygen
species mediated poly (ADP-ribose) polymerase-1 activation
signals apoptosis-inducing factor release from mitochondria.
Cancer Res., 2004, 64, 8960-8967.
Sidorkina, O.; Espey, M.G.; Miranda, K.M.; Wink, D.A.; Laval, J.
inhibition of poly (ADP-ribose polymerase (PARP) y nitric oxide
and reactive nitrogen oxide species. Free Radic. Biol. Med., 2003,
35, 1431-1438.
Yu, Z.; Kuncewicz, T.; Dubinsky, W.P.; Kone, B.C. Nitric oxidedependent negative feedback of PARP1 trans-activation of the
inducible nitric-oxide synthase gene. J. Biol. Chem., 2006, 281,
9101-9109.
Valko, M.; Leibfritz, D.; Moncol, J.; Cronin, M.T.D.; Mazur, M.;
Telser, J. Free radicals and antioxidants in normal physiological
functions and human disease. Int. J. Biochem. Cell Biol., 2007, 39,
44-84.
Cook, J.A.; Gius, D.; Wink, D.A.; Krishna, M.C.; Russo, A.;
Mitchell, J.B. Oxidative stress, redox, and the tumour
microenvironment. Semin. Radiat. Oncol., 2004, 14, 259-266.
Pennington, J.D.; Wang, T.J.C.; Nguyen, P.; Sun, L.; Bisht, K.;
Smart, D.; Gius, D. Redox-sensitive signalling factors as a novel
molecular targets for cancer therapy. Drug Resis. Updates, 2005, 8,
322-330.
Renschler, M.F. The emerging role of reactive oxygen species in
cancer therapy. Eur. J. Cancer, 40, 1934-1940.
Chen, W.; McBride, W.; Iwamoto, K.; Barber, C.; Wang, C.; Oh,
Y.; Liao, Y.; Hong, J.; de Vellis, J.; Shau, H. Induction of
radioprotective peroxiredoxin-I by ionizing irradiation. J. Neurosci.
Res., 2002, 70, 794-798.
Smart, D.; Ortiz, K.; Mattson, D.; Bradbury, C.; Bisht, K.; Sieck,
L.; Brechbiel, M.; Gius, D. Thioredoxin reductase as a potential
molecular target for anticancer agents that induce oxidative stress.
Cancer Res., 2004, 64, 6716-6724.
Bump, E.A.; Brown, J.M. Role of glutathione in the radiation
response of mammalian cells in vitro and in vivo. Pharmac. Ther.,
1990, 47, 117-136.
Kramer, R.A.; Zakher, J.; Kim, G. Role of the glutathione redox
cycle in acquired and de novo multidrug resistance. Science, 1988,
241, 694-697.
Wartenberg, M.; Hoffmann, E.; Schwindt, H.; Grünheck, F.;
Petros, J.; Arnold, J.R.S.; Hescheler, J.; Sauer, H. Reactive oxygen
species linked regulation of the multidrug resistance transporter Pglycoprotein in Nox-1 overexpressing prostate tumor spheroids.
FEBS lett., 2005, 579, 4541-4549.
Anestal, K.; Prast-Nielsen, S.; Cenas, N.; Arner, E.S.J. Cell death
by SecTRAPs: Thioredoxin reductase as a prooxidant killer of
cells. PLoS ONE, 2008, 3, e1846, 1-16.
Pompella, A.; Visvikis, A.; Paolicchi, A.; De Tata, V.; Casini, A.F.
The changing faces of glutathione, a cellular protagonist. Biochem.
Pharmacol., 2003, 66, 1499-1503.
Rebrin, I.; Sohal, R.S. Pro-oxidant shift in glutathione redox state
during aging. Adv. Drug Delivery Rev., 2008, 60, 1545-1552.
Ravi, D.; Muniyappa, H.; Das, K. Endogenous thioredoxin is
required for redox cycling of anthracyclines and p53-dependent
apoptosis in cancer cells. J. Biol. Chem., 2005, 280, 40084-40096.
Wang, J.; Yi, J. Cancer killing via ROS. Cancer Biol. Ther., 2008,
7, 1875-1884.
Caple, F.; Williams, E.A.; Spiers, A.; Tyson, J.; Burtle, B.; Daly,
A.K.; Mathers, J.C.; Hesketh, J.E. Inter-individual variation in
DNA damage and base excision repair in young, healthy nonsmokers: effects of dietary supplementation and phenotype. Br. J.
Nutr., 2009, 103, 1585-1593.
Base Excision Repair, the Redox Environment
[131]
[132]
[133]
[134]
[135]
[136]
[137]
[138]
[139]
[140]
[141]
[142]
[143]
[144]
Current Molecular Pharmacology, 2012, Vol. 5, No. 1
Larsson, S.C.; Bergkvist, L.; Wolk, A. Dietary carotenoids and risk
of hormone receptor-defined breast cancer in a prospective cohort
of Swedish women. Eur. J. Cancer, 2010, 46, 1079-1085.
Koran, R.; Hadari-Naor, I.; Zuck, E.; Rotem, C.; Liberman, U.A.;
Ravid, A. Vitamin D is a prooxiodant in breast cancer cells. Cancer
Res., 2001, 61, 1439-1444.
Hyodo, F.; Soule, B.P.; Matsumoto, K-I., Matsumoto, S.; Cook,
J.A.; Hyodo, E.; Sowers, A.L.; Krishna, M.C.; Mitchell, J.B.
Assessment of tissue redox status using metabolic responsive
contrast agents and magnetic resonance imaging. J. Pharm.
Pharmacol., 2008, 60, 1049-1060.
Hyodo, F.; Matsumoto, K-I.; Matsumoto, A.; Mitchell, J.B.;
Krishna, M.C. Probing the intracellular redox status of tumors with
magnetic resonance imaging and redox-sensitive contrast agents.
Cancer Res., 2006, 66, 9921-9928.
Matsumoto, K-I.; Hyodo, F.; Matsumoto, A.; Koretsky, A.P.;
Sowers, A.L.; Mitchell, J.B.; Krishna, M.C. High-resolution
mapping of tumor redox status by magnetic resonance imaging
using nitroxides as redox-sensitive contrast agents. Clin. Cancer
Res., 2006, 12, 2455-2462.
Hashemy, S.I.; Ungerstedt, J.S.; Avval, F.Z.; Holmgren, A.
Motexafin Gadolinium, a tumor-selective drug targeting
thioredoxin reductase and ribonucleotide reductase. J. Biol. Chem.,
2006, 281, 10691-10697.
Evens, A.M. Motexafin gadolinium: a redox-active tumor selective
agent for the treatment of cancer. Curr. Opin. Oncol., 2004, 16,
576-580.
Mehta, M.P.; Shapiro, W.R.; Phan, S.C.; Gervais, R.; Carrie, C.;
Chabot, P.; Patchell, R.A.; Glantz, M.J.; Recht, L.; Langer, C.; Sur,
R.K.; Roa, W.H.; Mahe, M.A.; Fortin, A.; Nieder, C.; Meyers,
C.A.; Smith, J.A.; Miller, R.A.; Renschler, M.F. Motexafin
gadolinium combined with prompt whole brain radiotherapy
prolongs time to neurologic progression in non-small-cell lung
cancer patients with brain metastases: results of a phase III trial.
Int. J. Radiat. Oncol. Biol. Phys., 2009, 73, 1069-1076.
Jing, Y.; Dai, J.; Chalmers-Redman, R.M.; Tatton, W.G.; Waxman,
S. Arsenic trioxide selectively induces acute promyelocytic
leukemia cell apoptosis via a hydrogen peroxide-dependent
pathway. Blood, 1999, 94, 2102-2111.
Platanias, L.C. Biological responses to arsenic compounds. J. Biol.
Chem., 2009, 284, 18583-18587.
Shen, Z.X.; Chen, G.Q.; Ni, J.H.; Li, X.S.; Xiong, S.M.; Qiu, Q.Y.;
Zhu, J.; Tang, W.; Sun, G.L.; Yang, K.Q.; Chen, Y.; Zhou, L.;
Fang, Z.W.; Wang, Y.T.; Ma, J.; Zhang, P.; Zhang, T.D.; Chen,
S.J.; Chen, Z.; Wang, Z.Y. Use of arsenic trioxide (As2O3) in the
treatment of acute promyelocytic leukemia (APL): II. Clinical
efficacy and pharmacokinetics in relapsed patients. Blood, 1997,
89, 3354-3360.
Soignet, S.L.; Maslak, P.; Wang, Z.G.; Jhanwar, S.; Calleja, E.;
Dardashti, L.J.; Corso, D.; DeBlasio, A.; Gabrilove, J.; Scheinberg,
D.A.; Pandolfi, P.P.; Warrell, R.P.J.J. Complete remission after
treatment of acute promyelocytic leukemia with arsenic trioxide. N.
Engl. J. Med., 1998, 339, 1341-1348.
Murphy, C.K.; Fey, E.J.; Watkins, B.A.; Wong, V.; Rothstein, D.;
Sonis, ST. Efficacy of superoxide dismutase mimetic M40403 in
attenuating radiation induced oral mucositis in hamsters. Clin.
Cancer Res., 2008, 14, 4292-4297.
Harrison, M.R.; Hahn, N.M.; Pili, R.; Oh, W.K.; Hammers, H.;
Sweeney, C.; Kim, K.; Perlman, S.; Arnott, J.; Sidor, C.; Wilding,
G.; Liu, G. A phase II study of 2-methoxyestradiol (2ME2) nano
crystal dispersion in patients with taxane refractory metastatic
castrate resistant prostate cancer (CRPC). Invest. New Drugs, 2011,
29(6), 1465-1474.
[145]
[146]
[147]
[148]
[149]
[150]
[151]
[152]
[153]
[154]
[155]
[156]
[157]
[158]
[159]
[160]
[161]
Received: June 16, 2010
101
Sweeney, C.; Liu, G.; Yiannoutsos, C.; Kolesar, J.; Horvath, D.;
Staab, M.J.; Fife, K.; Armstrong, V.; Treston, A.; Sidor, C.;
Wilding, G. A phase II multicentre, randomised, double blind,
safety trial assessing the pharmacokinetics, pharmacodynamics,
and efficacy of 2-methoxyestradiol capsules in hormone-refractory
prostate cancer. Clin. Cancer Res., 2005, 11, 6625-6633.
Liu, G.; Botting, C.H.; Evans, K.M.; Walton, J.A.G.; Xu, G.;
Slawin, A.M.Z.; Westwood, N.J. Optimisation of conoidin A, a
peroxiredoxin inhibitor. Chem. Med. Chem., 2010, 5, 41-45.
Bar-Am, O.; Weinreb, O.; Amit, T.; Youdim, M.B.H. The novel
cholinesterase-monoamine oxidase inhibitor and anti-oxidant,
ladostigil, confers neuroprotection in neuroblastoma cells and aged
rats. J. Mol. Neurosci., 2009, 37, 135-145.
Wang, T.; Tamae, D.; LeBon, T. ; Shively, J.E.; Yen, Y.; Li, J.J.
The role of peroxiredoxin II in radiation resistant MCF7 breast
cancer cells. Cancer Res., 2005, 65, 10338-10346.
Bradshaw, T.D.; Matthews, C.S.; Cookson, J.; Chew, E.H.; Shah,
M.; Bailey, K.; Monks, A.; Harris, E.; Westwell, A.D.; Wells, G.;
Laughton, C.A.; Stevens, M.F.G. Elucidation of thioredoxin as a
molecular target for antitumor quinols. Cancer Res., 2005, 65,
3911-3919.
Mukherjee, A.; Martin, S.G. The thioredoxin system: a key target
in tumour and endothelial cells. Br. J. Pharmacol., 2007, 151,
1167-1175.
Kirkpatrick, D.L.; Jimale, M.L.; King, K.M.; Chen, T. Synthesis
and evaluation of imidazolyl disulfides for selective cytotoxicity to
hypoxic EMT6 tumor cells in vitro. Eur J. Med. Chem., 1992, 27,
33-37.
Ramanathan, R.K.; Abbruzzese, J.; Dragovich, T.; Kirkpatrick, L.;
Guillen, J.M.; Baker, A.F.; Pestano, L.A.; Green, S.; VonHoff, D.
A randomised phase II study of PX-12, an inhibitor of thioredoxin
in patients with advanced cancer of the pancreas following
progression on gemcitabine containing combination. Cancer
Chemother. Pharmacol., 2011, 67(3), 503-509.
Greenstein, J.P.; Venrette, W.V.; White, J. The liver catalase
activity of tumour-bearing rats and the effect of extirpation of the
tumours. J. Biol. Chem., 1941, 141, 327-328.
Margoliash, E.; Novogrodsky, A.; Schejter, A. Irreversible reaction
of 3-amino-1:2:4-triazole and related inhibitors with the protein of
catalase. Biochem. J., 1960, 74, 339-348.
Hiasa, Y.; Ohshima, M.; Kitahori, Y.; Yuassa, T.; Fujita, T.; Iwata,
C. Promoting effects of 3-amino-1,2,4-triazole on the development
of thyroid tumours in rats treated with N-bis(2hydroxylpropyl)nitrosamine. Carcinogenesis, 1982, 3, 381-384.
Day, B.J. Catalase and glutathione peroxidase mimics. Biochem.
Biopharmacol., 2009, 77, 285-296.
Fishel, M.L.; Kelley, M.R. the DNA base excision repair protein
Ape1/Ref-1 as a therapeutic and chemopreventative target. Mol.
Aspects Med., 2007, 28, 375-395.
Nyland, R.L.; Luo, M.; Kelly, M.R.; Borch, R.F. Design and
synthesis of novel quinone inhibitors targeted to the redox function
of apurinic/apyrimidinic endonuclease/redox enhancing factor-1
(Ape1/Ref-1). J. Med. Chem., 2010, 53, 1200-1210.
Martin, S.A.; McCarthy, A.; Barber, L.J.; Burgess, D.J.; Parry, S.;
Lord, C.J.; Ashworth, A. Methotrexate induces oxidative DNA
damage and is selectively lethal to tumour cells with defects in the
DNA mismatch repair gene MSH2. EMBO Mol. Med., 2009, 1,
323-337.
Loft, S.; Møller, P.; Cooke, M.S.; Rozalski, R.; Olinski, R.
Antioxidant vitamins and cancer risk: is oxidative damage to DNA
a relevant biomarker. Eur. J. Nutr., 2008, 47(suppl. 2), 19-28.
Abbotts, R.; Madhusudan, S. Human AP endonuclease 1 (APE1):
from mechanistic insights to druggable target in cancer. Cancer
Treat. Rev., 2010, 36(5), 425-435.
Accepted: July 15, 2010
PMID: 22122466