The role of MTH1 in ultraviolet radiationinduced mutagenesis
Asal Fotouhi
Doctoral Thesis in Molecular Genetics.
Department of Molecular Biosciences, The Wenner-Gren Institute.
Stockholm University, Sweden.
2015
Cover illustrations by Farhad Heidarian
© Asal Fotouhi
ISBN 978-91-7649-096-9
Printed in Sweden by Universitetsservice AB (US-AB), Stockholm 2015
Distributor: Stockholm University
Dedicated to my mother
“No great thing is created suddenly”
Epictetus
Abstract
Ultraviolet radiation (UVR) is known to be highly mutagenic. What types of
DNA lesions that are induced by different UVR wavelengths are still a matter of
debate. UVR induces mutagenesis mostly by the formation of photoproducts and
the induction of reactive oxygen species (ROS). ROS can give rise to mutations via
oxidation of nucleotides in the DNA or the nucleotide pool. Oxidized nucleotides in
the nucleotide pool can thereby be incorporated into the DNA during replication and
ultimately give rise to mutations. MTH1 however, dephosphorylates oxidized
nucleotides in the nucleotide pool, in particular 8-oxo-dGTP and 2-OH-dATP, and
inhibits their incorporation into the DNA.
The aim of the present study was to investigate the role of MTH1 in
mutagenesis and cytogenetic damage induced by UVR in a human lymphoblastoid
TK6 cell line. The clonogenic survival, mutant frequency and micronucleus
frequency were measured following exposure to UVA, UVB and UVC in MTH1knockdown and wild-type TK6 cells. As a biomarker for oxidative damage the level
of intracellular and extracellular 8-oxo-dG was measured in TK6 cells exposed to
UVA. The mutational spectra of UVA-induced mutations at the thymidine kinase
gene in MTH1-knockdown and wild-type TK6 cells were investigated.
The results show that MTH1 protects against UVA and UVB mutagenesis
significantly. MTH1, however, has been shown to offer no protection against UVRinduced cytogenetic damage and is therefore suggested to mainly inhibit
mutagenesis. The mutational spectra show that GC>AT and AT>GC transitions are
the dominant mutation types in cells exposed to UVA.
In conclusion, MTH1 protects TK6 cells against mutagenesis induced by
longer wavelengths of UVR. This indicates that the nucleotide pool is a significant
target in mutagenesis for longer wavelengths of UVR. The type of mutations
induced by UVA, GC>AT and AT>GC, can be formed by the incorporation of 2-OHdATP from nucleotide pool into the DNA. UVA is therefore suggested to induce
mutations by induction of oxidized nucleotides such as 2-OH-dATP.
List of publications
This thesis is based on data presented in the following publications and manuscript:
Paper I
Fotouhi A., Skiold S., Shakeri-Manesh S., Osterman-Golkar S., Wojcik A., Jenssen
D., Harms-Ringdahl M., and Haghdoost S., Reduction of 8-oxodGTP in the
nucleotide pool by hMTH1 leads to reduction in mutations in the human
lymphoblastoid cell line TK6 exposed to UVA. Mutat Res, 2011. 715 (1-2): p. 13-18.
Paper II
Fotouhi A., Woldai Hagos W., Ilic M., Wojcik A., Harms-Ringdahl M., de Gruijl F.,
Mullenders L., Jansen J.G., and Haghdoost S., Analysis of mutant frequencies and
mutation spectra in hMTH1 knockdown TK6 cells exposed to UV radiation. Mutat
Res, 2013. 751-752: p. 8–14.
Paper III
Fotouhi A., Cornella N., Ramezani M., Wojcik A., and Haghdoost S., Investigation
of micronucleus induction in MTH1 knockdown cells exposed to UVA, UVB, and
UVC, 2014. Submitted Manuscript.
The following publications are not included in the thesis:
Manesh S.S., Deperas-Kaminska M., Fotouhi A., Sangsuwan T., Harms-Ringdahl
M., Wojcik A., and Haghdoost S., Mutations and chromosomal aberrations in
hMTH1-transfected and non-transfected TK6 cells after exposure to low dose rates
of gamma radiation. Radiat Environ Biophys, 2014. 53: p. 417-425.
Manesh S.S., Sangsuwan T., Fotouhi A., Emami N., and Haghdoost S.,
Cooperation of MTH1 and MYH proteins in response to oxidative stress induced by
chronic gamma radiation, 2014. Submitted Manuscript.
Permission to reproduce the published papers was obtained from the publishers.
Table of contents
Abbreviations .......................................................................................................... 1
Introduction ............................................................................................................. 2
Ultraviolet radiation ...................................................................................................... 2
UVR induced damage.................................................................................................. 3
DNA as a target of UVR ............................................................................................... 4
Repair of pyrimidine dimers ......................................................................................... 6
Mechanism of UVR-induced ROS................................................................................ 7
ROS-induced oxidative damage .................................................................................. 9
The nucleotide pool and DNA as targets of ROS ......................................................... 9
Repair of oxidative nucleotide damage ...................................................................... 11
Polymorphism in MTH1 and its role in mutagenesis and carcinogenesis ................... 13
UVR-induced cytogenetic damage............................................................................. 14
UVR-induced epigenetic alterations ........................................................................... 14
UVR-induced nucleotide pool imbalance ................................................................... 15
UVR-sensitivity syndromes ........................................................................................ 16
Aim ......................................................................................................................... 18
Methodology ......................................................................................................... 19
Cell culture ................................................................................................................ 19
Transfection of TK6 cells ........................................................................................... 19
Western blot .............................................................................................................. 19
Irradiation with UV ..................................................................................................... 20
Clonogenic survival.................................................................................................... 21
Mutant frequency ....................................................................................................... 21
Measurement of intra- and extracellular 8-oxo-dG and dG following UVA irradiation . 22
Isolation of mutants and RNA .................................................................................... 22
cDNA isolation ........................................................................................................... 23
Sequencing................................................................................................................ 23
Micronucleus assay ................................................................................................... 24
Quantitative Real-Time PCR ...................................................................................... 25
Statistical analysis ..................................................................................................... 25
Experimental design .................................................................................................. 26
Results................................................................................................................... 27
Paper I ....................................................................................................................... 27
Paper II ...................................................................................................................... 29
Paper III ..................................................................................................................... 31
Discussion ............................................................................................................ 32
Ongoing studies and future perspective ............................................................ 35
Acknowledgements .............................................................................................. 36
References ............................................................................................................ 38
Abbreviations
BER
Base excision repair
CPD
Cyclobutane pyrimidine dimer
dA
2’-Deoxyadenosine
dC
2’-Deoxycytosine
dG
2’-Deoxyguanosine
dT
2’-Deoxythymidine
dNTP
Deoxyribonucleoside-triphosphate
dTTP
Deoxyribothymidine-triphosphate
dCTP
Deoxyribocytidine-triphosphate
DNMT
DNA methyltransferase
DSB
Double strand break
hMYH
Human MutY homolog protein
hOGG1
Human MutM homolog protein
H2O2
Hydrogen peroxide
MN
Micronucleus
MTH1
Human MutT homolog protein 1
NER
Nucleotide excision repair
N-Tr
Non-transfected
•
OH
Hydroxyl radical
2-OH-dA
2-Hydroxy-2’-deoxyadenosine
2-OH-dAMP
2-Hydroxy-2’-deoxyadenosine 5′-monohosphate
2-OH-dATP
2-Hydroxy-2’-deoxyadenosine 5′-triphosphate
8-oxo-dG
8-Oxo-7,8-dihydro-2’-deoxyguanosine
8-oxo-dGMP
8-Oxo-7,8-dihydro-2’-deoxyguanosine-5’-monophosphate
8-oxo-dGTP
8-Oxo-7,8-dihydro-2’-deoxyguanosine-5’-triphosphate
PCR
Polymerase chain reaction
6-4PP
Pyrimidine (6-4) pyrimidone photoproduct
ROS
Reactive oxygen species
TK6 cells
Human B lymphoblastoid cells
Tk
Thymidine kinase
UVA
Ultraviolet A radiation, λ = 320-400 nm
UVB
Ultraviolet B radiation, λ = 280-320 nm
UVC
Ultraviolet C radiation, λ = 200-280 nm
UVR
Ultraviolet radiation
1
Introduction
Ultraviolet radiation
The Sun is a vital source of energy for life on Earth. Sunlight provides us with
warmth, light and oxygen (by driving the photosynthesis) for our well-being. The
range of shorter wavelengths of the sunlight, called ultraviolet radiation (UVR), was
discovered in 1801 by the German physicist Johann Wilhelm Ritter [1]. It comprises
wavelengths from 200 up to 400 nm and can be classified into three types;
ultraviolet A (UVA 320-400 nm), ultraviolet B (UVB 280-320 nm), and ultraviolet C
(UVC 200-280 nm). Approximately 95% of the UVR reaching Earth is UVA and 5%
UVB, while most of UVB and all UVC are absorbed by the ozone layer [2].
UVR is essential for the body as it induces vitamin D production which is
important for calcium absorption [3]. However, too much of UVR can be dangerous
and lead to acute effects such as sunburn and long-term effects such as skin
cancer and cataract [4]. Skin cancer is one of the most frequently diagnosed cancer
types in the world. According to WHO statistics, globally, one in every three cancers
diagnosed is skin cancer. Only in Sweden there are 8000 new cases of different
types of skin cancer diagnosed year 2010. In many Asian countries where the
majority of people avoid sun exposure there is a lower incidence of skin cancer [5].
Despite the high incidence of skin cancers, sun exposure to humans increases due
to traveling, tanning devices and depletion of the ozone layer.
UVR is an exogenous agent that does not penetrate the human body any
deeper than the skin, and is absorbed by the skin layers in a wave-dependent
manner (figure 1). Whilst UVB only reaches the outer layers of the skin (epidermis),
UVA infiltrates deeper into the dermis [6]. The skin is protected against the
deleterious effects of UVR by the production of melanin. Melanin is produced by the
melanocytes in the dermis and is thereafter transported to the keratinocytes in the
epidermis to shield the DNA [7].
2
shorter wavelengths however are directly absorbed by DNA. The UV absorption
spectrum for DNA ranges between 220 to 300 nm with the highest absorption peak
at 260 nm. The longer wavelengths of UVR are therefore suggested to induce
damage in an indirect manner [2, 8]. The indirect or direct mechanisms are however
not limited to UVA or UVB. There are studies observing the same type of DNA
damage induced by UVA and UVB, which means that UVA is also able to induce
damage to the DNA through photo-induced reactions [9]. The direct mechanism
triggered by UVA may act via photosensitized triplet energy transfer from an excited
sensitizer to DNA [10, 11]. There are also studies showing that UVB induces
photosensitized processes [12].
Although oxidative nucleotide damage and pyrimidine dimers are the most
studied types of UVR-induced lesions, UVR can also induce oxidative protein
damages or DNA strand breaks [2, 13]. All damages mentioned except pyrimidine
dimers could also be induced by other agents, which make pyrimidine dimers a
signature of UVR exposure.
DNA as a target of UVR
The photo-induced reactions involve absorption of UVR by DNA leading to
formation of covalent bonds between two adjacent pyrimidines to give so called
pyrimidine dimers. There are two types of pyrimidine dimers, differing in the location
of the covalent bonds that are formed between the pyrimidines; cyclobutane
pyrimidine dimers (CPDs) and pyrimidine (6-4) pyrimidone photoproducts (6-4PPs)
(figure 2). CPDs are formed when two bonds connect the C5 and C6 of adjacent
pyrimidine dimers, while 6-4PPs are formed when the C6 position of one pyrimidine
dimer is covalently connected to the C4 position of an adjacent pyrimidine [14]. The
6-4PPs are converted to their Dewar valence isomers when exposed to longer
wavelengths of UVR and can revert to 6-4PPs when exposed to shorter
wavelengths of UVR [15]. The most commonly observed CPDs and 6-4PPs are at
TT and TC sequences, respectively. The CPDs and 6-4PPs are formed at a ratio of
3:1 which makes CPDs the major UV-induced lesion [11]. UV-induced DNA purine
photoproducts have also been recognized, but to a lesser extent [2].
4
Repair of pyrimidine dimers
The repair of CPDs and 6-4PPs is carried out by nucleotide excision repair
(NER). NER is divided into two pathways; global genome repair (GGR) and the
transcription coupled repair (TCR). GGR operates on damages anywhere in the
whole genome while TCR operates on damages in the actively transcribed strand.
During NER, a short single-stranded DNA segment including the damaged DNA is
excised and removed. The gap is thereafter filled by DNA polymerase and ligated
by DNA ligase [18] (figure 4). Damage, repaired by GGR, can be recognized by
XPC-hHR23B-Cen2 or DDB complexes depending on the type of damage. The
XPC-hHR23B-Cen2 recognizes damages such as 6-4PPs while the DDB complex
recognizes damages such as CPDs. The ten-component transcription factor protein
complex (II), TFIIH, is thereafter recruited to the damage site and unwinds the DNA
creating an open-complex structure of about 30 nucleotides around the lesion. XPA
can then bind together with RPA (a single-strand DNA binding protein) to the
damage site to confirm the presence of the damage and stabilizes the opencomplex. Thereafter the damage can be excised by the endonucleases XPG which
splices on the 3’ side of the damage and ERCC1-XPF which splices on the 5’ side
of the damage. The resulting gap will be filled by either DNA polymerase delta or
epsilon assisted by PCNA. The remaining nick can then finally be ligated by a DNA
ligase III. GGR and TCR both share the same proteins except for XPC, which is
replaced by the CSA and CSB in TCR. The RNA polymerase II is stalled by the
lesion which can attract CSA and CSB to recruit the entire repair protein apparatus
to the damage [18].
6
ROS-induced oxidative damage
ROS can be produced by exogenous factors such as gamma radiation,
chemicals or - as already mentioned - UVR. ROS are however also produced
endogenously by several cellular organells such as the mitochrondria, peroxisomes
and the endoplasmic reticulum [21-24]. ROS are continously produced by cells to
help maintain homeostasis and play an important role as signaling molecules. The
list of ROS-mediated pathways includes the generation of ATP in mitochondria [25],
apoptosis in defective cells and, killing of micro-organisms and cancer cells by the
respiratory burst from phagocytes [26]. High levels of ROS can damage cellular
proteins, lipids, DNA, and mtDNA that can lead to diseases such as cancer and
neurodegeneration disorders [27-29]. The rate of endogenously formed DNA
lesions formed by ROS is estimated to be in the range of 10 000 to 20 000/cell/day
[28, 30] which is expected to increase during oxidative stress. The cell is protected
against the harmful effects of ROS by the action of antioxidants. Oxidative stress
arises when the level of ROS exceeds the level of the cell’s antioxidant defense
mechanisms [31]. Among the enzymes included in the antioxidant defense
mechanisms are superoxide dismutase, catalase, and glutathione peroxidase. The
highly reactive superoxide anions (O2•· ) can be converted to hydrogen peroxide
(H2O2) by superoxide dismutase. Hydrogen peroxide can thereafter be converted to
the most reactive radicals, the hydroxyl radical (•OH), by a Fenton reaction.
Catalase or glutathione peroxidase act to prevent the damaging effect of •OH by
removing H2O2 [31].
The nucleotide pool and DNA as targets of ROS
ROS can react with nucleotides in DNA as well as in the nucleotide pool [32].
The most readily oxidized nucleotide base is guanine due to its low redox potential
[33]. Formation of oxidized guanine was first reported in 1984 by Kasai and
Nishimura where they found that if deoxyguanosine is oxidized at the C8 position it
will form 8-oxo-7,8-dihydro-2’-deoxyguanosine (8-oxo-dG) (figure 6A) [34]. This is,
however, not the case for 2-hydroxy-2’-deoxyadenosine (2-OH-dA) which is instead
oxidized at the C2 position (figure 6B) [32]. DNA in the nucleus is tightly packed
9
Repair of oxidative nucleotide damage
In order to correctly pass the genetic information to the next generation, the
genome must be accurately replicated in each cell division. Hence, the involvement
of high fidelity DNA polymerases is essential. However, it has been established that
oxidized nucleotides in DNA or in the nucleotide pool can promote erroneous
incorporation of nucleotides by DNA polymerases during DNA replication [46, 47].
Different types of mutations (transitions and transversions) can arise depending on
the base that is modified. 8-Oxo-dGTP can be misincorporated into the DNA
opposite dA leading to TA>GC transversion after replication. 2-OH-dATP can be
misincorporated into the DNA opposite dG, dC or dT leading to CG>AT, GC>AT,
and AT>GC transitions, respectively, after replication. Direct modifications in the
DNA such as 8-oxo-dG can base pair with dA during replication and lead to GC>TA
transversion (figure 7) [48-50].
To counteract the mutations induced by ROS, cells have developed multiple
defense and repair mechanisms. Once ROS react with a DNA base, base excision
repair (BER) pathways will excise the modified base, e.g. 8-oxo-dG and 2-OH-dA.
The main components of BER for repairing 8-oxo-dG and 2-OH-dA are hOGG1 and
hMYH proteins, both of which comprise DNA glycosylase activities. hOGG1 exerts
its effect by removing 8-oxo-G in DNA, allowing further repair proteins to recover the
damage. If 8-oxo-dG is presented to the DNA during replication, dA can be
misincorporated opposite to 8-oxo-dG. This mismatch can be repaired by the action
of hMYH, which removes dA and allows repair of 8-oxo-dG by hOGG1 [51-53].
11
Polymorphism in MTH1 and its role in mutagenesis and carcinogenesis
Several studies that have been investigating the function of MTH1 in vitro
and in vivo show the importance of MTH1 in cell maintenance. MTH1 is encoded by
the MTH1 gene that is localized on chromosome 7p22 and consists of five exons
and encompasses a size of 8 kb [59]. However, genetic polymorphism is found in
MTH1 resulting in different variants of the protein. A single nucleotide polymorphism
(SNP) at the first nucleotide of codon 83, which results in an amino acid change
from valine (Val: GTG) to methionine (Met: ATG) was found in the MTH1 gene. This
SNP decreased the activity of MTH1 in vitro and has been shown to increase the
risk of cancer in the stomach [60]. Patients with tumor and diseases such as
Parkinson and Alzheimer have been shown to have increased levels of 8-oxo-dG
and need therefore more MTH1 to handle the oxidative stress [61]. Indeed, an overexpression of MTH1 at the mRNA level in human renal cell carcinomas [62], breast
cancer [63] and lung cancer cell lines [64] has been observed. Increased levels of
MTH1 have been found in brain tumors [65] and lung cancer cells [66]. The
substantia nigra of Parkinson disease patients also shows an increased level of
MTH1 [67]. A recent study by the group of Helleday, has been looking at the
survival of cancer cells with a knockdown of MTH1. This study indicates that cancer
cells need MTH1 for their survival [68]. We have previously shown that a low
expression of MTH1 leads to higher formation of oxidized dGTP and mutagenesis in
a human lymphoblastoid cell line [69, 70]. MTH1 has also been shown to have an
important role in normal cells. MTH1 null mice (MTH1-/-), lacking expression of the
MTH1 protein, had a 2-fold increase in spontaneous mutation rate, and an increase
of tumor incidents in lung, liver and stomach [71]. Another study with MTH1-null
mouse embryo fibroblasts cells showed that the H2O2 effects, such as lower survival
and increased levels of 8-oxo-dG, were suppressed by the wild-type human MTH1
[72].
13
UVR-induced cytogenetic damage
UVR is able to induce both ROS as well as pyrimidine dimers, which can
both lead to DNA breaks by different mechanisms. ROS can induce single strand
(SSB) and double strand (DSB) breaks either directly or indirectly via oxidizing
nucleotides such as dG/dGTP in DNA/nucleotide pool. Hydroxyl radicals, the most
reactive oxygen species, are able to interact with DNA and cause SSBs, whereby
two neighboring SSBs on opposite strand can in turn lead to a DSB [73, 74]. ROSoxidized bases as well as pyrimidine dimers in close proximity and on opposite
complementary DNA strands during excision repair can leave gaps that would form
a DSB [75, 76]. Unrepaired DSB can further lead to chromosomal breakages
causing micronucleus (MN) induction. MN are chromosome/chromatid fragments or
whole chromosomes that are lagging during mitosis and are not included in the
daughter nuclei due to chromosome breakage or a defective spindle apparatus.
These fragments or whole chromosomes are then enclosed by a nuclear membrane
and are visualized as a small nucleus in the cytoplasm [77]. Since UVR is able to
induce both ROS and pyrimidine dimers it is a potent chromosomal damage
inducer. There are several in vitro and in vivo studies showing that UVA and UVB
are both able to induce MN [78, 79]. There are however fewer studies investigating
the involvement of UVC in MN induction, even though UVC is becoming
increasingly of interest due to depletion of the ozone layer. The exact mechanism
by which UVR induces DSBs is not clear. For example, there are contradicting
results on whether UVA is able to induce DSBs [80, 81].
UVR-induced epigenetic alterations
Both genetic and epigenetic events are linked to the development of skin
cancer. Whilst most studies focus on genetic events following exposure to UVR,
studies of epigenetic effects have recently come into focus. Epigenetic effects are
mechanisms that can regulate gene expression. One mechanism involved in
epigenetic events is DNA base methylation (gene silencing). DNA base methylation
is the addition of a methyl group to a cytosine converting it to 5-methylcytosine. The
cytosines that are methylated are typically found to be part of cytosine
phosphodiester guanine (CpG) sites found in regions known as CpG islands [82].
14
CpG islands are usually localized to promoter regions of the gene. There are
specific proteins that bind to methylated CpGs and lead to repression of
transcription. Thus, when CpG islands are hyper-methylated the gene may be
repressed whilst when hypo-methylated the gene may be expressed [83].
Methylations are catalyzed by DNA methyltransferases; DNMT1 is responsible for
the maintenance of DNA methylation while DNMT3A and DNMT3B play their main
role in de novo methylation [84].
UVR-induced nucleotide pool imbalance
The cell needs a balanced amount of dNTPs in the pool, with accurate
proportions in both the nucleus and the mitochondria. If there is an imbalance in the
dNTP concentration, it can lead to mutagenesis as well as chromosomal
abnormalities, DNA deletions, and cell death with characteristics of apoptosis [85,
86]. dNTPs can either be synthesized via the de-novo pathway or the salvage
pathway. The dNTPs can also be transferred to the mitochondria [85]. The
synthesis pathways of dNTPs are controlled through allosteric regulations.
Mutations affecting the allosteric sites can lead to loss of control and cause
imbalance in the nucleotide pool. The ribonucleotide reductase (RNR) enzyme has
a dominant role in dNTP synthesis, converting NDPs to dNTPs. A defective RNR
has been shown to lead to an imbalanced nucleotide pool and increased
spontaneous mutation rate [87, 88]. DNA replication and DNA repair are
mechanisms requiring dNTPs in adequate and balanced amounts. A nucleotide
pool imbalance may induce aberrant DNA replication and DNA repair by
misincorporation of an elevated dNTP from the pool into the DNA [89]. Nucleotide
pool imbalance can appear after exposure to UVR and other physical factors. ROS
production by UVR could increase oxidation of dNTPs thus leading to lower levels
of non-oxidized dNTPs in the pool causing an imbalance. In many studies UVC (254
nm) has been shown to alter the dNTPs concentration, promoting an imbalance; an
increase of both dATP and dTTP while less of an increase or even a drop of dCTP
and no change in dGTP has been reported [90, 91].
15
UVR-sensitivity syndromes
UVR and DNA repair deficiencies can cause photoaging [92], erythema [93],
immunosuppression [94] and photosensitive syndromes such as cutaneous
malignant melanoma (CMM) and non-melanomas [95] and NER-deficient
syndromes [96]
CMM arising from epidermal melanocytes, is very aggressive and exhibits a
high metastatic feature, whereby the majority of CMM incidences lead to death [97].
Xeroderma pigmentosum patients (having deficient NER repair protein) have been
observed to have an increased onset of CMM. Most of the xeroderma pigmentosum
patients show melanoma in the face, indicating that CMM arises from sun exposure
[98]. This suggests that deficiencies in DNA repair leads to UVR-induced CMM,
among several other mutated essential genes such as CDKN2, CDK4, XRCC3 and
BRAF which have been shown to contribute to the CMM development [99-101].
Different types of mutations have been observed in the mentioned genes related to
CMM. The CDKN2 gene showed the UVR-signature damage, C>T transversions
[102], while the BRAF gene showed T>A transversions which does not indicate any
UVR type of mutation [103]. The UVR relation to CMM is not clear and is discussed
by Moan et al. where arguments are listed for and against a relationship between
UVR and CMM [104]. Other factors than UVR that influence the risk of an individual
developing CMM include skin type, family history of CMM, a higher number of naevi
and freckles and increasing age [105].
Non-melanomas are the most common type of skin cancers and are, in
contrast to CMM, more likely to have a strong relationship with UVR exposure.
Similar to CMM, skin type is a factor influencing the development of nonmelanomas in individuals [105]. The two major forms of non-melanomas are basal
cell carcinoma (BCC) and squamous cell carcinoma (SCC). Both SCC and BCC
arise from epidermal keratinocytes. BCC being the more common type, grows
slowly and rarely metastasizes, in contrast to the rarer type, SCC that is more likely
to metastasize [106]. BCC and SCC rarely result in death [107]. Mutations in the
p53 gene have been observed to be related to non-melanomas [108, 109]. The
mutations found in the p53 gene of BCCs and SCCs indicate point mutations being
C>T and CC>TT changes at pyrimidine sites suggesting the involvement of the
shorter wavelengths of UVR [110]. A study by Van Kranen et al. showed that
16
mutations in the p53 gene were observed more frequently in UVB-induced tumors
than UVA-induced tumors. This indicates that the main types of damage in nonmelanomas are pyrimidine dimers and that UVB is responsible for those [111].
Defective NER proteins can lead to rare recessive photosensitivity
syndromes such as xeroderma pigmentosum (XP), Cockayne syndrome (CS) and
trichothiodystrophy (TTD) (table 1). Patients with these syndromes suffer from
increased sensitivity to the Sun, whilst XP patients differ from CS and TTD patients
by their high susceptibility of developing cancer. Mutations in 7 XP genes (XPAXPG) have been found to cause XP. CS is caused by mutations in the CSA or CSB
gene whereas mutations in the XPB, XPD or TTD-A gene cause TTD [96].
Table 1. DNA repair deficient syndromes.
Syndrome
Affected gene
Xeroderma pigmentosum (XP)
XP-A to XP-G
Cockayne syndrome (CS)
CSA and CSB
thiodystrophy (TTD)
TFIIH subunits: TTD-A, XPB or XPD
17
Aim
It has previously been reported by our group that low doses of ionizing
radiation, i.e. the mGy range, induce an endogenous release of ROS that can be
monitored indirectly by the subsequent formation of extracellular 8-oxo-dG [112]. It
was proposed that extracellular 8-oxo-dG originates from 8-oxo-dGTP, as the
nucleotide pool is a significant target of ROS. The results showed that the
production of 8-oxo-dGTP in the nucleotide pool by ionizing radiation is caused by
ROS, which are generated by a radiation-triggered endogenous stress response.
Based on these observations we hypothesized that UV radiation can also induce
ROS, which can give rise to dNTP oxidation and consequently to a higher mutation
frequency. There is an ongoing debate regarding UVR mutagenicity. Although it is
known that UVR can induce both indirect and direct DNA damage, the exact
mechanism is still unclear. The aim of the present project was to investigate the role
of MTH1 and dNTP oxidation in UVR-induced mutagenesis. For this purpose we
stably transfected human lymphoblastoid TK6 cells with shRNA against MTH1.
MTH1 prevents incorporation of oxidized nucleotides from the nucleotide pool into
the DNA. The hypothesis was that the longer wavelengths of UVR would mainly
modify dNTPs, which may lead to an increased mutation frequency, originating from
the incorporation of modified dNTP into the DNA. Clonogenic survival, mutant
frequency, the level of 8-oxo-dG and MN frequency following exposure to UVR have
also been investigated. Furthermore, the mutation spectrum in the Tk gene of UVAirradiated cells has been analyzed.
18
Methodology
Cell culture
Human lymphoblastoid cells (the TK6 cell line) were cultured in complete
RPMI-1640 medium. The cells were kept in flasks at 37 °C in 5% CO2. A cell
concentration of 1.0 - 1.5×105 cells/ml was kept throughout the whole experiment.
Transfection of TK6 cells
TK6 cells were stably transfected using a protocol provided by SigmaAldrich.
Hexadimethrine bromide was added to the cells making up a final
concentration of 8 µg/ml to increase the efficiency of the transfection. Cells were
transfected with short hairpin RNA and incubated for 20 hours at 37 °C in 5% CO2.
Following the incubation, the transfected cells were selected with medium
containing puromycin, which was changed every 2-3 days. Thereafter, the cells
were casted in low gelling agarose and incubated for 10 days at 37 °C in 5% CO2
for colony formation. The colonies were isolated and cultured in complete RPMI1640 to expand the colony. The level of hMTH1 was determined (by Western blot)
in the cells from 20 transfected colonies and the colony with lowest expression level
of hMTH1 was selected for this study.
Western blot
The MTH1 expression level in transfected TK6 cells was confirmed with the
Western blot assay as described in paper I [69]. The cells were lysed with Laemmli
buffer supplemented with proteinase inhibitor cocktail tablet. The gel was immersed
in MOPS (Invitrogen) and loaded with PageRuler Plus Prestained protein ladder
and with the cell samples (5-7µg). Electrophoresis was run at 100 V for 1.5-2 hours.
To transfer the proteins from the gel to a PVDF membrane, blotting sandwich was
assembled and run overnight at 30 V. On the following day, the membrane was
incubated with 5% fat-free milk for 1.5 hours to block unspecific binding sites on the
membrane. The membrane was washed with TBS and thereafter incubated with
primary
antibody
(rabbit
anti-MTH1)
and
secondary
(anti-rabbit
IgG-HRP
conjugated) antibody. The membrane was incubated in ECL plus substrate for 2
19
minutes in the dark and visualized with Fuji CCD camera. The bands on the picture
were analyzed with the Gelpro Analyzer.
Irradiation with UV
Prior to irradiation the cells were washed with RPMI without phenol red
(RPMI w/o) to eliminate any formation of toxic compounds from phenol red during
exposure to UV light. The exposure was from above and the cells were in a petri
dish without the lid in order to not reduce the radiation fluence. The cells were
always kept on ice prior to, during and after irradiation to avoid heating by the UVA
light. The UVA source was an Osram UltraMed 400W lamp with 4.5 mm Sekurit
glass, heat protection filter and blue glass filter (figure 8). The fluence of the source
was 122 W/m2, corresponding to 122 J/sec/m2. For irradiation with UVB, a corona
mini dose UV240T lamp, 230 V 50 Hz, 70 W, with a fluence of 1.4 W/m2 was used.
Irradiation with UVC was performed using a low-pressure mercury lamp (Philips UV,
15 W) with more than 80% output at 254 nm at a fluence of 0.18 W/m2 monitored by
a radiometer (Ultra-violet products, Inc., model J-260 Digital radiometer, with a
calibrated probe).
Figure 8. The UVA source with cells prepared for irradiation. The construction of
UVB and UVC has a similar setup.
20
Clonogenic survival
The clonogenic survival of MTH1-knockdown and wild-type TK6 cells was
investigated following exposure to different doses of UVA, UVB and UVC. The
doses that were applied for UVA were 0, 36, 73, 145 and 290 kJ/m 2. The doses
applied for UVB were 0, 20, 40, 60, 80, 120, 160, and 180 J/m2. The doses applied
for UVC were 0, 1, 5, 10, 15, 20, 30, and 40 J/m2. Following irradiation a defined
number of cells (50-800 cells/well) were casted into a 6-well plate with low gelling
agarose prepared in complete RPMI. The plate was then incubated at 37 °C in 5%
CO2 for 10 days to allow colony formation of surviving cells.
Mutant frequency
A mutant frequency assay was performed in order to investigate the mutant
frequency of MTH1-knockdown and wild-type TK6 cells following UVA, UVB and
UVC irradiation. The doses used for this assay were acquired from the clonogenic
survival results. For UVA, a dose of 50% and 80% of IC50 (dose at which 50% of the
cells were killed) was used corresponding to 18 and 29 kJ/m2, respectively. For
UVB and UVC a dose of IC50 was used; 50 and 7 J/m2, respectively. For each
experiment a sham-irradiated sample was also prepared as a control for the
determination of the background mutant frequency. Following irradiation the cells
were cultured and incubated in T75 flasks with complete RPMI for 10-14 days at 37
°C in 5% CO2. Thereafter a defined number of cells (400 000 – 1000 000 cells/well)
was cast in 4 wells of a 6-well plate with low gelling agarose prepared in complete
RPMI supplemented with 5 µg/ml trifluorothymidine for selection of the mutants. The
remaining 2 wells were used for estimation of the clonogenic efficiency (cells
without any treatment). The plate was then incubated at 37 °C in 5% CO2 for 10
days for colony formation. The colonies/mutants were counted to calculate the
number of mutants per 100 000 surviving cells.
21
Measurement of intra- and extracellular 8-oxo-dG and dG following UVA
irradiation
The concentration of intra- and extracellular 8-oxo-dG were measured in
wild-type and MTH1-knockdown TK6 cells following exposure to UVA with doses of;
0, 18, 29 and 73 kJ/m2. The highest dose was only used for the measurement of
intracellular dG. The intra- and extracellular 8-oxo-dG (after conversion of 8-oxodGTP/dGDP/dGMP to 8-oxo-dG) levels were measured using ELISA while
intracellular dG (after conversion of dGTP/dGDP/dGMP to dG) levels were
measured using HPLC. The cells were centrifuged to separate pellet (intracellular)
and supernatant (extracellular). For the analysis of the intracellular 8-oxo-dG, the
pellet was mixed with ethanol/double distilled water (1:1) and incubated overnight at
4 °C on shaker to extract the low-molecular-weight intracellular components.
The samples were mixed with internal standard (80 µM BrdU), frozen at -20
°C, lyophilized and then mixed with 500 µl sterile water containing 5% Dimethyl
sulfoxide (DMSO). Following filtration to remove high-molecular-weight compounds,
40u calf intestine alkaline phosphatase (CIAP) was added using a kit supplied from
In Vitro Sweden AB. The mixture was incubated for 2 hours at 37 °C for the
dephosphorylation. The samples were divided into 2 tubes, one for measuring 8oxo-dG with ELISA and the other for measuring dG with HPLC [69].
To measure intra- and extracellular 8-oxo-dG with ELISA, an ELISA kit was
purchased from Health Biomarkers Sweden AB. The detailed method for detection
of 8-oxo-dG has been described in the first publication.
Isolation of mutants and RNA
Several mutant frequency assays were performed as described above to
isolate mutants exposed to a UVA dose of 0 and 32 kJ/m2 (IC50). Almost 50 mutant
colonies resistant to trifluorothymidine were picked up from each of the four treated
groups: MTH1-knockdown/wild-type and UVA exposed/non-exposed TK6 cells.
RNA was extracted from each mutant colony with a kit supplied from Sigma Aldrich
(GeneElute Mammalian Total RNA miniprep Kit). The cells were lysed and
thereafter centrifuged through a GenElute column to remove cellular debris and
DNA. The filtrate was mixed with an equal amount of 70% ethanol and thereafter
through another column to collect the RNA. The quality of RNA was checked by
22
agarose gel electrophoresis whereby the 28S and 18S rRNA should appear. The
RNA concentration was measured using a NanoDrop spectrophotometer (ND8000).
cDNA isolation
This part was performed in collaboration with the group of prof. Leon
Mullenders at Leiden University Medical Center, department of Toxicogenetics, the
Netherlands. The RNA was converted to cDNA using a kit from Invitrogen/Life
technologies (Cat. No: 18080-051). Firstly, 2 µg of total RNA was mixed with oligodT primers and dNTPs. The mixture was thereafter incubated at 65 °C for 5 minutes
and placed on ice for at least 1 minute. Then, First strand buffer (5x), MgCl 2 (25
mM), DTT (0.1 M), µl RNaseOUT (inhibitor) (40 U/µl) and superscript III RT (reverse
transcriptase) were added to the mixture and incubated at 50 °C for 50 minutes.
The cDNA synthesis reaction was stopped by heating the mixture to 85 °C for 5
minutes.
Sequencing
To identify the mutations induced by UVA, the Tk gene was sequenced for
each cDNA corresponding to each mutant. The primers used were called Tk-WF1,
WR1 and WNF1 & WNR1 (900 bp) (table 2). Each reaction comprised: cDNA, 4 µl
dNTPs (2.5 mM), 5x GoTaq Buffer, GoTaq, forward and reverse primers and water.
For most of the mutants, WF1 and WR1, one PCR round was enough whilst some
mutants that give weak bands needed a second round of PCR with WNF1 and
WNR1. Before sending the PCR products for sequencing, they were purified using a
QIAquick PCR Purification Kit. Chromas Version 1.5 software was used to visualize
and analyze the DNA sequence. DNAMAN software Version 5.2.9 was used to align
the mutant DNA sequence with the wild-type DNA sequence in order to determine
the mutation.
23
Table 2. Sequences of primers for thymidine kinase
Primer pairs
Length
Primer
Primer sequence
1st amplification primers
900 bp
WF1
GGAGAGTACTCGGGTTCGTG
WR1
CAGCCACACAAAGGAGAG
WNF1
GGAGAGTACTCGGGTTCGTG
WNR1
CAGCCACACAAAGGAGAG
2nd amplification primers
900 bp
Micronucleus assay
MN frequency was used as an endpoint for cytogenetic damage in MTH1knockdown and wild-type TK6 cells following exposure to UVA, UVB and UVC. The
cells were exposed to IC80 corresponding to 73 kJ/m2 for UVA, 124 J/m2 for UVB,
18 J/m2 for UVC, and 1 Gy for gamma rays. For each experiment a sham-irradiated
sample was prepared as a control for the determination of the background MN
frequency. Following irradiation, cells from each treatment were incubated in flasks
with RPMI and Cytochalasin B (final concentration 5.6 µg/ml). The cells were
incubated for 24, 30 and 46 hours at 37 °C and thereafter harvested for the MN
assay. The cells were collected into a tube and centrifuged, leaving approximately
0.5 ml of supernatant. Cellular swelling was achieved by the use of a hypotonic
solution, 0.14 M KCl and for fixation first with methanol: 0.9% NaCl: acetic acid
(12:13:3) and secondly with methanol: acetic acid (4:1). Following fixation, the cells
were dropped on a glass slide and stained with 5% Giemsa in phosphate-buffered
saline. The slides were visualized in a light microscopy with a 40× objective,
whereby the MN were then scored as described by Fenech et al. [113]. The MN
frequency and replication index was then investigated. 500 binucleated cells per
treatment were scored for MN frequency and 500 cells per treatment were scored
for the replication index.
24
Quantitative Real-Time PCR
The epigenetic effect of UVA was investigated by measuring the mRNA
levels of DNMT1 and DNMT3a in the TK6 lymphoblastoid cells. 6 × 106 cells were
irradiated with a UVA dose of 32 kJ/m2 corresponding to IC50 and were thereafter
incubated at 37 °C. For each experiment a sham-irradiated sample was also
prepared as a control for the determination of the background mRNA level. After 3
hours, 24 hours and 1 week the RNA was isolated whereby 1 µg RNA was
reversely transcribed into cDNA as described above. After several experiments the
protocol was optimized for the target genes (DNMT1 and DNMT3a) and the
reference genes (GAPDH and TBP) (table 3). The cDNA was diluted to the optimal
concentration 3.125 ng/µl prior mixing the reactions. The reactions were mixed and
transferred into a 96-well plate. 14.8 µl of cDNA was added to a total volume of 20
µl that contained 300 nM of each primer (forward and reverse), and 5 × HOT
FIREPol EvaGreen qPCR Mix Plus (Solis BioDyne Estonia). The 96-well plate was
put into the LightCycler 480. The PCR reaction was initiated by denaturing the
cDNA and activating the enzyme at 95 °C for 15 minutes, followed by amplification
for 50 cycles at 95 °C for 15 seconds, 60 °C for 1 minute and 72 °C for 20 seconds.
The data was analyzed with the LightCycler 480 SW 1.5.
Table 3. Primers used for quantitative real-time PCR.
Gene
Forward (5’>3’)
DNMT1
TACCTGGACGACCCTGACCTC
Reverse (5’>3’)
CGTTGGCATCAAAGATGGACA
Length
Reference
103 bp
[114]
DNMT3a TATTGATGAGCGCACAAGAGAGC GGGTGTTCCAGGGTAACATTGAG
111 bp
[114]
GAPDH
CAGCCTCAAGATCATCAGCA
TGTGGTCATGAGTCCTTCCA
106 bp
[115]
TBP
GATCAGAACAACAGCCTGCC
TTCTGAATAGGCTGTGGGGT
131 bp
qPrimerDepot
Statistical analysis
The student’s t-test was applied comparing the results for clonogenic survival
and for the levels of intra- and extracellular 8-oxo-dG. For the mutant frequency, the
multiple comparison (ANOVA) with the Tukey’s method as post hoc analysis was
25
Results
Paper I
Title: “Reduction of 8-oxodGTP in the nucleotide pool by MTH1 leads to
reduction in mutations in the human lymphoblastoid cell line TK6 exposed to
UVA”
The DNA has long been considered the most significant genotypic target for
exogenous and endogenous factors. DNA is however intricately packed by proteins
into nucleosomes, which offers an efficient protection. On the other hand, the
nucleotides in the nucleotide pool are free and could be more susceptible to
damage. Recent studies show that the nucleotide pool rather than the DNA is the
main target for oxidative damage [36, 112, 116]. Oxidized nucleotides in the
nucleotide pool may be incorporated into DNA and if not repaired lead to mutations.
The aim of this work was to investigate if the nucleotide pool is the main
target for UVA induced mutations. A human cellular model system; lymphoblastoids
with a knockdown of the nucleotide pool sanitization enzyme MTH1, was used.
These cells are therefore less capable of handling oxidized nucleotides in the
nucleotide pool. We investigated clonogenic survival, mutant frequency and the
intra- and extracellular 8-oxo-dG in MTH1-knockdown and wild-type TK6 cells
following exposure to UVA.
The results indicate a clear decrease in survival of both MTH1-knockdown
and wild-type TK6 cells with higher dose of UVA but do not show a significant
difference between the two groups. Significantly increased levels of mutants were
observed after UVA exposure in both MTH1-knockdown and wild-type TK6 cells
when compared with non-exposed cells. A significantly higher mutant frequency
was observed in the MTH1-knockdown TK6 cells when compared to the wild-type
TK6 cells, indicating a protective role of MTH1 towards UVA induced oxidative
damage in the nucleotide pool. The higher mutant frequency in MTH1-knockdown
TK6 cells also indicates that UVA causes nucleotide oxidation in the nucleotide
pool. When looking at the intra- and extracellular 8-oxo-dG levels, significantly
increased levels of intracellular 8-oxo-dG and significantly decreased levels of
extracellular 8-oxo-dG could be seen in MTH1-knockdown TK6 cells following
exposure to UVA, when compared to the wild-type TK6 cells (figure 10). This clearly
27
Paper II
Title: “Analysis of mutant frequencies and mutation spectra in MTH1
knockdown TK6 cells exposed to UV radiation”
As a continuation of paper I, we were also interested to investigate whether
UVB and UVC also induce oxidized nucleotide damages and in this case, if the
nucleotide pool is an important target. The same type of cells were used; MTH1knockdown and wild-type TK6 cells. We investigated clonogenic survival and
mutant frequency in the cells with a knockdown in MTH1 (MTH1-knockdown) and
cells with normal levels of MTH1 (wild-type) following exposure to UVB and UVC.
The results indicate that there is a clear decrease in cell survival with
increasing doses of UVB and UVC in both MTH1-knockdown and wild-type TK6
cells, however, there is no significant difference between the MTH1-knockdown and
wild-type TK6 cells. There was a significant increase (p<0.001) in mutant frequency
in both MTH1-knockdown and wild-type TK6 cells following exposure to UVB and
UVC when compared with non-exposed cells. There was a significant increase
(p=0.017) in mutant frequency between the MTH1-knockdown and wild-type TK6
cells when exposed to UVB but not UVC. This indicates that MTH1 has (as for UVA)
a protective role in UVB-induced mutagenesis and that the nucleotide pool is a
significant target for UVB-induced mutagenesis.
Since MTH1 has a protective role in mutagenesis originating from the
nucleotide pool, the affected nucleotides involved in the mechanism could be
oxidized guanine (8-oxo-dGTP) and/or oxidized adenine (2-OH-dATP). To
investigate the mechanism of UVA-induced damage, we also aimed to investigate
the type of genetic changes (mutations) induced in MTH1-knockdown and wild-type
TK6 cells exposed to UVA by sequencing the thymidine kinase (Tk) gene.
Firstly, a set of mutant frequency experiments was performed with MTH1knockdown/wild-type and UVA exposed/non-exposed TK6 cells. Mutants (mutated
in thymidine kinase) were isolated, RNA was extracted and converted to cDNA. The
cDNAs were sequenced in the Tk gene to find out the mutation spectra in all 4
groups (MTH1-knockdown/wild-type and exposed/non-exposed TK6 cells). The
results indicated that UVA induced mostly GC>AT and AT>GC transitions. More
specifically, GC>AT transitions dominated in wild-type TK6 cells, whereas AT>GC
transitions dominated in MTH1-knockdown TK6 cells. These mutations are
29
Paper III
Title: “Investigation of micronucleus induction in MTH1 knockdown cells
exposed to UVA, UVB and UVC”
Paper I and II have been focusing on UVR induced mutagenesis, whereas
here the focus is mainly on UVR induced cytogenetic damage.
UVR can also
induce DSBs, which is the primary cause of chromosomal aberrations and MN.
UVR induced DSBs can arise either during repair of pyrimidine dimers/oxidative
DNA damage that lay in close proximity on the DNA or by the attack of ROS directly
on the DNA.
To assess the cytogenetic damage, MN assay was used as an endpoint. We
wanted to see whether UVR induces cytogenetic damage such as MN and in that
case, whether MTH1 can protect against cytogenetic damage induced by UVR.
MTH1-knockdown and wild-type TK6 cells were exposed to UVA, UVB and UVC
and thereafter investigated for MN induction 24, 30 and 46 hours following
irradiation. Cells exposed to gamma radiation were used as a positive control since
it is already known that gamma radiation induce MN [117].
We found out that UVA is able to induce MN significantly after all three
incubation times, whereas UVB and UVC were able to induce MN significantly only
after the later incubation time. This suggests that UVB- and UVC-induced
pyrimidine dimers need more time to be transformed to DSBs and are therefore Sphase dependent whereas UVA is able to induce DSBs directly through the action
of ROS. Our results show that there is no significant difference between the MTH1knockdown and wild-type TK6 cells. This observation suggests that MTH1 has no
role in the protection against MN induction and that the MN are most probably not
induced via oxidized nucleotides (dGTP and dATP) in the nucleotide pool but from
the direct action of ROS on DNA.
Main findings:
x
UVA induces cytogenetic damage through a mechanism that is not dependent on that the
cells are cycling (proliferating).
x
UVA is suggested to induce cytogenetic damage through generation of ROS.
x
UVB and UVC induce cytogenetic damage only in cells that are cycling (proliferating).
x
UVB and UVC might induce cytogenetic damage through the formation of pyrimidine dimers.
31
Discussion
This project has been carried out using the human TK6 lymphoblastoid cell
line. This cell line was used since it is a human cell line and useful for these
mechanistic studies. Although UVR is mainly targeting the skin cells such as
keratinocytes and melanocytes, the longer wavelengths of UVR can penetrate as
far as into the dermis where blood vessels are located.
The
nucleotide
sanitization
enzyme,
MTH1,
protects
the
cells
by
dephosphorylating 8-oxo-dGTP and 2-OH-dATP to monophosphates and thereby
preventing their incorporation into DNA. Single nucleotide polymorphisms (SNPs)
have been found in the MTH1 gene, affecting the activity of the protein [60]. This
makes it interesting to investigate how different levels of MTH1 in cells may affect
the sensitivity in aspects of mutagenicity. In our present work the protective role of
MTH1 in mutagenesis and cytogenetic damage induced by UVR has been
investigated.
Our results show that MTH1 protects cells more efficiently against mutations
induced by longer wavelengths rather than shorter wavelengths of UVR (paper I
and paper II). This suggests that oxidative stress plays a greater role at longer
wavelengths of UVR. This phenomenon has been observed previously in a study by
Zhang et al. [118]. However, Zhang’s group was looking at the damaging effects
solely in the DNA, whereas in the current project, a biomarker was used to measure
oxidative damage (8-oxo-dG) both in the intracellular milieu as well as in the
extracellular milieu to be able to investigate the damaging effects in the nucleotide
pool.
The
analysis
of
intra-
and
extracellular
8-oxo-dG
was
done
by
dephosphorylating the 8-oxo-dGTP, 8-oxo-dGDP, and 8-oxo-dGMP to 8-oxo-dG.
The dephosphorylation together with the use of ethanol and DMSO during
extraction allowed us to reduce experimental errors induced by possible oxidation of
nucleotides or phosphorylation/dephosphorylation during the extraction [43]. The
results show that there is an increase of 8-oxo-dGTP in the intracellular milieu and a
decrease of 8-oxo-dG in the extracellular milieu of MHT1-knockdown TK6 cells
when compared with wild-type TK6 cells. These findings show that MTH1 has antimutagenic properties by reducing the incorporation of 8-oxo-dG into DNA following
exposure to UVA, which supports the idea that the nucleotide pool is a significant
32
target for UVA. The specific mutation spectrum following exposure to UVA in the Tk
gene shows that the major types of mutational events are GC>AT and AT>GC
transitions (paper II). These mutations are suggested to be induced by 2-OH-dATP,
which is prone to mispair with a C or T during replication (figure 7). We therefore
conclude that the nucleotide pool is a significant target for mutations induced by
UVA. Even though most studies have focused on the role of 8-oxo-dG, this study
shows that 2-OH-dATP should also be taken into account.
The same type of mutations, in the Tk gene, was found (paper II) in both
MTH1-knockdown and wild-type TK6 cells with and without exposure to UVA, i.e.
mainly GC>AT and AT>GC transitions. These results indicate that UVA enhanced
the levels of the type of mutations that we observed in the non-exposed cells.
Therefore, the mutagenic effect of UVA could be a consequence of an increased
endogenous production of ROS, as has been shown to be the case after exposure
to low doses of ionizing radiation [112].
A similar trend as for UVA is observed for UVB irradiation in terms of a
significantly higher mutant frequency in MTH1-knockdown TK6 cells when
compared to wild-type TK6 cells. We therefore suggest that the nucleotide pool also
is a main target for mutations induced by UVB. As no significant difference was
seen in mutant frequencies between MTH1-knockdown and wild-type TK6 cells
following exposure to UVC, one can conclude that the mutagenesis after UVC
exposure is caused mainly through formation of photoproducts, which cannot be
protected by the MTH1 protein.
Our results indicate that the nucleotide pool is a significant target for
mutagenesis. We could protect our cellular nucleotide pool by minimizing the level
of ROS and oxidized nucleotides. This can be done by the action of antioxidants
and repair proteins such as MTH1. Antioxidants such as SOD, glutathione
peroxidase are the body’s own defense mechanism. Antioxidants can also be taken
up from the food such as fruits and vegetables (vitamin C, vitamin E), tea (flavonoid)
and tomato juice (lycopene) [31, 119]. If the level of ROS is not controlled by the
antioxidants, this could lead to photoaging and cancer [120, 121]. Interestingly, the
same type of mutation that we observed after UVA exposure, GC>AT, have been
found to be the predominant mutation in multiple melanoma cell lines [122]. This
suggests that UVA is part of the factors responsible for malign melanoma formation.
There are however contradicting results on this matter, argued by Moan et al. [123].
33
Nonmelanomas (BCC and SCC) strongly seem to be related to UVR exposure and
is mostly formed on the face which is the mostly exposed area of the body. In
contrast, CMM cases have shown larger formation on the trunk, which is not directly
exposed to UVR [123].
Other UVR-induced damage than mutations such as cytogenetic damage
(MN) have also been observed in this study (Paper III). MN is induced with an
earlier response to UVA exposure compared to UVB and UVC exposure. This is
suggested to be due to the different types of damage induced by the different
wavelengths of UVR. Our data indicate that mutations from photoproducts (such as
CPD and 6-4PP) are S-phase dependent lesions, while ROS are able to induce
DSBs independently of the cell cycle stage. In this case, UVA is suggested to
induce MN by the direct action of ROS while UVB and UVC might induce MN
through generation of photoproducts that are transformed into DBSs during DNA
replication. Interestingly, MTH1 has no protective role against the MN induction by
UVA, UVB or UVC. This implies, as already suggested, that these cytogenetic
events are induced by factors other than UVR-induced oxidized nucleotides in the
nucleotide pool.
Our conclusion is that MTH1 is an important protein for protection against
mutagenesis following exposure to longer wavelengths of UVR. Given that
polymorphism in the MTH1 gene gives rise to MTH1 protein with different levels of
activity, MTH1 may be one possible genetic factor that relate to the mechanisms
behind individual sensitivity to develop skin cancer. We have also shown that the
nucleotide pool is the main target in mutagenesis of the longer wavelengths of UVR.
More specifically, the major types of mutation induced by UVA are GC>AT and
AT>GC suggested to arise from 2-OH-dATP.
34
Ongoing studies and future perspective
We have shown that the nucleotide pool is an important target for longer
wavelengths of UVR in inducing mutagenesis and for a comprehensive
understanding of the mechanisms behind, further research will be needed. We
suggest that the mutations induced by UVA (GC>AT and AT>GC) arise from 2-OHdATP. In order to verify this hypothesis, intra- and extracellular 2-OH-dATP in
response to UVA in our TK6 cell model system should be analyzed. According to
our results, UVC does not induce oxidative nucleotide damage that contribute to the
mutagenicity whereas we have shown that UVB does but to a lower degree than
UVA. Sequencing the Tk gene in mutants and measuring 8-oxo-dGTP and 2-OHdATP in cells exposed to UVB and UVC would also provide us with better
knowledge of UVR damage at short wavelengths.
Epigenetics is suggested to have a high impact on the changes in gene
expression needed for the development of malignant melanoma and other cancers
[82]. Up to now, very few studies have been done on the epigenetic effects of UVR.
In a recent study the effect of UVB on the global DNA methylation and DNMT1
mRNA expression were investigated in healthy subjects and patients with the
autoimmune inflammatory disease, systemic lupus erythematosus (SLE). Most of
these patients are photosensitive. An inhibitory effect of UVB on both global DNA
methylation and DNMT1 mRNA expression levels were observed resulting in hypomethylation. The mechanism by which UVB induces hypo-methylation however is
unclear and further studies are required [124].
We have in our lab established a protocol for analysis of DNMT1 and
DNMT3a at the mRNA level using quantitative real-time PCR with the LightCycler
480. The protocol is adopted for the human lymphoblastoid TK6 cells. The aim is to
investigate if there is any epigenetic effect in TK6 cells when exposed to UVA.
Although the experimental variation was large at 1 week after exposure there was a
tendency of an mRNA increase for both DNMT1 and DNMT3a (data not shown),
however additional experiments must be done to complement the data. It would be
of interest to perform experiments with longer incubation times to see whether or
not the mRNA levels remain increased and if the changes in mRNA levels reflect in
changes of protein levels.
35
Acknowledgements
I would first like to thank Dr. Siamak Haghdoost and Prof. Mats Harms-Ringdahl
for accepting me as an intern during summer 2008, when it all began.
Dr. Siamak Haghdoost, thank you for being available for discussions and help in
the laboratory. It has been great being a part of your research. I appreciate your
patience in teaching and educating me for improving my skills in science.
Having Prof. Andrzej Wojcik as a co-supervisor have been a privilege. I never
needed Google around you with all the knowledge that you have. Thank you for all
the theoretical and practical support.
Prof. Mats Harms-ringdahl, you are the pride of the CRPR-group. You are not only
an outstanding professor but also a wonderful human being with a lot of kindness.
Your presence always spreads a smile.
I would also like to thank my co-supervisor Dr. Natalia Kotova who always had
good suggestions and discussions.
The group of CRPR have brought joy to the daily routines of a PhD-student. Without
you the days would pass slower. Thanks to all of you:
Sara Shakeri Manesh: I am so glad that I had you by my side during my whole
PhD project, both at work and after work. You made me not to lose hope and
continue in hopeless times of a PhD-career. It has been a pleasure to work with you
and I hope that our roads will cross once again in the future.
Marina: My friend from far behind! All the fun we had as lab partners in 2005 at
Molecular biology started a lifetime of friendship. With you joining CRPR the fun
continued. I miss working with you!
Eliana: It has been great to have you here in Sweden at CRPR. You became a very
good colleague and a friend to me. Thank you for all the time we spent at work and
after work.
Traimate: You are the expert in western blot and have always been a big help in
the laboratory. What would the lab be without you? Thank you so much for all the
help and for being such a nice person.
Siv: Thank you for all the help with your excellent correction skills. I could always
count on you.
Marta: It still feels empty since you left! It has been a pleasure getting help with
micronucleus scoring from a girl with chromosome aberrations expertise.
Sara Skiöld: Thank you for always being so kind and for the nice time together at
work and travelling. My suitcase always felt heavier in comparison ;)
Alice: It has been fun having you as a colleague. Thank you for sharing your expert
knowledge in gamma-H2AX and tiramisu with me.
Ainars: It has been great working in the same group with you. You were the most
organized person in the group.
Elina: It has been nice to have you as a colleague and to share the same office with
you. I am very thankful for your advices during my PhD-project.
Karl: Putting me and you in the same office made us realize that even people with
different music taste can have fun and come along just fine.
36
Lovisa: Welcome to CRPR and I know you will have a great time here!
Also a big thanks to the other colleagues at the MBW. You all contributed to the
warmth atmosphere at MBW.
A special thanks to my former supervisor at Helmholtz institute, Dr. Michael
Rosemann. Thank you for your expert advices in quantitative real-time PCR. You
have always been very kind and helpful.
I would like to thank our collaborators at the Department of Toxicogenetics and
Dermatology of Leiden University Medical Center. Thank you Dr. Jacob G. Jansen
for your quick answers and helpful discussions. Thank you Winta Woldai Hagos for
your excellent lab work with analyzing the Tk gene. I am thankful to prof. Leon
Mullenders for making this collaboration possible.
Thank you my dearest family and friends for seeing me as a doctor from the
moment I got accepted to this PhD-project.
Thank you grandmother, Shahnaz (mami), for always taking care of us and always
believing in me. You really have a heart of gold.
Thank you my dear brother Afra for always believing in me. Spending time with you
always make me forget about any struggle in life, almost like going back in time to
our childhood.
My father, Dr. Farzin, I am very proud of you and I thank you for all your support.
Your texts at Monday mornings have always been a good start of the week.
Shafagh, thank you for all your support. I am so glad to have you in my life. You
really are a super-woman always helping out and spreading joy to everyone around
you.
A special thanks goes to Homan. Having you in my life at the end of my PhDproject gave me the extra strength that I needed. Thank you for believing in me and
for all support from both you and your loving family; Farnoosh, Hooshang,
mamani, Ahmad, Dina, Johan, Pegah, Reza, and Elara.
I would also like to thank Fereshteh, Arman, Marjan, Ashkan, Azadeh and
Melody for being there for me.
Last but not least, the person that influenced me to do a PhD-project, my mother
Afsaneh. You have always been there for me, not only as a mother but as a friend
too. I always have you to thank for everything that I have in my life.
37
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