Oxidatively Induced DNA Damage and Its Repair in Cancer

Miral Dizdaroglu

Biomolecular Measurement Division, National Institute of Standards and Technology, 100

Bureau Drive, MS 8311, Gaithersburg, MD 20899, USA

Corresponding author. Tel.: +1-301-975-2581; fax: +1-301-975-8505

1 ABSTRACT

Oxidatively induced DNA damage is caused in living organisms by endogenous and exogenous reactive species. DNA lesions resulting from this type of damage are mutagenic and cytotoxic and, if not repaired, can cause genetic instability that may lead to disease processes including carcinogenesis. Living organisms possess DNA repair mechanisms that include a variety of pathways to repair multiple DNA lesions.

Mutations and polymorphisms also occur in DNA repair genes adversely affecting

DNA repair systems. Cancer tissues overexpress DNA repair proteins and thus develop

greater DNA repair capacity than normal tissues. Increased DNA repair in tumors that

removes DNA lesions before they become toxic is a major mechanism for development

of resistance to therapy, affecting patient survival. Accumulated evidence suggests that

DNA repair capacity may be a predictive biomarker for patient response to therapy.

Thus, knowledge of DNA protein expressions in normal and cancerous tissues may

help predict and guide development of treatments and yield the best therapeutic

response. DNA repair proteins constitute targets for inhibitors to overcome the

resistance of tumors to therapy. Inhibitors of DNA repair for combination therapy or as

single agents for monotherapy may help selectively kill tumors, potentially leading to

personalized therapy. Numerous inhibitors have been developed and are being tested in

clinical trials. The efficacy of some inhibitors in therapy has been demonstrated in

patients. More developments of inhibitors of DNA repair proteins are globally

underway to help eradicate cancer.

Keywords:

Cancer therapy

DNA damage

DNA repair

DNA glycosylases

Inhibitors

Contents

1. Introduction

2. Mechanistic aspects of oxidatively induced DNA damage

2.1. Purines

2.2. Pyrimidines

2.3. Sugar moiety

2.4. Tandem lesions

2.4.1. 8,5'-Cyclopurine-2'-deoxynucleosides

2.4.2. Base-base tandem lesions

2.4.3. DNA-protein cross-links

2.4.4. Clustered lesions

3. Cellular repair of oxidatively induced DNA lesions

3.1. Base excision repair

3.1.1. Substrate specificities of prokaryotic DNA glycosylases

3.1.2. Substrate specificities of eukaryotic DNA glycosylases

3.1.3. Repair of sugar lesions

4. Genetic effects of oxidatively induced DNA lesions

4.1. Purine-derived lesions

4.2. Pyrimidine-derived lesions

4.3. 8,5'-Cyclopurine-2'-deoxynucleosides

4.4. Sugar lesions

5. Oxidatively induced DNA damage and cancer

5.1. Role of DNA glycosylases of BER in carcinogenesis

5.1.1. OGG1

5.1.2. NEIL proteins

5.1.3. NTH1

5.2. Other BER proteins

5.2.1. APE1

5.2.2. Pol β

5.3. DNA lesions and DNA repair proteins as biomarkers

5.3.1. DNA lesions as biomarkers

5.3.2. BER proteins as biomarkers

5.3.3. BER proteins as therapy targets

6. Conclusions

References

● ●─ ─ Abbreviations: RS, reactive species; OH, hydroxyl radical; O2 , superoxide radical; eaq ,

hydrated electron; k, reaction rate constant; 8-OH-Gua, 8-hydroxyguanine; FapyGua, 2,6- diamino-4-hydroxy-5-formamidopyrimidine; 8-OH-Ade, 8-hydroxyadenine; FapyAde, 4,6- diamino-5-formamidopyrimidine; Sp, spiroiminohydantoin; Gh, 5-guanidinohydantoin; 5-

OHMe-Ura, 5-(hydroxymethyl)uracil; 5-OH-Cyt, 5-hydroxycytosine; 5-OH-Ura, 5- hydroxyuracil; cdA, 8,5'-cyclo-2'-deoxyadenosine; cdG, 8,5'-cyclo-2'-deoxyguanosine; Fo, formamido residue; Thy-Tyr cross-link, 3-[1,3-dihydro-2,4-dioxopyrimidin-5-yl)-methyl]-L- tyrosine; BER, base excision repair; NER, nucleotide excision repair; MMR, mismatch repair; AP site, apyrimidinic/apurinic site; Me-FapyGua, 2,6-diamino-4-hydroxy-N7-methyl-

5-formamidopyrimidine; APE1, apurinic/apyrimidinic endonuclease 1; dRP, 2'-deoxyribose phosphate; Pol β; DNA polymerase β; TS, thymidylate synthetase.

1. Introduction

Reactive species (RS) including free radicals derived from either oxygen or nitrogen are generated in aerobic organisms by cellular metabolism and by exogenous sources such as ionizing radiations, UV radiation, redox cycling drugs, carcinogenic compounds, environmental toxins, etc. [1]. Antioxidant defense mechanisms exist in living organisms to encounter the production and effects of RS. If the prooxidant-antioxidant balance is disturbed in favor of the former, a state of oxidative stress can occur, leading to oxidative damage to biomolecules including DNA, proteins and lipids [1,2]. Consequences of oxidative stress can

be manyfold depending on its severity and the cell type [1]. Among others, these may include

increased genetic instability, proliferation, reduction of antioxidants, cell death, apoptosis and

angiogenesis [1]. Oxidative stress can also drive the onset of inflammation, which produces

RS and is a hallmark of cancer, predisposing individuals to different types of cancers [3-5].

The acute inflammatory response recruits neutrophils that can damage DNA [4,6]. Reactive

species are involved in carcinogenesis by damaging DNA and by modulating certain cellular

pathways[1,4]. These species can be radicals or non-radicals. Among the oxygen-derived

radicals, the hydroxyl radical (●OH) is the most reactive one and reacts with biological

molecules such as DNA constituents at or near diffusion-controlled rates [7]. Other radicals

●─ ● ● such as superoxide radical (O2 ), hydroperoxyl radical (HO2 ), peroxyl radicals (RO2 ),

● 1 + alkoxyl radicals (RO ) and singlet oxygen (O2 Σg ) possess very low or intermediate

reactivity. Non-radical H2O2 is not reactive, unless its reaction with transition metal ions

converts it into ●OH [1]. Nitric oxide (NO●) is also a free radical and possesses low

●─ reactivity; however, its reaction with O2 is diffusion-controlled, yielding peroxynitrite

(ONOO─) [8]. Peroxynitrite is a fairly unreactive non-radical. On the other hand, its

protonated form peroxynitrous acid (ONOOH) can undergo homolytic fission to yield ●OH

● and NO2 , although this reaction may not be favored [1]. Ionizing radiations also generate

● ● ─ OH and, in addition, H atom (H ) (also a free radical) and hydrated electron (eaq ) from

cellular water [9]. Reactions of these endogenously and exogenously generated species with

the DNA bases and sugar moiety result in the formation of a multitude of modifications

(reviewed in [9,10]). This type of damage, which is called oxidatively induced DNA damage, can be repaired in living cells by a variety of repair mechanisms [11]. Oxidatively induced

DNA modifications that escape repair before replication may lead to mutagenesis, which is

well known to be a fundamental part of the molecular basis of all cancers [11-13]. Mutations

occur throughout the genome, including in genes that maintain genetic stability, leading to

genetic instability, which is a hallmark of cancer [3,14,15]. Genetic instability may affect

many types of enzymes in various pathways including DNA repair [11]. In healthy cells,

there is a balance between DNA damage and DNA repair. In cancer cells, however, this

balance may be disturbed in favor of DNA damage, overwhelming DNA repair capacity of

cells and thus resulting in mutations at high frequency. Increase in DNA repair capacity may

also occur in cancer cells, causing therapy resistance [16-19]. There is mounting evidence

that oxidatively induced DNA damage by endogenous and exogenous sources may be a

significant source of mutations and genomic instability, and thus an important contributor to

carcinogenesis [11,20-22].

2. Mechanistic aspects of oxidatively induced DNA damage

2.1. Purines

Mechanistic aspects of oxidatively induced DNA damage has extensively been reviewed

in the past and just recently [9,10,23]. Thus, only a brief summary of this field will be given

here. Of the RS, ●OH is the most damaging species to DNA and other biological molecules.

Its reactions by addition to the double bonds of purines and pyrimidines in DNA are

diffusion-controlled with second-order reaction rate constants (k) of 4‒9 x 109 dm3 mol‒1 s‒1

[7,9]. Abstraction of H● from the five C-atoms of the sugar moiety and from the methyl group of thymine also occurs, albeit by slower rates with k ≈ 2 x 109 M‒1 s‒1 [7,9]. Ionizing

─ radiation-generated eaq also adds to the double bonds of DNA bases at diffusion-controlled

rates (k = 0.9‒1.7 x 109 dm3 mol‒1 s‒1); however, the addition reactions of H● are slower, but

8 3 ‒1 ‒1 ─ ● still occur at appreciable rates (k = 1‒5 x 10 dm mol s ) [23-27]. Reactions of eaq and H

with the sugar moiety of DNA are negligible. Hydroxyl radical preferentially adds to the sites

of double bonds of purines and pyrimidines with the highest electron density because of its

electrophilic nature. Addition to guanine generates C4-OH‒, C5-OH‒ and C8-OH‒adduct

● ● radicals [10,23,28]. In addition, an H abstraction by OH from the NH2 group at C2 of

guanine has been claimed to occur at a rate of ≈ 65%, practically eliminating the addition of

●OH to C4 [29,30]. However, theoretical and experimental studies unequivocally showed that this reaction is energetically not favored and does not occur to an appreciable extent [9,10,31-

36]. Addition reactions of ●OH with adenine mainly produces C4-OH‒ and C8-OH‒adduct radicals, although the C5-OH‒adduct radical is also formed, but to a much lesser extent

[23,37,38]. The addition of ●OH to the C2 of adenine also occurs to an extent of 2%. The C4-

OH‒ and C5-OH‒adduct radicals of guanine and adenine dehydrate to yield Gua(‒H)● and

Ade(‒H)● radicals, respectively, that may be reduced to reconstitute Gua and Ade [23,39].

Gua(‒H)● protonates to give rise to guanine radical cation (Gua●+), which can be converted

into the C8-OH‒adduct radical upon hydration (HO‒ addition) [40-42]. Direct effect of

ionizing radiation also generates Gua●+; therefore, the direct effect and the indirect effect of

ionizing radiation may lead to the same products of guanine [43,44]. Ade(‒H)● may also

protonate to give Ade●+, which may yield the C8-OH‒adduct radical upon hydration. This is

the same adduct radical formed by direct addition of ●OH to the C8 of adenine. The addition of ●OH to the C2 of adenine also occurs to an extent of 2%.

The major products of guanine in DNA result from the reactions of the C8-OH‒adduct radical, one-electron oxidation of which gives rise to 8-hydroxyguanine (8-OH-Gua) [45-50].

In an exothermic reaction, 8-OH-Gua tautomerizes into 8-oxoguanine, which is its

predominant keto form [51-53]. In contrast to oxidation, the C8-OH‒adduct radical of

guanine can undergo a β-fragmentation leading to unimolecular opening of the imidazole ring

(k = 2 x 105 s‒1) [23,39,54,55], followed by one-electron reduction to yield 2,6-diamino-4- hydroxy-5-formamidopyrimidine (FapyGua) (reviewed in [10,23]). The formation of 8-OH-

Gua increases in the presence of O2, whereas ring-opening leading to FapyGua is favored at low O2 concentrations and can compete with the bimolecular oxidation. Under hypoxic conditions of the cell nucleus, therefore, the ring-opening of the C8-OH‒adduct radical followed by reduction may be a favorable reaction. This notion is strongly supported by the formation of FapyGua in living cells with comparable yields to that of 8-OH-Gua (reviewed in [56]). Adenine undergoes analogous reactions, generating 8-hydroxyadenine (8-OH-Ade) and 4,6-diamino-5-formamidopyrimidine (FapyAde) (reviewed in [56]. Although called

pyrimidines, FapyGua and FapyAde are distinguished from pyrimidines by the position of

their glycosidic bond attached to the sugar moiety in DNA through the amino group at C6 of

the pyrimidine ring. It should be emphasized that, chemically and mechanistically, and also in

terms of biological effects, these formamidopyrimidines are different from their methylated

counterparts (reviewed in [56]). The oxidation of the C2-OH‒adduct radical of adenine

results in the formation of 2-hydroxyadenine [57,58].

Oxygen reacts with the OH‒adduct radicals of guanine and adenine at different rates. The

6 3 C4-OH‒adduct radical of guanine does not react with O2 at appreciable rates (k ≤ 10 dm

‒1 ‒1 9 mol s ); however, O2 rapidly reacts with the C4-OH‒adduct radical of adenine (k = 1 x 10

3 ‒1 ‒1 dm mol s )[38]. On the other hand, the reactions of O2 with the C8-OH‒adduct radicals of

guanine and adenine are diffusion-controlled (k = 4 x 109 dm3 mol‒1 s‒1) [38,39]. This is

10 likely to be the reason for the preferred formation of 8-OH-Gua and 8-OH-Ade in the

presence of oxygen. The ring-opening leading to formamidopyrimidines, however, may

compete with this reaction and thus be equally efficient at the hypoxic conditions of the cell

nucleus. The abundant formation of formamidopyrimidines in vivo supports this notion

● (reviewed in [10]). The reaction of O2 with Gua(‒H) has been reported to lead to the

formation of 2,5-diamino-4H-imidazol-4-one and 2,2,4-triamino-5(2H)-oxazolone [59,60].

●─ ● However, this mechanism has not been confirmed. Instead, the addition of O2 to Gua(–H) has been shown to be kinetically more favored reaction in nucleosides and DNA (k = 3–4.7 x

109 dm3 mol‒1 s‒1) [39,61,62]. This reaction generates guanine hydroperoxide that undergoes several reactions to give rise to 2,5-diamino-4H-imidazol-4-one, the slow hydrolysis of which results in the formation of 2,2,4-triamino-5(2H)-oxazolone [59,62-64].

Because of its low reduction potential (0.74 V) compared to that of guanine (1.29 V), 8-

OH-Gua is even more prone to oxidation than guanine by a number of oxidizing agents

2─ including ionizing radiation, metal ions, peroxynitrate and IrCl6 [65]. Its oxidation gives

rise to 8-OH-Gua●+, which readily hydrates (HO─ addition) and produces the 5-OH–adduct

radical of 8-OH-Gua. One-electron oxidation of this radical leads to 5-OH-8-OH-Gua, the

isomerization (acyl shift) of which gives rise to spiroiminohydantoin (Sp) and 5-

guanidinohydantoin (Gh) by loss of CO2 depending on reaction conditions [66-72]. Cadet et

al. have misassigned the structure of spiroiminohydantoin as 4,8-dihydro-4-hydroxy-8- oxoguanine for almost two decades and have used it for a marker of single oxygen-induced damage to Gua [73-77]. Later on, the correct structure of this compound as spiroiminohydantoin has been elucidated using the synthesized authentic material and a number of analytical techniques [66-70]. Single oxygen also reacts with 8-OH-Gua to give rise to oxaluric acid and parabanic acid among other products [78,79]. Moreover, the reaction

●─ ●+ ● 9 3 ‒1 of O2 with 8-OH-Gua and its deprotonated form 8-OH-Gua(‒H ) (k = 3 x 10 dm mol

11 s‒1) yields 5-hydroperoxide of 8-OH-Gua whose facile decomposition leads to the formation

of oxaluric acid and parabanic acid [80]. This area has extensively been reviewed elsewhere

[72,81]. All these data unequivocally show the possible effect of numerous factors on 8-OH-

Gua leading to its decomposition, and thus on its measured level in DNA. It should be

pointed out that this mounting evidence stands in stark contrast to the claim by the European

Standards Committee on Oxidative DNA Damage (ESCODD) about the so-called “correct” or

“established” value of the background level of 8-OH-Gua in living organisms and about the

validity of its related recommendations to editors and reviewers of manuscripts not to accept

manuscript with levels of 8-OH-Gua above the “established” value [82-84].

─ Ionizing radiation-generated eaq reacts with guanine and adenine at diffusion-controlled

rates, giving rise to radical anions Gua●─ and Ade●─, respectively (k = 3.3‒6 x 109 dm3 mol‒1

s‒1) [25-27]. These radical anions rapidly protonate in reaction with water, generating Gua(‒

H●) and Ade(‒H●), which subsequently yield the C8-H‒adduct radicals of guanine and

adenine, respectively, by water-assisted tautomerization [23,26,27,85-87]. Addition of H● at

C8 of guanine and adenine also generates the C8-H‒adduct radicals [26]. No products of

these adduct radicals have been identified in DNA. This is likely due to electron transfer from

these radicals to other DNA bases prior to formation of final products [87].

2.2. Pyrimidines

Hydroxyl radical reacts with thymine and cytosine at diffusion control rates (k = 6.4‒6.8 x

109 dm3 mol‒1 s‒1) by addition to the C5‒C6 double bonds producing C5-OH‒ and C6-OH‒

adduct radicals [25,88]. Abstraction of H● from the methyl group of thymine also occurs,

albeit to a lesser extent, giving rise to an allyl radical [89,90]. Addition of ●OH occurs more

at C5 than that at C6 because of the higher electron density at C5. Thymine and cytosine

radicals are oxidized or reduced depending on their redox properties, experimental conditions

12 and the presence or absence of oxygen, producing a variety of products (reviewed in [9,10]).

In the absence of oxygen, the C5-OH‒ and C6-OH‒adduct radicals of thymine and cytosine

undergo oxidation and reaction with water (HO─ addition) to yield thymine glycol (Thy

glycol) and cytosine glycol, respectively. 5-(Hydroxymethyl)uracil (5-OHMe-Ura) is formed by oxidation of the allyl radical of thymine followed by HO─ addition. The adduct radicals

are also reduced, producing 5-hydroxy-6-hydrothymine, 6-hydroxy-5-hydrothymine and 5-

hydroxy-6-hydrocytosine. Oxygen adds to the thymine and cytosine radicals at diffusion-

controlled rates generating peroxyl radicals (k ≈ 2 x 109 dm3 mol‒1 s‒1). 5-Hydroxy-6- hydrothymine, 6-hydroxy-5-hydrothymine and 5-hydroxy-6-hydrocytosine are thus not formed under oxygenated conditions. Further reactions of peroxyl radicals yield thymine glycol, 5-hydroxy-5-methylhydantoin, 5-OHMe-Ura, 5-formyluracil, cytosine glycol, dialuric acid, alloxan, 5-hydroxyhydantoin, isodialuric acid and 5,6-dihydroxycytosine. Cytosine products undergo dehydration and deamination to yield 5-hydroxycytosine (5-OH-Cyt), uracil glycol, 5-hydroxyuracil (5-OH-Ura) and 5-hydroxy-6-hydrouracil [91]. Hydrated electron reacts with thymine and cytosine by addition to 5,6-double bonds (k = 1.3‒1.8 x 1010

dm3 mol‒1 s‒1) giving rise to electron adducts (anion radicals). Addition of H● also takes

place, albeit at lower rates (k ≈ 1‒1.8 x 108 dm3 mol‒1 s‒1) and generates H-adduct radicals.

Protonation of anion radicals also produce H-adduct radicals, which are converted to 5,6- dihydrothymine and 5,6-dihydrocytosine by reduction. The deamination of the latter yields

5,6-dihydrouracil. In the presence of oxygen, these products are not formed because of

─ ● diffusion-controlled reactions of O2 with eaq and H , preventing the addition reactions of

these two species. However, 5,6-dihydropyrimidines may be formed in DNA in vivo because

of the hypoxic conditions of the cell nucleus.

Fig. 1 illustrates the main products of DNA bases. Many of these products were identified

in DNA in vitro, in cultured mammalian cells, and in animal and human tissues (reviewed in

[9,10,22,92]). Their types and yields depend on experimental conditions, the presence or absence of O2, cellular redox environment, disease conditions, DNA repair capacity,

scavenger concentration, among others.

2.3. Sugar moiety

Hydroxyl radical abstracts H● from all five carbons of 2'-deoxyribose in DNA (k = 2.5 x

109 dm3 mol‒1 s‒1) in the order of H5' > H4' > H3' ≈ H2' ≈ H1', leading to C-centered radicals

[9,25,93-95]. The order of abstraction follows the exposure to solvent with the C4'- and the

C5'-positions being the most accessible to solvent and from the minor groove. Reactions of

─ ● ● eaq and H are negligible. The extent of OH attack on 2'-deoxyribose in DNA may amount to ≈ 20%, although this ratio may depend on the cellular environment[9]. Further reactions of

the C-centered radicals of 2'-deoxyribose in the presence or absence of O2 cause DNA strand

breaks and release of intact DNA bases, and generate products that are either freed from

DNA or remained within DNA or are bound to DNA as end groups of broken DNA strands.

In 1970s, the oxidatively induced products of 2'-deoxyribose in DNA have been identified and reaction mechanisms of product formation and DNA strand breaks have been elucidated

[96-101]. The reactions resulting from the C4' radical in the absence of O2, leading to strand

breaks and products, were the first understood mechanistically [9,96]. These reactions still

remain the most-widely studied mechanism of product formation and strand breakage in

DNA. Fig. 2 illustrates the products of 2'-deoxyribose in DNA. Extensive reviews of the

mechanisms and product formation can be found elsewhere [9,10,102,103].

2.4. Tandem lesions

2.4.1. 8,5'-Cyclopurine-2'-deoxynucleosides

The H● abstraction by ●OH from C5' of 2'-deoxyribose causes the formation of the tandem

lesions 8,5'-cyclopurine-2'-deoxynucleosides in DNA. The stereospecific attack of the C5'-

14 centered radical at the C8 of purine nucleosides leads to C5'‒C8-intramolecular cyclization and an N-centered purine radical. The oxidation of this radical causes the formation of (5'R)-

and (5'S)-8,5'-cyclopurine-2'-deoxynucleosides. This reaction has been first discovered to

take place in adenosine-5'-monophosphate [104]. Subsequent studies showed that this

reaction also occurs in DNA generating both (5'R)- and (5'S)-diastereomers of 8,5'-cyclo-2'-

deoxyadenosine (R-cdA and S-cdA) and 8,5'-cyclo-2'-deoxyguanosine (R-cdG and S-cdG)

(reviewed in [105]). Fig. 1 illustrates the structures of these compounds. The C5'‒C8-

intramolecular cyclization is inhibited by O2 because of its rapid reaction with C-centered

9 3 ‒1 ‒1 radicals (k = 1.9 x 10 dm mol s ) [9,105,106]. At low O2 concentrations, however, 8,5'-

cyclopurine-2'-deoxynucleosides are formed, suggesting that a competition takes place

between the C5'‒C8-intramolecular cyclization and the reaction of O2 with the C5'-radical

[105,107]. This competition may also occur in living cells because of hypoxic conditions in the cell nucleus and steric hindrances. Indeed, R-cdA, S-cdA, R-cdG and S-cdG have been observed in cultured mammalian cells, and in human and animal tissues at background levels or at elevated levels depending on disease states, aging, DNA repair deficiency, gene knockouts, environmental pollutants or exposure to ionizing radiation [108-126]. R-cdA and S-cdA have also been detected in human urine [127]. Moreover, these compounds have been found in urine of atherosclerosis patients at significantly greater concentrations than in that of healthy individuals [128]. These findings suggested that R-cdA and S-cdA combined with the noninvasive nature of urine collection may be used as potential disease biomarkers for basic research, and for clinical and epidemiological studies. The identification of the 8,5'- cyclopurine-2'-deoxynucleosides in human and animal tissues at background levels or after exposure to DNA-damaging agents, and in human urine is in stark contrast to a claim that the

“estimated” levels of these compounds in vivo were too low; therefore, they would not be detectable by any means, unless very high and biologically irrelevant ionizing radiation doses

15 were used [92,129-131]. It should be emphasized again that the background levels of R-cdA and S-cdA (or R-cdG and S-cdG) in cells could not be measured and, interestingly, were only estimated, without providing any data to support this claim. Moreover, human monocytes in culture and biologically irrelevant high radiation doses have been used only, and no data on human or animal tissues have been provided unlike the studies cited above. These facts make the aforementioned claim baseless and unscientific.

2.4.2. Base-base tandem lesions

Adjacent, intrastrand and interstrand base-base tandem lesions have been identified mostly in vitro in oligodeoxynucleotides and DNA upon exposure to ionizing radiation or to other

●OH-generating agents. It is out of the scope of this paper to review all the work done in this field and cite all the references. Briefly, identified adjacent tandem lesions in

oligodeoxynucleotides and DNA consisted of an 8-OH-Gua residue and a formamido residue

(Fo) as 8-OH-Gua/Fo or Fo/8-OH-Gua [132-143]. A mechanism has been proposed, which

involves the one-electron oxidation of a neighboring Gua by the C5-OH-C6-peroxyl radical of Thy followed by hydration of Gua●+ and oxidation to form 8-OH-Gua and the decomposition of the C5-OH-C6-oxyl radical of Thy yielding Fo [141]. However, the

significantly lower reduction potential of a peroxyl radical than that of Gua renders this

reaction endothermic. Thus, this mechanism has been dismissed as a very unlikely one [9].

These tandem lesions (8-OH-Gua/Fo or Fo/8-OH-Gua) have not yet been identified in

cellular DNA [92]. Intrastrand cross-links between C8 of guanine or adenine and the methyl

group of thymine have been observed as Gua[8,5-Me]Thy and Thy[5-Me,8]Gua [138,144-

149], or Ade[8,5-Me]Thy and Thy[5-Me,8]Ade [145,150]. Intrastrand cross-linking between guanine and cytosine (Gua[8,5]Cyt), guanine and 5-methylcytosine (Gua[8,5-Me]MeCyt), and guanine and thymine (Gua[8,N3]Thy) have also been reported [151-154]. An interstrand cross-link has been shown to occur between the amino group of adenine on one DNA strand

16 and the allyl radical of thymine on the other DNA strand [155-158]. In studies in vivo, however, only Gua[8,5-Me]Thy and Gua[8,5]Cyt have been identified in γ-irradiated cultured cells and in animal tissues [151,159-161]. Fig. 3 illustrates the structures of Gua[8,5-Me]Thy and Gua[8,5]Cyt. Extensive reviews including reaction mechanisms of this field can be found elsewhere [10,92,149].

2.4.3. DNA-protein cross-links

Hydroxyl radical reactions with DNA bases and proteins in chromatin, which generate

DNA base radicals and amino acid radicals, cause the formation of covalent DNA-protein cross-linking [162-168]. Mechanisms of cross-linking may involve the reaction between a

DNA base radical and an amino acid or an amino acid radical and a DNA base or a DNA base radical and an amino acid radical. For example, a Thy-Tyr cross-link (3-[1,3-dihydro-

2,4-dioxopyrimidin-5-yl)-methyl]-L-tyrosine) has been identified in γ-irradiated mixtures of

Thy with Tyr or with a peptide containing Tyr, and its structure has been elucidated [169-

174]. The structure of this DNA-protein cross-link is illustrated in Fig. 3. Subsequently, Thy-

Tyr cross-links have been detected in mammalian chromatin upon exposure to γ-irradiation or by treatment with H2O2/iron or copper ions [175,176]. Its formation has been proposed to result from the addition of the allyl radical of Thy (see above) to C3 of Tyr, followed by oxidation or from the combination of the allyl radical of Thy with the phenoxyl radical of

Tyr. The latter is formed by ●OH addition to the Tyr ring followed by water elimination

[177]. The cross-linking was not inhibited by O2, most likely because the allyl radical of Thy

adds to Tyr in close proximity without first reacting with O2. The Thy-Tyr cross-link has also been observed in mammalian cells upon exposure to ionizing radiation, H2O2 or Fe(II)-ions

[178,179], and in renal chromatin of rats upon treatment with a renal carcinogen [180].

Additional DNA-protein cross-links have been identified in γ-irradiated mammalian chromatin in vitro between Thy and Gly, Ala, Val, Leu, Ile, Thr and Lys, and between Cyt

17 and Tyr [175,181,182]. A lysine-guanine cross-linking involving a guanine radical cation has been observed in aerated aqueous solution of a thymine-guanine-thymine oligodeoxynucleotide in the presence of a trilysine peptide due to riboflavin-mediated photosensitization; however, the corresponding Nϵ-(guanine-8-yl)-lysine cross-link has not been identified in mammalian chromatin [183].

2.4.4. Clustered lesions

Clustered lesions in DNA are also known as locally multiply damaged sites and are produced almost exclusively by ionizing radiations [184-189]. These lesions can be formed on the same strand or on opposite strands within one or two helical turns of DNA and can persist in living cells due to resistance to DNA repair by DNA glycosylases or by endonucleases. This field has extensively been reviewed [190].

3. Cellular repair of oxidatively induced DNA lesions

Living organisms evolved to possess DNA repair mechanisms to repair DNA damage and thus to protect the genetic stability for survival (reviewed in [11,191,192]). Failure to repair

DNA damage may lead to detrimental biological consequences for organisms. There are numerous DNA repair mechanisms. Oxidatively induced DNA lesions are generally repaired by base excision repair (BER) and, to a lesser extent, by nucleotide excision repair (NER) both of which include multiple steps and enzymes (reviewed in [11]). DNA lesions paired with a cognate DNA base are repaired by mismatch repair (MMR) [193-196]. There is repair in the nucleotide pool where modified 2'-deoxynucleoside triphosphates are dephosporylated by MutT in E.coli and its homolog MTH1 in human and other mammalian cells, and are thus prevented from being incorporated into DNA by DNA polymerases during DNA replication

[197-202]. MTH1 has been found to be overexpressed in many cancers [203]. Moreover, cancer cells have been shown to require MTH1 for efficient survival, suggesting that this

18 protein may be targeted as an anticancer therapeutic approach [204]. DNA single-strand breaks are repaired by mechanisms similar to those in BER, whereas homologous recombination or non-homologous end-joining mechanisms act on double-strand breaks

(reviewed in [11,205,206]).

3.1. Base excision repair

Base excision repair is highly conserved during evolution from bacteria to humans. It starts

with the removal (excision) of a DNA base lesion from DNA by a DNA glycosylase that

hydrolyzes the N-glycosidic bond between the sugar moiety and the modified base, leaving

behind an abasic site, also called an apyrimidinic/apurinic (AP) site. DNA glycosylases are

either monofunctional removing the DNA lesion only or possess an associated AP-lyase

activity. The 3'-phosphodiester bond of the AP site is hydrolyzed by a β- or β-δ-elimination mechanism that generates a strand break with a 3' α,β-unsaturated aldehyde (β-elimination) or a 5'-phosphate group (β-δ-elimination) [11]. Subsequently, AP-endonucleases, DNA polymerases and DNA ligases process AP sites to restore the DNA structure. The lyase activity is generally associated with DNA glycosylases specific for oxidatively induced DNA base lesions. BER consists of short-patch and long-patch pathways. The former is initiated by a bifunctional DNA glycosylase, whereas a monofunctional DNA glycosylase can start either pathway [207]. In general, the short-patch pathway is initiated by DNA glycosylases, whereas

AP sites resulting from spontaneous hydrolysis of DNA bases are repaired by the long-patch pathway [208,209]. DNA glycosylases are highly conserved from bacteria to humans and are divided into two families, the Nth superfamily and the Fpg/Nei family, on the basis of structure and sequence homology [208,210-212]. The members of the Nth superfamily are widely found in bacteria, archaea and eukaryotes, whereas those of the Fpg/Nei family are sparsely distributed across the phyla. All members of these families use a common

19 mechanism for catalysis that includes several steps [213,214]. DNA glycosylases in the

Fpg/Nei family are characterized by a helix-two turn-helix (H2TH) motif and a zinc or zincless finger motif for DNA binding. They also have a conserved N-terminus with a Pro residue, which is essential for catalysis. The Fpg/Nei family members are named after bacterial members formamidopyrimidine glycosylase (Fpg, also called MutM) and endonuclease VIII (Nei), and also include NEIL1, NEIL2 and NEIL3; the Nth superfamily contains E. coli endonuclease III (Nth), E. coli MutY, yeast Ntg1 and Ntg2, MUTYH, OGG1 and AlkA [215].

DNA glycosylases have distinct substrate specificities, although there exists a redundancy with respect to overlapping substrates. The determination of substrate specificities of DNA glycosylases has been performed using various substrates, methods and techniques. In general, most studies used oligodeoxynucleotides with a single DNA lesion incorporated at a specific position. The use of such substrates and applied analytical methods permitted to study the excision of a single modified DNA base only at a time. A different concept that uses damaged DNA with multiple lesions and the technique of gas chromatography-mass spectrometry (GC-MS) has been proposed for the determination of substrate specificities and excision kinetics of DNA glycosylases [49]. This technique permits the simultaneous

identification and quantification of multiple modified bases from all four DNA bases in a

given DNA sample. Therefore, it enables the determination of substrate specificities and

excision kinetics of DNA glycosylases by identifying which lesions are or are not excised

from DNA by a given DNA glycosylase. Subsequently, substrate specificities and excision

kinetics of numerous DNA glycosylases has extensively been studied (reviewed in

[22,216,217]).

3.1.1. Substrate specificities of prokaryotic DNA glycosylases

The concept described above has been used for the first time to investigate the substrate

specificity of E. coli Fpg [41]. This enzyme had originally been shown to recognize and remove purine-derived lesions with an opened imidazole ring such as 2,6-diamino-4- hydroxy-N7-methyl-5-formamidopyrimidine (Me-FapyGua) (derived from 7-methylguanine) and FapyAde [218-220]. Subsequent work reported the excision by E. coli Fpg of 8-OH-Gua

[221], and pyrimidine-derived lesions 5-OH-Cyt and 5-OH-Ura from oligodeoxynucleotides

[222]. The use of GC-MS demonstrated the excision of 8-OH-Gua, FapyGua and FapyAde by this enzyme, but no excision of pyrimidine-derived lesions from damaged DNA containing

multiple lesions [41] (for the structures of these compounds see Figure 1). A subsequent

study reported the excision of these three lesions by similar Michaelis-Menten kinetics [223].

These results clearly showed that FapyGua and FapyAde may also be the main substrates of

E. coli Fpg in cells, in contrast to the claim of 8-OH-Gua being the main physiological

substrate of this enzyme without providing any data on the former two compounds [224,225].

Five mutant forms of E. coli Fpg have been generated and used to investigate the effect of

single point mutations in the fpg gene targeting highly conserved amino acids on the

specificity of this enzyme [226]. The results showed that single mutations targeting amino

acids Lys-57, Lys-155 and Pro-2 dramatically affected the specificity up to a complete loss of

activity. A protein homologous to E. coli Fpg has been isolated from the bacterium

Deinococcus radiodurans that exhibits resistance to the effects of extreme doses of ionizing

radiation and other DNA-damaging agents [227,228]. This enzyme designated Deinococcus

radiodurans Fpg efficiently excised 8-OH-Gua, FapyGua and FapyAde similar to E. coli Fpg,

but with significantly different excision kinetics [229].

E. coli Nth of the Nth superfamily exhibits a broad substrate specificity for cytosine- and

thymine-derived lesions [222,230-234]. The use of GC-MS extended the substrate specificity

E. coli Nth for pyrimidine-derived lesions and also included purine-derived FapyAde

[235,236]. Most of the pyrimidine-derived lesions listed in Figure 1 have ben found to be the

substrates of this enzyme. Another DNA glycosylase endonuclease VIII (Nei) of E. coli exhibits strong homology to E. coli Fpg and other bacterial Fpg proteins, but no significant sequence similarity to E. coli Nth [237-240]. Both enzymes have overlapping substrate specificity [222,234]. The use of the GC-MS and damaged DNA with multiple lesions extended the substrate specificity of E. coli Nei and showed that this enzyme also excises

FapyAde as E. coli Nth does [241]. E. coli uracil DNA glycosylase (UNG), which is specific for removal of uracil from DNA [242], has also been found to act on cytosine-derived products 5-OH-Ura and isodialuric acid (5,6-dihydroxyuracil) [243,244]. E. coli MutY of the

Nth superfamily removes adenine paired with 8-OH-Gua [245].

3.1.2. Substrate specificities of eukaryotic DNA glycosylases

Structural and functional homologues of E. coli Nth have been found in yeast and humans

[246,247]. Schizosaccharomyces pombe Nth (S. pombe Nth) efficiently excised 5-OH-Cyt, 5-

OH-Ura, Thy glycol, 5-OH-6-HThy and 5,6-diOH-Cyt from DNA, exhibiting a narrower substrate specificity than E. coli Nth [246,248]. Human NTH1 acted on the same DNA lesions as S. pombe Nth, albeit with significant differences in excision kinetics [249].

Efficient excision of purine-derived FapyAde by NTH1 has also been observed [250].

FapyAde accumulated in nth1–/– mice, providing the evidence that FapyAde is the

physiological substrate of NTH1 [250,251]. (Fig. 4) illustrates the levels of FapyAde and 8-

OH-Gua in livers of nth1–/–, ogg1–/– and ogg1–/–/nth1–/– mice, demonstrating the accumulation

of FapyAde in nth1–/– mice when compared to wt mice, but not in ogg1–/– mice, and that of 8-

OH-Gua in ogg1–/–/mice, but not in nth1–/– mice. As expected, both compounds accumulated

in double knockout animals. The use of liver mitochondrial and nuclear extracts of wild type

(wt)-mice and nth1‒/‒, ogg1–/– and ogg1–/–/nth1–/– mice, and oligodeoxynucleotides containing

FapyAde confirmed this finding and also that FapyAde is not a substrate of OGG1 [250], as

had been previously shown in in vitro experiments [252]. The repair of 5,6-dihydrouracil has

also been found to be reduced, but not nullified in nth1–/– mice [253].

Schizosaccharomyces cerevisia possesses two DNA glycosylases/AP lyases named Ntg1

and Ntg2, which relate to each other and to their functional homologue E. coli Nth [254-256].

These DNA glycosylases have been shown to excise a number of pyrimidine-derived lesions, and purine-derived lesions FapyAde and FapyGua [257,258]. Thus, the cross-activities of

Ntg1 and Ntg2 are clearly different from E. coli Nth and Nei in that they efficiently remove

FapyGua in addition to FapyAde. Human uracil DNA glycosylase, which is mainly specific for removal of uracil from DNA [259], has been found to also excise cytosine-derived

products isodialuric acid, 5-OH-Ura and alloxan from DNA containing multiple lesions

[260,261]. Similarly, human SMUG1 exhibited specificity for the same lesions [261].

Functional homologues of E. coli Fpg, named OGG1, have been discovered in eukaryotes

[262-266]. All these DNA glycosylases exhibited an identical substrate specificity with the

excision of 8-OH-Gua and FapyGua from damaged DNA with multiple lesions, although excision kinetics somewhat varied among the enzymes [252,264,267-269]. The failure of

FapyAde excision by OGG1 from eukaryotes indicates significant differences between these enzymes and E. coli Fpg. Two different types of human OGG1 (hOGG1) have been discovered and designated α-hOGG1 and β-hOGG1 [263]. α-hOGG1 is targeted to the nucleus, whereas β-hOGG1 is located in the mitochondrion [270,271]. Two forms of α-

hOGG1 due to a polymorphism at codon 326, α-hOGG1-Ser326 and α-hOGG1-Cys326, have

been found in human cells [272,273]. α-hOGG1-Ser326 and α-hOGG1-Cys326 efficiently excised FapyGua and 8-OH-Gua from damaged DNA with multiple lesions, with the former exhibiting ≈ 2-fold greater activity than the latter [252]. Both forms had a greater preference for FapyGua than 8-OH-Gua. Two mutated forms of α-hOGG1-Ser326, α-hOGG1-Gln46 and

α-hOGG1-His154, have been found in tumor cells [268,274,275]. Both forms exhibited

efficient activity on FapyGua and 8-OH-Gua; however, their activity was lower than that of

α-hOGG1-Ser326 [268]. MUTYH, which is a homologue of E. coli MutY, also plays a role in the repair of 8-OH-Gua by removing adenine paired with it [193,276,277].

E. coli Nei-like DNA glycosylases have been discovered in eukaryotes, and named

NEIL1, NEIL2 and NEIL3 [278-285]. NEIL1 is cell cycle regulated and may thus be associated with the replication fork [286-289]. It is located to both the nucleus and mitochondrion, lending credit to its importance in maintaining the genetic stability [250].

NEIL1 mainly removes FapyAde and FapyGua from damaged DNA with multiple lesions,

and also Thy glycol and 5-OH-5-MeHyd albeit to a lesser extent [278,290]. In agreement

with in vitro studies, FapyAde and FapyGua have been shown to be the main physiological

substrates of NEIL1 in vivo [251,291]. NEIL1 exhibits no detectable activity toward 8-OH-

Gua in vitro or in vivo [122,278,290-293]. Recently, an additional specificity of NEIL1 has

been discovered. The accumulation of R-cdA and S-cdA in liver DNA of neil1‒/‒ mice has

been observed [122]. Since R-cdA and S-cdA are repaired by NER and not by BER [108,294-

297,297,298], this finding suggested that NEIL1 may be involved in NER in addition to its function as a DNA glycosylase in BER. To this end, there is evidence that NEIL1 may

interact with proteins of the NER complex. For example, similar to NEIL1, the Cockayne

syndrome complementation group B protein (CSB) plays a role in the repair of S-cdA [120],

and stimulates the action of NEIL1 on FapyAde and FapyGua [291]. This is in contrast to its lack of interaction with OGG1 [299]. Moreover, CSB and NEIL1 coimmunoprecipitate and

colocalize in HeLa cells, indicating that these proteins cooperate in the repair of

formamidopyrimidines [291]. NEIL1 may also interact with other NER proteins and

accelerate the repair of R-cdA and S-cdA, since it cannot itself initiate the BER at these

lesions.

The other Nei-like DNA glycosylase NEIL2 has been shown to preferentially excise pyrimidine-derived lesions from oligodeoxynucleotides with bubble structures [280,300]. It also exhibited specificity for removal of Sp and Gh from oligodeoxynucleotides [301]. Thus far, however, no excision by NEIL2 of any base lesions from DNA with multiple lesions has been observed. FapyAde and FapyGua have also been found to be the main substrates of

NEIL3; however, the efficient removal by this enzyme of pyrimidine-derived lesions has been observed from DNA with multiple lesions as well [293]. 8-OH-Gua has been shown to be not a substrate of NEIL3, either; nevertheless, its oxidation products Sp and Gh were efficiently removed by NEIL3 and also by NEIL1 from synthetic oligodeoxynucleotides

[301,302].

3.1.3. Repair of sugar lesions

Sugar lesions within DNA or bound to DNA as end groups (Figure 2), the oxidatively induced formation of which was discussed above, constitute AP sites with a modified 2'- deoxyribose moiety in DNA. On the other hand, AP sites with the intact 2'-deoxyribose moiety are formed in DNA of living cells by spontaneous hydrolysis of the glycosidic bond that occurs several thousand times per day per cell and also by the action of DNA glycosylases on sites with damaged DNA bases (reviewed in [11,303]). BER is the primary pathway that repairs AP sites in mammalian cells. Although multifunctional DNA glycosylases can cleave AP sites, apurinic/apyrimidinic endonuclease 1 (APE1) processes most AP sites via hydrolysis of the phosphate bond 5' to the AP site creating a single strand break with a 3'-OH group and a 5' terminal 2'-deoxyribose phosphate (dRP) residue

[304,305]. Subsequently, DNA polymerase β (Pol β) and other enzymes repair the remaining nick. In certain circumstances, AP sites are also repaired by BER via long-patch repair [306].

2-Deoxypentose-4-ulose, erythrose and 2-deoxyribonic acid (or its lactone form) within

DNA, and 2-deoxytetradialdose as an end group (Figure 2) are the substrates of the first step

25 of BER involving APE1, Pol β and Pol λ; however, they are not processed as efficiently as

AP sites with the intact 2'-deoxyribose moiety [307-319]. 2-Deoxyribonic acid is repaired

almost exclusively by long-patch BER [315,320-322]. The sugar lesions can cause

irreversible inhibition of BER enzymes and the formation of interstrand DNA cross-links

[317,319,323-333]. Furthermore, DNA-protein cross-links occur between 2-deoxyribonic acid and the proteins E. coli endonuclease III, Pol β and histones [310,311,315,321,334,335].

Thus, the action of APE1 in short-patch BER on 2-deoxyribonic acid is stalled by the formation DNA-protein cross-links between this lesion and Pol β [315]. On the other hand, long-patch BER prevents the formation of DNA-protein cross-links between 2-deoxyribonic acid and Pol β [320].

3.2. Nucleotide excision repair

Nucleotide excision repair removes bulky DNA-distorting lesions from DNA [209,336-

341]. Global genome repair and transcription-coupled repair constitute two distinct mechanisms of NER and are responsible for the repair of the entire genome and preferential repair of transcribing DNA strands, respectively. NER has also been reported to repair oxidatively induced lesions such as thymine glycol and 8-OH-Gua [342-344]. Almost three decades ago, it has been proposed that 8,5'-cyclopurine-2'-deoxynucleosides would not be repaired by BER because of the 8,5'-covalent bond between the base and sugar moieties, and thus they would likely be subject to repair by NER [108,294]. Indeed, NER, not BER, has been shown to repair R-cdA and S-cdA, with the former being repaired more efficiently than the latter [295,296]. On the other hand, inefficient repair of R-cdG and S-cdG has been reported by E. coli NER enzymes in vitro [345]. Recently, a number of DNA glycosylases including Fpg, NEIL1 and OGG1 have been tested on oligodeoxynucleotides containing cdA or cdG; however, no cleavage has been detected, confirming the lack of activity of BER on

8,5'-cyclopurine-2'-deoxynucleosides [297]. Furthermore, these DNA glycosylases at high

concentrations failed to form DNA-protein complexes with oligodeoxynucleotides containing

S-cdA or S-cdG. In contrast, HeLa cell extracts excised 24-32 base-pair fragments from long

double-stranded oligodeoxynucleotides containing S-cdG or S-cdA, with a greater efficiency

for the former than the latter [297]. The efficiency of repair depended on the complementary

base opposite the lesion. Just recently, the structural basis for the recognition of 8,5'-

cyclopurine-2'-deoxynucleosides by NER has been investigated in detail [298]. Using

extracts of HeLa cells and oligodeoxynucleotides containing R-cdA, S-cdA, R-cdG or S-cdG with identical sequence contexts, NER has been shown to excise the R-diastereomers of both cdA and cdG with a ≈ 2-fold greater efficiency than their S-diastereomers. However, the overall excision efficiencies between cdA and cdG were similar. The R-diastereomers of cdA and cdG caused greater distortion of the DNA backbone than their S-diastereomers, correlating with NER incision efficiencies. Recently, the apurinic/apyrimidinic endonucleases, E. coli Xth and human APE1 have been reported to remove S-cdA at the 3' terminus of duplex DNA, but not that located at 1 or more nucleotides away from this end

[346]. This mechanism has been suggested as a complementary pathway to NER to remove

S-cdA and possibly other 8,5'-cyclo-2'-deoxynucleosides as well. This is an intriguing mechanism; however, the formation of an 8,5'-cyclo-2'-deoxynucleoside as the end unit of a broken DNA strand must occur for this mechanism to be active. In support of this notion, evidence has recently been provided for the incorporation of R- and S-diastereomers of cdATP into DNA by replicative DNA polymerases, inhibiting further DNA synthesis and thus generating a DNA strand with a cdA at the 3'-terminus [347].

4. Genetic effects of oxidatively induced DNA lesions

Failure to repair DNA lesions before replication of DNA-damaged cells may lead to cell death, cytotoxicity and mutagenicity, and ultimately to disease processes including carcinogenesis. If a DNA lesion is not removed from DNA, it may be tolerated and bypassed by DNA polymerases, which may mispair it with a non-cognate intact DNA base, leading to a mutation following the next step of replication. On the other hand, a DNA lesion can block the action of DNA polymerases to perform DNA synthesis, thus becoming a lethal lesion leading to cell death. The interplay between the repair and the two types of replication may ultimately determine the future of a DNA-damaged cell and potentially that of the organism.

It should be mentioned that a DNA lesion can also pair with a cognate DNA base, which will make it neither lethal nor mutagenic [348].

4.1. Purine -derived lesions

Among the oxidatively induced DNA lesions, 8-OH-Gua has been the most investigated lesion in terms of its biological effects, perhaps at the expense of the other equally important lesions. In an early work, this lesion has been shown to induce numerous mutations in vitro that included a G → T transversion mutation indicating mispairing of 8-OH-Gua with Ade

[349]. In a subsequent work, the genetic effects of 8-OH-Gua have been determined after transfection of the single-stranded site-specifically modified viral genome into wild-type E. coli [350]. This work provided direct evidence that 8-OH-Gua is premutagenic in vivo and leads to the G → T transversion mutation as a major mutagenic event. The bypass efficiency of 8-OH-Gua generally amounted to 85-90% [351]. Frequency of mispairing depended on the polymerase [352,353]. 8-OH-Gua has also been shown to pair with cognate Cyt, albeit to a lesser extent, causing no mutations [350,352,354-356]. Fig. 5 illustrates the 8-OH-Gua•Ade mispair [356]. Oxidation products of 8-OH-Gua such as Sp and Gh have been found to

28 exhibit mutagenic effects as well as cytotoxic effects depending on the polymerase, sequence context, etc. (reviewed in [72]). The other equally important guanine-derived lesion FapyGua has been shown to also mispair with non-cognate Ade (Fig. 5), leading to G → T transversion mutations [357,358]. In simian kidney cells, FapyGua has even been more mutagenic than 8-

OH-Gua [359,360], although a weak mutagenicity for FapyGua in E. coli has been observed

[361]. A subsequent work unequivocally demonstrated significant in vivo mutagenicity of

FapyGua in an E. coli triple mutant lacking Fpg, Nei and MutY glycosylase activities by expressing glycosylase domains of mouse NEIL1 and NEIL3 in these cells [293]. G → T transversion mutations are the second most common somatic mutations found in human cancers, constituting 14.6% of all mutations in the tumor suppressor gene TP53 [362]. Of course, this does not mean that these mutations entirely result from 8-OH-Gua and FapyGua.

Other DNA lesions may lead to such mutations as well.

8-OH-Ade has been shown to pair with cognate Thy and to mispair with Gua and Ade depending on the polymerase [363-366]. Fig. 5 illustrates the 8-OH-Ade•Gua mispair [367].

The lack of mutagenicity of 8-OH-Ade in bacteria has been reported [368]. On the other, this lesion caused A → G transition and A → C transversion mutations in mammalian cells at a mutation frequency of ≈1% only [366,369]. FapyAde, which has the precursor C8-OH‒ adduct radical of Ade in common with 8-OH-Ade [10], has been found to direct Klenow exo‒

fragment to misincorporate Ade opposite itself, potentially leading to A → T transversions

[370]. The FapyAde•Ade mispair is illustrated in Fig. 5 [370]. Both 8-OH-Ade and FapyAde

were very weakly mutagenic in simian kidney cells when compared to 8-OH-Gua and

FapyGua [359]. In the case of another adenine-derived product, 2-OH-Ade, DNA polymerases inserted all DNA bases opposite this lesion with the possibility of leading to all mutations involving Ade [371,372]. In E. coli, the frequency and the spectrum of the mutations depended on the sequence contexts and the lagging/leadings template strands

[373]. In simian COS-7 cells, the primary mutation has been a ‒1 deletion followed by A →

G transitions and A → T transversions [374]. The observed mutation frequencies (0.6 % ‒ 0.1

%) have been comparable with those of 8-OH-Gua in NIH3T3 cells [375,376]. The results

suggested the formation of 2-OH-Ade•Cyt and 2-OH-Ade•Ade mispairs in living cells. In

addition, 2-OH-Ade formed in the nucleotide pool induced G•C → T•A transversions in E.

coli [377]. Moreover, 2-OH-dATP led to G•C → A•T transitions and G•C → T•A

transversions during in vitro replication using HeLa cell extracts, albeit to a lesser extent

[378].

4.2. Pyrimidine-derived lesions

Thymine glycol is one of the major and most investigated products of thymine. Its biological effects have also been investigated extensively. Thymine glycol correctly pairs with Ade and is thus poorly mutagenic [348,379-382]. In some sequence contexts, however, it is bypassed by DNA glycosylases and can pair with non-cognate Gua to a low extent, leading to T → C transitions [383-386]. In general, Thy glycol constitutes a strong block to

DNA polymerases and is thus a lethal lesion [348,379,382,383,386-391]. 5-Hydroxy-6- hydrothymine also strongly blocks an E. coli DNA polymerase and is a lethal lesion [392].

5,6-Dihydrothymine pairs with cognate Ade and is not a block to DNA polymerases; therefore, it is neither lethal nor mutagenic [393,394].

Deamination and dehydration reactions of cytosine glycol result in the formation of 5-OH-

Cyt, 5-OH-Ura and uracil glycol [91,235]. These lesions can exist in DNA simultaneously

[235]. Uracil glycol, 5-OH-Cyt and 5-OH-Ura pair with non-cognate Ade leading to C → T transitions [348,380,386,395-397]. Fig. 6 illustrates of the pathway of the formation of C →

T transitions due to mispairing of 5-OH-Ura and 5-OH-Cyt with Ade. The incorporation of the intact base opposite these lesions depended on the sequence context. Thus, C → G

30 transversions have also been observed as a result of the mispairing of 5-OH-Cyt with Cyt

[398]. The minor anionic imino tautomer of 5-OH-Cyt has been shown to mispair with Ade

(Fig. 5) and to be the likely source of C → T transitions caused by this lesion [399]. In E. coli, 5-OH-Ura and uracil glycol led to C → T transitions with high frequency (83% and

80%, respectively), whereas 5-OH-Cyt elicited the same mutations with a lower frequency

(0.05%) and C → G transversions even to a much lesser extent [397]. These results suggested

that 5-OH-Ura and uracil glycol may be the main cause of C → T transitions in cells. These

transition mutations have been found to be the most frequently occurring mutations in human tumors and in TP53 [362,400], and from oxidatively induced DNA damage [396,401].

However, this does not mean that 5-OH-Ura and uracil glycol would be the only source of C

→ T mutations. For example, errors by replicative DNA polymerases [402,403], and deamination of 5-methylcytosine can also cause these mutations at high frequency [15,404].

4.3. Sugar lesions

AP sites with an intact 2'-deoxyribose are mutagenic [405,406]. Similarly, the sugar lesions 2-deoxypentose-4-ulose, erythrose and 2-deoxyribonic acid exhibit mutagenic effects

[322,407-412]. They also form interstrand DNA cross-links and DNA-protein cross-links, and thus are converted to other types of DNA damage such as DNA strand breaks that may additionally be deleterious to living cells [310,311,315,319,321,329-334,413]. The ability of

2-deoxypentose-4-ulose and 2-deoxytetradialdose to inactivate Pol β and Pol λ indicates that these lesions may be a significant source of the cytotoxicity caused by DNA-damaging agents

[317,326-328]. Furthermore, there is evidence that strand scission at AP sites and at the sugar lesions 2-deoxypentose-4-ulose and 2-deoxyribonic acid are accelerated in nucleosome core particles [319,414-418]. In early studies, the treatment of DNA with neocarzinostatin chromophore (NCS) resulted in significant amounts of G•C → A•T transitions, pointing to 2-

31 deoxyribonic acid as the highly likely source of these mutations [419,420]. Indeed, 2- deoxyribonic acid has been identified at the site of NCS-induced Cyt release in a certain sequence [421,422], and confirmed to be a source of transition mutations [407,409].

Incorporation of Ade by Klenow opposite this lesion is on a par with the observed mutations

[423]. In addition, 2-deoxyribonic acid constitutes a stronger block to the translesional synthesis by Klenow than the AP site with an intact 2'-deoxyribose [423,424]. Cross-linking

occurring between 2-deoxyribonic acid and histones may also contribute to genotoxic effects

of this lesion in DNA [315,335]. Similar to 2-deoxyribonic acid, Ade is preferentially

incorporated opposite erythrose within DNA, which exhibits an effect on Klenow exo‒

similar to that of the AP site with an intact 2'-deoxyribose moiety [313]. Thus, erythrose may be a mutagenic lesion as well. Gua was also incorporated, albeit to a much lesser extent, but not Cyt or Thy to any significant extent. On the other hand, Pol V in E. coli preferentially incorporated Gua opposite 2-deoxyribonic acid, even more than opposite the AP site, and discriminated between Gua and Ade [425]. Moreover, 2-deoxyribonic acid induced significant Gua incorporation in E. coli under SOS conditions [411]. In the case of 2-

deoxypentose-4-ulose, Klenow exo‒ efficiently incorporated Ade opposite, followed by Gua

with ≈ 10-fold greater preference for the former than the latter [426]. Klenow exo+ exhibited

slightly less preference for Ade incorporation than Klenow exo‒. Although the sugar lesions

and AP sites with an intact 2'-deoxyribose moiety have substantial structural differences, they

exhibit similar effects on the activities of Klenow exo+/exo‒ fragments and bypass polymerases. Taken together, sugar lesions in DNA (Fig. 2) apparently exhibit significant mutagenic and genotoxic effects that rival those exhibited by modified DNA bases.

4.4. Tandem lesions

4.4.1. 8,5'-Cyclopurine-2'-deoxynucleosides

Genetic effects of 8,5'-cyclopurine-2'-deoxynucleosides have drawn much attention in the past decade and have been investigated quite extensively. S-cdA blocks transcription and several DNA polymerases, reduces transcription and causes transcriptional mutagenesis

[295,296,427-429]. Multiple nucleotide deletions occur due to incorporation of adenosine opposite to the next 5'- to S-cdA by RNA polymerase II [429]. Recently, S-cdG has been found to be a strong block to replication and a potent DNA polymerase V-dependent mutagenic lesion in E. coli, leading mainly to G → A transitions, indicating it mispairs with

Thy, and to G → T transversions to a lesser extent [345]. Fig. 5 illustrates the S-cdG•Thy mispair [297,430]. The pathway leading to G → A transitions by this mispair is shown in Fig.

6. Similarly, S-cdG and S-cdA strongly block DNA replication, and also cause G → A transitions and A → T transversions in five different strains of E. coli to an extent of 20% and

11%, respectively [431]. On the other hand, both human polymerase η and S. cerevisiae polymerase η bypassed S-cdG and S-cdA accurately and efficiently, indicating that mutagenic bypass of these lesions may relate to other DNA polymerases [432]. Indeed, S-cdG and S- cdA have been shown to be strong blocks to replication by DNA polymerase IV, exo-free

Klenov fragment and Dpo4 [433]. Furthermore, the same study showed the occurrence of both A → T transversions and A → G transitions caused by S-cdA in equal frequency in wild-type E. coli; however, the results with DNA polymerase IV-deficient strain suggested that DNA polymerase IV had played a role in A → G transitions caused by S-cdA. DNA polymerase IV incorporated Cyt and Thy opposite S-cdA with almost equal efficiency, whereas the cognate Cyt was efficiently inserted opposite S-cdG by this polymerase. On the other hand, the incorporation of Thy by the exo-free Klenov fragment was more than twice

33 that of Cyt by DNA polymerase IV [433]. The S-cdA•Thy pair (Fig. 5) has been found to be

very stable when compared to the Ade•Thy pair [434].

4.4.2. Tandem lesions

Two tandem lesions identified in vivo, i.e., Gua[8,5]Cyt and Gua [8,5-Me]Thy (Fig. 3),

have been shown to be cytotoxic and mutagenic [151,159,160]. In E. coli, Gua[8,5]Cyt blocked DNA replication and exhibited significant mutagenicity in vivo due to misincorporation opposite the 5'-guanine moiety that included G → T and, to a lesser extent,

G → C transversion mutations. DNA polymerase V was responsible for the mutagenicity of this lesion. S. cerevisiae DNA polymerase η inserted the cognate dGMP opposite 3'-cytosine moiety, indicating that the 3'-cytosine moiety of this lesion is not mutagenic. However, this polymerase inserted non-cognate dAMP and dGMP opposite the 5'-guanine moiety, which may result in G → T and G → C transversion mutations. The Klenow fragment of E. coli

DNA polymerase I stopped synthesis after incorporating the cognate dAMP opposite 3'- thymine moiety of Gua[8,5-Me]Thy; however, S. cerevisiae DNA polymerase η performed synthesis past this lesion and incorporated non-cognate dAMP and dGTP opposite 5'-guanine.

Thus, Gua[8,5-Me]Thy may give rise to G → T and G → C transversion mutations.

Gua[8,5]Cyt and Gua[8,5-Me]Thy may distort the DNA structure, leading to lack of recognition by DNA polymerases of the hydrogen bonding property of the 5'-guanine moiety that assumes a syn configuration in contrast to the anti configuration of the 3'-cytosine moiety

[151]. Thus, purine nucleotides may be inserted opposite 5'-guanine more efficiently than pyrimidine nucleotides. Mutagenic effects of the Thy-Tyr cross-link have not been investigated.

5. Oxidatively induced DNA damage and cancer

DNA repair is essential to life. Thus, unrepaired DNA lesions may lead to detrimental consequences in living organisms. DNA lesions that escape repair may accumulate in the genome, progressively leading to increase in mutation rate, i.e., mutator phenotype, and thus in genetic instability, a hallmark of cancer [3,12,14,15,191,435,436]. Defective DNA repair is associated with carcinogenesis [14,15,19,437-455]. Persistent oxidative stress exists in cancer

[456]. In agreement with this fact, precancerous and cancerous tissues or cancer cell lines have been shown to contain oxidatively induced DNA lesions at elevated levels when compared to surrounding cancer-free tissues or to normal cell lines [112,113,116,117,457-

470]. Most of these lesions are mutagenic (see above) and may thus play a significant role in carcinogenesis and other disease processes. DNA repair genes are also prone to mutations.

Germline mutations cause genetic instability and predisposition to cancer [191,449].

Polymorphisms of DNA repair genes including the BER genes also increase the cancer risk

and may determine the outcome of disease for patients [191,444,449,454,471-477]. Therapy

resistance occurs in tumors and may be due to defects in DNA repair, adversely affecting the

outcome of patient survival [16-18,449,454,478-480]. Several studies reported lower levels of

ethano-DNA adducts and oxidatively induced DNA lesions in cancerous tissues than in

surrounding non-cancerous tissues, providing the evidence that DNA repair may be

upregulated in cancerous tissues [123,481,482]. Upregulated DNA repair in cancerous tissues

may cause resistance to therapeutic agents. Despite their adverse effects, DNA repair

alterations may have the potential to help develop the concept of personalized cancer therapy

and may also serve as promising predictive cancer biomarkers [480].

5.1. Role of DNA glycosylases of BER in carcinogenesis

As mentioned above, most of oxidatively induced DNA base lesions, except for 8,5'-cyclo-

2'-deoxynucleosides which are subject to NER, are repaired by BER with the action of

various DNA glycosylases in the first step of this mechanism. BER removes numerous

endogenously and exogenously induced lesions from cellular DNA per day and thus protects

the genetic stability and plays an important role in disease prevention including cancer

prevention. Thus, defects in BER are associated with neurological disorders and cancer

[214,476,477,483]. Polymorphic variants of DNA glycosylases have been found in human

populations in connection to various cancer incidences. Moreover, mouse models with

knockouts of DNA glycosylase genes have been developed to study resulting phenotypes.

5.1.1. OGG1

OGG1 has been one of the most investigated DNA glycosylases. Among many oxidatively induced DNA lesions (Fig. 1), 8-OH-Gua and FapyGua are the main physiological substrates of human OGG1 (hOGG1) and its eukaryotic homologues. This fact has been proven in

experiments in vitro and in vivo (discussed above). When compared to wt-mice, simultaneous accumulation of both lesions has been demonstrated in livers of ogg1‒/‒ mice; however, no

accumulation of FapyAde has been observed [122,250]. These observations are on a par with the in vitro findings of the substrate specificity of OGG1 (see above). These results have also been confirmed using liver mitochondrial and nuclear extracts of wt-mice and ogg1‒/‒ mice,

and oligodeoxynucleotides containing FapyAde, FapyGua or 8-OH-Gua at a defined position

[250]. Human ogg1 gene contains a variety of single-nucleotide polymorphisms

(http://www.ncbi.nlm.nih.gov/sites/entrez?db=snp) [273,275,452,484-486]. The most

common polymorphic variant of hOGG1 is OGG1-Cys326 with a high frequency in human population [452]. Many studies provided the evidence for the association of this variant with

36 the risk of a number of cancers [272,274,275,447,452,484,487-511]. On the other hand, no risk of some cancers with OGG1-Cys326 has been demonstrated [512-514]. Other polymorphic variants of hOGG1 such as OGG1-His154, OGG1-Gln46, OGG1-Gln209,

OGG1-Thr321, OGG1-His154, OGG1-131Gln have also been found in some cancer cell lines and tissues [268,272,274,275,515,516]. Differences between the activities of the variants of

OGG1 and the wt-OGG1 have been observed. OGG1-Cys326, OGG1-His154, OGG1-Gln46 and OGG1-Asn322 exhibited significantly lower activities than that of wt-OGG1

[252,268,517]. In contrast, excision kinetics of one variant, i.e., OGG1-Val288 has been

found to be similar to that of wt-OGG1 [517]. Low OGG1 activity has been shown to

constitute a risk factor in some types of cancers [447,518-521]. The expression of OGG1 in

eighteen human cancer and three normal cell lines has been studied [468]. Sixteen of the

cancer cell lines exhibited overexpression of OGG1; however, two of them had even lower

expression of OGG1 than normal cell lines. Increased levels of 8-OH-Gua have been

observed in these two cancer cell lines. Their mitochondria also exhibited compromised

repair of 8-OH-Gua, suggesting OGG1 plays a role in the maintenance of the mitochondrial

genome. Overall, the results implicated 8-OH-Gua repair defects in certain lung cancers.

Despite all these findings discussed above, ogg1‒/‒ mice have been found to be viable with no tumor formation, although they exhibited organ-specific accumulation of 8-OH-Gua and

elevated G → T transversion mutations [522-524]. Treatment of such mice with KBrO3

caused a dramatic increase in the level of 8-OH-Gua in both livers and kidneys; however, no

tumor formation has been observed in these organs [525-527]. Furthermore, exposure of both

wt-mice and ogg1–/– mice to low doses of ionizing radiation, ogg1–/– mice exhibited a

significant increase in G → T transversions in their brains; however, no tumor development

has been observed [528]. On the other hand, an increase in spontaneous lung adenomas and

carcinomas along with a significant accumulation of 8-OH-Gua has been observed in a

37 different strain of ogg1‒/‒ mice [529]. However, 8-OH-Gua has been the only measured lesion

among many other DNA lesions (Fig. 1) to unequivocally conclude that this effect had been due to its accumulation only. Additional knockout of the mth1 gene in these mice further increased the level of 8-OH-Gua with no significant increase in carcinogenesis. Exposure to chronic UVB radiation increased the level of 8-OH-Gua in ogg1–/– mice and to make them susceptible to skin carcinogenesis [530]. Again, no other lesion among many DNA lesions has been measured to draw useful conclusions about the role of 8-OH-Gua in the observed carcinogenesis. Due to all these facts discussed above, there seems to be no clear association of ogg1 deficiencies and the accumulation of 8-OH-Gua with human carcinogenesis. On the other hand, mice with deficiencies in both ogg1 and mutyh exhibited the incidence of tumors predominantly in lung and ovarian tumors, and lymphomas with accompanying G → T transversions in the majority of lung tumors at codon 12 of the K-ras oncogene [531]. Mutyh–

/– mice developed no tumors similar to ogg1–/– mice. In another study using the same strain of

mutyh–/– mice, no age-associated accumulation of 8-OH-Gua has been observed in several organs except for liver; however, mutyh–/–/ogg1–/– mice accumulated 8-OH-Gua and exhibited

the incidence of lung and small intestine cancers [532]. These findings may indicate the

requirement for carcinogenesis of synergistic occurrence of mutations in several DNA repair

genes.

5.1.2. NEIL proteins

Extensive studies in the past decade demonstrated a critical role for NEIL1 in the

maintenance of the genetic stability and disease prevention. Polymorphic variants of NEIL1

have been discovered in humans. Three NEIL1 variants, NEIL-Arg242, NEIL1-Arg245 and

NEIL1-Gly334 have been found in human gastric cancers; however, they exhibited activity similar to that of wt-NEIL1 [533]. A NEIL1 deletion variant with no activity has also been observed in gastric cancers [533]. Three promoter polymorphisms in neil1 have been

38 identified in gastric cancers, although the consequences of these mutations have not been

determined [534]. Accumulation of mutations in the hprt locus and their increase by

oxidative stress have been observed in neil1-knockdown human bronchial cells and Chinese

hamster ovary cells [535]. Knocking down neil1 rendered embryonic stem cells more sensitive to killing effects of ionizing radiation [536]. Exposure of human carcinoma cells to

oxidative stress increased expression of NEIL1 [537]. NEIL1-Asp83 and NEIL1-Lys181

variants have been found in patients with cholagiocarcinoma and primary sclerosing

cholangitis, respectively [538]. Rare NEIL1-Ser203 and NEIL1-Gln339 variants have been

observed in patients with colorectal adenomas; however, the latter variant also existed in a

control individual [516]. Four polymorphisms have been reported for the human neil1 [292].

Corresponding variants NEIL1-Cys82, NEIL1-Asp83, NEIL1-Asn252 and NEIL1-Arg136 have been isolated and characterized [292]. An AP site- or a Thy glycol-containing oligodeoxynucleotide, and DNA substrates with multiple lesions have been used to test the

AP-lyase and glycosylase activities of these variants in comparison to wt-NEIL1. NEIL1-

Cys82 and NEIL1-Asn252 exhibited a β,δ-elimination activity on the AP-site as wt-NEIL1.

However, NEIL1-Asp83 had a β-elimination activity only and NEIL1-Arg136 exhibited no activity at all. An efficient excision of FapyAde and FapyGua from DNA with multiple lesions and that of Thy glycol from an oligodeoxynucleotide by wt-NEIL1, NEIL1-Cys82 and NEIL1-Asn252 has been observed. In contrast, no DNA glycosylase activity whatsoever has been detected for NEIL1-Asp83 and NEIL1-Arg136. The measurement of the specificity constants revealed a preference of NEIL1 and its active variants for FapyAde over FapyGua, perhaps because FapyGua is also a substrate of OGG1, which does not act on FapyAde, as was discussed above. Overall, this work suggested that individuals with neil1 mutations may be at risk for disease development.

A neil1–/– mouse model has been developed to investigate the consequences of NEIL1

deficiency in vivo [251,539-541]. Male neil1–/– mice developed obesity by 24 months of age,

accompanied by severe fatty liver, increased circulating lipids and hyperinsulinemia, which

are collectively called metabolic syndrome. Female neil1–/– mice also developed obesity, but

to a lesser extent than male neil1–/– mice [539]. In this context, there is evidence that

metabolic syndrome may be associated with certain types of cancer [542-545]. When exposed

to oxidative stress, male neil1–/– mice gained more weight than wt-mice [539,541]. Mutations

increased in mitochondrial (mt) DNA and the mtDNA content decreased in livers of these

mice. All these results strongly suggested that NEIL1 may play an important role in disease

prevention and that NEIL1 deficiency may lower the threshold for tolerance of oxidatively

induced DNA damage.

In another study, nth1–/– and neil1–/–/nth1–/– mice in addition to neil1–/– mice have been developed to understand the roles of NEIL1 and NTH1 in carcinogenesis [251]. Very few tumors have been observed in the first year of these animals’ lives. During the second year, however, neil1–/– and nth1–/– male and female mice developed pulmonary tumors and

hepatocellular tumors to a low extent up to ≈ 4 % and ≈ 15%, respectively. However, nth1–/–

/neil1–/ – mice exhibited a dramatic increase in both cancer incidences at a much greater rate

(up to ≈ 75 %) than neil1–/– or nth1–/– animals [251]. These results are illustrated in Fig. 7.

Pulmonary tumors contained activating GGT → GAT transitions in codon 12 of their K-ras

oncogene. This is in contrast to the activating GGT → GTT transversions in codon 12 of the

K-ras oncogene of the pathologically similar pulmonary tumors in ogg1–/–/mutyh1–/– mice

[531]. Oxidatively induced DNA lesions have been measured in DNA of livers, kidneys and brains of wild type and knockout animals. The results revealed significant accumulation of

FapyAde in all three organs of neil1–/– and neil1–/–/nth1–/– mice, and in kidneys of nth1–/–

mice, and that of FapyGua in livers and kidneys of neil1–/– and neil1–/–/nth1–/– mice, but not in

40 brains. In contrast, no accumulation of 8-OH-Gua has been observed in these organs of any

knockout animals. Enhanced levels of FapyAde and FapyGua in knockout animals is on a par

with the substrate specificity of NEIL1 observed in vitro using DNA containing multiple

lesions (see above), and provide additional evidence for FapyAde and FapyGua, but not for

8-OH-Gua, to be the in vivo substrates of NEIL1. Significant accumulation of FapyAde and

FapyGua in cancer-prone knockout mice strongly suggests a role for these compounds in

carcinogenesis, and for the involvement of NEIL1 and NTH1 in cancer prevention. The

absence of 8-OH-Gua accumulation and GGT → GTT transversions of codon 12 in the K-ras

oncogene, which is typical of tumors in ogg1–/– and muty–/– mice, unequivocally excludes the

involvement of 8-OH-Gua in the tumor incidences observed in neil1–/– and nth1–/– animals.

There is now compelling evidence that NEIL1 is not simply a so-called backup enzyme for

other DNA glycosylases as had been assumed originally, but plays an important role in the

prevention of cancer and metabolic syndrome-associated diseases. DNA glycosylase activity

of NEIL1, which is quite distinct from those of most other known DNA glycosylases, and its

potential role in NER makes it a unique DNA repair enzyme. In addition, NEIL1 may play a

primary role in transcription- and replication-coupled repair [278,300]. Moreover, the ability of NEIL1 along with NEIL3 in the prevention of mutagenesis in vivo has recently been demonstrated [293]. The expression of NEIL1 or NEIL3 in an E. coli fpg mutY nei mutant strain significantly reduced frequency of spontaneously occurring high G → T transversions.

A greater level of FapyGua has been observed in the fpg mutY nei mutant than in the wild

type strain. The expression of NEIL1 or NEIL3 in the mutant strain significantly reduced the

level of FapyGua, confirming the specificity of NEIL1 for in vivo repair of FapyGua and

providing the evidence that NEIL3 also recognizes FapyGua in vivo. The decrease in both the

mutation frequency and the level of FapyGua suggested that the G → T transversions resulted

from FapyGua to a great extent. In fact, FapyGua has been shown to cause this type of

41 mutations [357,359]. The lack of NEIL1 activity on 8-OH-Gua in vitro and in vivo provided the evidence that 8-OH-Gua does not contribute to the adverse effects of NEIL1 deficiency in vivo. Taken together all the works surveyed above, further studies are warranted on the role of the NEIL1 substrates, FapyAde, FapyGua, R-cdA and S-cdA in carcinogenesis and

metabolic syndrome observed in neil1–/– animals.

Any role of NEIL2 or NEIL3 in carcinogenesis has not yet been investigated in detail.

Three polymorphic variants of NEIL2 have been found in patients with familial colorectal

cancer [546]. Two of these variants have also been detected in multiple colorectal carcinomas,

but also in controls and an additional variant has been found in a patient, but not in controls

[516]. A number of NEIL3 variants have been detected in multiple colorectal adenomas with

only one variant being present in a patient, but not in controls [516]. Polymorphic variants of

NEIL2 and NEIL3 have been evaluated neither for their action nor for their association with

any disease [477]. Further work will be necessary to elucidate the role of NEIL2 and NEIL3

deficiencies in carcinogenesis.

5.1.3. NTH1

Nth1–/– mice have been generated to study the effect of deficiencies in the nth1 gene

[250,251,547,548]. Several studies showed no phenotypic abnormalities in nth1–/– mice

[547,548]. Mice have been viable and exhibited similarity to wt mice in early life, and have

shown no tumor formation or increased phenotypic aberrations. No carcinogenesis has been

observed in nth1–/–/ogg1–/– mice, either [548,549]. These findings indicate that other repair

enzymes may compensate for the lack of NTH1. In humans, altered expression of the nth1

gene has recently been detected in eight gastric cancer lines [550]. Reduced mRNA

expression of NTH1 and its abnormal cytoplasmic localization have also been observed in

some primary gastric cancers. These findings pointed to a possible involvement of NTH1

deficiency in gastric cancer. Furthermore, two polymorphisms have been found in the nth1

42 promoter region; however, no association between these polymorphisms and gastric cancer has been observed. Cytoplasmic localization of NTH1 has also been detected in some primary colorectal cancers[551]. Low expression of NTH1 in the nucleus due to cytoplasmic localization in cancer cells may lead to accumulation in the nucleus of oxidatively induced

DNA lesions that are the substrates of NTH1. In another study, nth1–/– male and female mice

developed pulmonary and hepatocellular tumors as they aged; however, double knockout

animals with nth1–/–/neil1–/ exhibited a dramatic increase in both cancer incidences [251].

5.2. Other BER proteins

5.2.1. APE1

In mammalian cells, APE1, which is the mammalian ortolog of E. coli exonuclease III

family of endonucleases, provides over 95% of the total AP endonuclease function

[304,305,552-555]. In addition, APE1 exhibits multiple functions including 3'-

phosphodiestrease, 3'-5' exonuclease, 3'-phosphatase and nucleotide incision repair (NIR)

activities, transcription and redox regulations, involvement in RNA repair and metabolism

[555-559]. Critical nature of APE1 functions is evidenced by early embryonic lethality in

mice with both deleted alleles of ape1, and by increased oxidative stress, spontaneous

mutagenesis and cancer incidences, and reduced survival of pups and embryos in APE1

heterozygous mice [560-564]. Other adverse effects are caused by depletion, inhibition or

downregulation of APE1, and defects in its activity; these include apoptosis [565,566],

sensitization to DNA-damaging agents [567], and loss of neuronal function and development

of neurodegenerative disease [568-570]. There is evidence for the association of APE1

polymorphisms with disposition to cancer [571,572]. Various variants of APE1 have been

identified in the human population with the potential to lead to variations in protein activity

or expression level [485,555,573-575] (see also the NCBI database, www.ncbi.nlm.nih.gov).

The majority of the amino acid substitutions is located in the repair nuclease domain of the protein, whereas the redox regulatory portion (REF-1) contains several of the substitutions

[575]. The most common Asp148Glu variant has been shown to have an allele frequency of ≈

0.38 [485,573,574]. The involvement of this variant has been suggested in cancer risk such as

melanoma [576-578], lung cancer [579], breast cancer [580], colorectal cancer [581-583] and

amyotropic lateral sclerosis (ALS) [584], and in ionizing radiation sensitivity [585]; however,

some other studies found no cancer susceptibility [586-590]. On the other hand, Asp148Glu

variant has been shown to possess a normal endonuclease activity [573]. Other variants

Leu104Arg, Glu126Asp and Arg237Ala exhibited a reduction in the endonuclease activity up

to ≈ 60%, while the activities of Gly241Arg and Gly306Ala variants were similar to that of

wt-APE1 [573]. Arg237Cys and Pro112Leu variants have been identified in 3 of 20

endometrial tumors [485,574]. The former displayed significant defects in several protein

activities including exonuclease function, possibly representing a reduced-function

susceptibility allele, while the latter had wt-APE1 activity, thus it seems unlikely to be involved in carcinogenesis [575]. Variants Gln51His, Ile64Val, Asp148Glu, Pro311Ser,

Gly241Arg and Ala317Val, which are not associated with human disease, exhibited no effect on the protein structure/function [575]. In terms of the intracellular localization, various

APE1 variants displayed a pattern similar to that of wt-APE1 [575]. The search in disease-

associated variations in ape1 in a large number of cancer cell lines found no novel APE1

amino acid substitutions with only common Asp148Glu and Gln51His variants observed

[575].

5.2.2. Pol β

Pol β is found in all vertebrate species and belongs to the X-family of DNA polymerases

[591-593]. In BER, following the action of APE1 that leaves one nucleotide gap with a 3'-OH and a 5'-terminal dRP, Pol β binds to the gap, performs DNA synthesis with its DNA

44 polymerase activity, filling in the gap, and removes the blocking dRP-moiety with its dRP

lyase activity, paving the way for DNA ligase 1 or a complex of X-ray repair complementing

protein 1 (XRCC1) and DNA ligase 3 to seal the resulting gap to complete the repair

[592,594-596]. The polymerase activity of Pol β is also necessary in long-patch BER

[597,598]. Pol β lacks 3' to 5' proofreading exonuclease activity and displays a moderate

fidelity with ≈ 1 error/3000 nucleotides synthesized, which is much higher than that of other

DNA polymerases [599-601]. This makes Pol β a relatively error prone polymerase.

Misinsertions introduced by Pol β during its DNA polymerase activity in BER may generate

mutations, indicating that Pol β may play a role in the etiology of cancer [602,603]. On the

other hand, Pol β appears to be essential for survival and fetal development as pol β‒/‒

animals exhibit embryonic lethality [592,604]. Mutations in pol β have been identified in

several human carcinomas and mouse lymphomas [605-611]. Variants of Pol β have been

found in approximately 30% of human tumors that also contained wt-Pol β; however, these

variants were not present in normal tissues of the same patients [602,606-608,610-615].

Approximately half of the tumors have been found to express Pol β variants with single

amino acid substitutions, whereas one third of the tumors expressed a deletion variant with

missing amino acids in the positions between 208 and 236 (Pol βΔ208-236) that likely results from alternative splicing [616,617]. This variant has been identified in several cancers

[610,612,613]; however, it has also been detected in normal tissues [612,616]. In addition, about 10% of the tumors have been shown to contain a truncated form of Pol β due to frame shift mutations. All these Pol β variants may cause defects in BER by synthesizing DNA with low fidelity, leading to genomic instability, thus to mutations leading to cancer. These mutations are not among the common polymorphisms found in pol β [574,602]. A strong association of overall survival with single nucleotide polymorphisms of pol β, i.e., the pol β

A165G and T2133G phenotypes, has been found in a large number of patients with

45 pancreatic adenocarcinoma [618]. In contrast, the homozygous variant genotype of pol β had

a significant protective effect on overall survival. The median survival time was 35.7 months

for patients with at least one of the two homozygous variant pol β 165 GG and 2133 CC

genotypes, whereas those patients carrying the pol β 165 AA/AG and 2133 TT/TC genotypes

had a median survival time of 14.8 months.

A Pol β Lys289Met variant along with wt-Pol β has been identified in a human colon

carcinoma [612]. The expression of this variant in mouse cells led to a significant increase in

the mutation frequency with the mutational spectrum being different from that of wt-Pol β

[619]. In addition, the Lys289Met variant displayed a lower fidelity than wt-Pol β and

misincorporated nucleotides during BER. The frequency of C → G transversion mutations in

cells with Lys289Met variant expression was much greater than those in wild type cells.

Another Ile260Met variant of Pol β has been identified in a prostate carcinoma [606]. This

variant has been shown to have a mutational spectrum different from that of wt-Pol β and to possess a sequence-specific mutator activity [620,621]. Expression of the Lys289Met and

Ile260Met variants in mouse cells has been shown to result in permanent cellular

transformation, indicating that these variants induce mutations during BER by aberrant gap filling, which is likely to be different from the function of wt-Pol β [620]. These results led to the suggestion that the mutations may occur in key growth control cells leading to cellular transformation. By genotyping of pol β, two exonic germline variants Arg137Gln and

Pro242Arg of Pol β have been found with allele frequencies of 9% and 3%, respectively

[622]. The Arg137Gln variant displayed a lower DNA polymerase activity than wt-Pol β

[623]. Patients with lung cancer who carry the Pro242Arg variant exhibited a decrease in survival [624]. A gastric cancer-associated variant Glu295Lys of Pol β has been shown to have no DNA polymerase activity, leading to unfilled gap in BER and inducing cellular transformation [625]. Another gastric cancer-associated Leu22Pro variant exhibited DNA

46 polymerase activity, albeit lesser than that of wt-Pol β, but lacked dRP lyase activity and

could not support BER [626]. The studies of Pol β mutants lent credence to the mutator

phenotype hypothesis [15] and suggested that BER is a tumor suppressor mechanism [603].

Pol β has been shown to form complexes with DNA lesions such as mutagenic 8-OH-Gua

in the confines of its active site [592,627]. It preferentially inserted the correct base cytosine

opposite 8-OH-Gua rather than adenine, although this depended on the sequence context and

the insertion of adenine also occurred [628,629]. However, 8-OH-Gua has been the only

DNA lesion investigated so far. Pol β may also form complexes with other oxidatively

induced DNA base lesions, causing misinsertions and subsequent mutations. Taken together,

accumulated evidence clearly points to a role of Pol β and its variants in the etiology of

cancer.

5.3. DNA lesions and DNA repair proteins as biomarkers

5.3.1.DNA lesions as biomarkers

Evidence accumulated over several decades suggests that oxidatively induced DNA

lesions may be used as potential disease biomarkers and cancer risk assessment. Elevated

levels of such DNA lesions in cancerous tissues and in BER enzyme-knockout animals or in

animals that developed cancer upon exposure to environmental toxins, as discussed above,

strongly support this notion. For this purpose, however, accurate measurements of oxidatively

induced DNA lesions in tissues would be absolutely necessary. Mass spectrometry-based

assays using stable isotope-labeled internal standards have been developed for such

measurements and successfully applied over the past two decades or so. It is out of the scope

of this article to review the entire literature on this subject. The reader is referred to reviews

and other articles in this field (see e.g., [105,121,124,251,630]). Noninvasive procedures to

collect samples such as urine and the measurement of oxidatively induced DNA lesions

47 therein drew significant attention from many laboratories. A large number of studies have been conducted to measure these lesions as non-invasive biomarkers for diagnosis, early detection and therapy monitoring, and also for epidemiological studies. Various protocols have been developed to measure DNA lesions in urine. These included the use of mass spectrometry, HPLC with electrochemical detection and enzyme-linked immunosorbant assay

(ELISA). First studies measured thymine glycol and 2'-deoxythymidine glycol (dT glycol)

[631], and 8-hydroxy-2'-deoxyguanosine (8-OH-dG) [632]. Subsequently, 8-OH-dG and its free base 8-OH-Gua, albeit to a lesser extent, have mainly been measured [633-656]. Other lesions FapyGua, 8-OH-Ade and 5-OH-Ura have also been found in urine [633,644], although much less attention was paid to these lesions than 8-OH-dG and 8-OH-Gua. There have been significant differences between the measurements in different laboratories and about the source of these DNA lesions in urine. A European Standards Committee on Urinary

(DNA) Analysis (ESCULA) has been established to help reach a consensus between results

in different laboratories in Europa, USA and Asia [649]. A recent comprehensive study

involving a large number of laboratories investigated both human and methodological factors

influencing measurements of 8-OH-dG in urine using ELISA and chromatographic

techniques including mass spectrometry [656]. Chromatographic techniques performed better

than ELISA in terms of high agreement across urine samples from different subjects. As for

the source, several studies showed that diet and cell death do not contribute to the appearance

of DNA lesions in urine [635,639,645,647,649,657,658]. BER has been suggested to be

responsible for the presence of 8-OH-Gua in urine because of the specificity of OGG1 of

BER for this lesion [640]. However, DNA glycosylases such as OGG1 would not remove 8-

OH-dG from DNA because they are specific for modified DNA bases, not for modified 2'-

deoxynucleosides [217]. NER is not likely to be responsible for 8-OH-dG in urine, either,

because no oligonucleotides containing 8-OH-dG have been identified in urine [639]. Repair

48 in the nucleotide pool appears to be a major source of 8-OH-dG and possibly that of dT glycol and other modified 2'-deoxynucleosides [657,659]. MTH1 is the best characterized enzyme in the nucleotide pool that hydrolyzes 8-OH-dGTP to 8-OH-dGMP and thus prevents its incorporation into DNA [197,660]. Dephosphorylation of the latter would give rise to 8-

OH-dG, which might be removed from cells and ultimately appear in urine. Still, the precise nature of the presence of 8-OH-dG in urine remains unclear [661].

Recently, the presence of R-cdA and S-cdA has been discovered in human urine [127]. A methodology has been developed to simultaneously measure these compounds and 8-OH-dG

in urine using LC-MS/MS with isotope-dilution. Since 8,5'-cyclopurine-2'-deoxynucleosides are repaired by NER, not by BER [108,294-296], their presence in urine has been suggested to result from this repair pathway[127]. However, the repair of their triphosphates in the nucleotide pool followed by the excretion into urine as described above cannot be excluded, either. In an application to a disease state, R-cdA and S-cdA have been found in urine of atherosclerosis patients at significantly greater concentrations than in that of healthy individuals [128] (Fig. 8). The statistical difference was highly significant. 8-OH-dG has been simultaneously measured. Its concentrations in urine of patients were also significantly greater than those in controls. However, the significance of the data for R-cdA and S-cdA was greater than that for 8-OH-dG. The concentrations of R-cdA and S-cdA in urine were about two magnitudes of order less than that of 8-OH-dG. 8-OH-Gua has also been measured by another method. No statistical significance has been found between the levels of 8-OH-Gua in controls and patients, indicating that this compound may not be a reliable biomarker.

Simultaneous measurement of R-cdA and S-cdA along with 8-OH-dG in urine would be more advantageous for reliable results than the measurement of one lesion only. Taken together, the accurate and reproducible measurement of R-cdA and S-cdA in human urine, and their extraordinary chemical stability and clear origin render these compounds ideal as potential

49 disease biomarkers in urine for early detection, testing of drugs, monitoring the therapy and epidemiological studies. Further studies on the use of these unique compounds as biomarkers for cancer and other diseases are warranted.

5.3.2. BER proteins as biomarkers

Ionizing radiation and most chemotherapeutic agents kill tumor cells by damaging DNA.

However, their effectiveness may be influenced by the efficiency of DNA repair capacity in

tumors [449,454,480,662-664]. Overexpression of DNA repair proteins that may increase the

DNA repair capacity is common in cancer. Since malignant tumors possess increased level of

oxidatively induced DNA damage [112,113,116,117,457-470], the overexpression of DNA

repair proteins may be required to encounter high level of DNA damage in tumors. Increased

rate of DNA damage and subsequent mutations may lead to genetic instability and cell death

late in tumor evolution. However, rapidly developing tumors that overexpress DNA repair

proteins may have an evolutionary advantage for survival and thus develop greater DNA

repair capacity than normal tissues. Effective DNA repair in tumors that removes DNA

lesions before they become toxic is a major mechanism for resistance to therapy, and may

affect the outcome of therapy and thus determine the patient survival. In the normal

population, cancer susceptibility may also be influenced by repair capacity [449]. Increase in

DNA repair capacity allows cancer cells to develop multi-drug resistance. Accumulated

evidence strongly suggests that DNA repair capacity might be a predictive biomarker of

patient response to therapy [480,663]. Determination of the overexpression or

underexpression of DNA repair proteins in normal and cancer tissues might help predict and

guide treatments. In this context, BER proteins have emerged as biomarkers for prediction of

tumor response and prognosis of treatment outcome [449,454,479,480,555,572,663,665-667].

5.3.3. BER proteins as therapy targets

Since BER proteins are responsible for the repair of a multiplicity of oxidatively induced

DNA lesions, they would be logical targets for inhibition to effectively achieve therapies to

overcome the resistance of tumors to treatment, effecting apoptosis or cell death instead of

DNA repair [555]. Development of inhibitors of BER proteins and other DNA repair proteins

for combination therapy or as single agents for monotherapy will help selectively kill tumors.

The achievement of this goal will potentially lead to personalized cancer therapy [480]. In

this respect, a thorough understanding of DNA repair pathways and functions of proteins

involved will be of fundamental importance for the use of DNA repair proteins as biomarkers

and targets for improving therapy of cancer [668].

Success with the inhibitors of poly(ADP ribose) polymerase 1 (PARP1), which is the major enzyme of the members of the PARP superfamily and involved in BER, brought to attention the inhibition of BER proteins in tumors as a potential and promising concept for cancer therapy (reviewed in [667]). Inhibition of PARP activity preventing the repair of methylation-induced DNA strand breaks and generating cytotoxicity gave the impetus for development of DNA repair inhibitors for cancer therapy [669]. In particular, high efficacy of

PARP inhibitors as single agents for monotherapy has been demonstrated in patients with

inherited breast and ovarian cancers that contain deficient brca1 and brca2 genes [670,671].

About a decade ago, PARP inhibitors started entering clinical trials [672,673]. Clinical trials

are in progress now and numerous PARP inhibitors that have been developed by the

academia and the industry are being tested. There is a wealth of literature dealing with PARP

inhibitors and their use in cancer therapy. More information can be found in several recent

review articles [454,663,667,674,675].

APE1 is another BER protein for which intense efforts are being underway worldwide to

find inhibitors of its activities [555]. Overexpression of APE1 has been observed in multiple

51 human cancers and associated with resistance to chemotherapy and radiation therapy, and with poor survival[555,571,572,676]. In this context, numerous studies provided the evidence for the expression and also subcellular localization of APE1 to be of great predictive and prognostic value. Thus, strict nuclear localization of APE1 has been found to associate with good prognosis, whereas its combined nuclear and cytoplasmic localization correlated with poor survival. However, elevated levels of APE1 with predominantly nuclear localization have also been found in various cancers [677-681]. These contradictory observations indicate that APE1 subcellular localization may vary among cancer types. Mounting evidence suggests that inhibition of APE1 functions increases cellular sensitivity to DNA-damaging agents. As discussed above, this protein is not only a DNA repair enzyme, but also exhibits other important functions and aspects. Because of these reasons, APE1 is generally accepted as being an excellent target for development of inhibitors as anticancer agents for monotherapy and/or to enhance the efficacy of present drugs and ionizing radiation in cancer therapy [555,572,682]. Since 2005, thousands of structurally diverse compounds have been screened to find inhibitors of APE1 using high-throughput screens [471,555,572,682-686].

Inhibitors that block the DNA repair function of APE1 work either by binding to DNA to

inhibit its endonuclease activity or by binding to APE1 to inhibit its AP site activity. Some

inhibitors of APE1 endonuclease function are in Phase I trials [555]. Inhibition of the redox

activity of APE1 blocks various cellular pathways including multiple tumor signaling

pathways involved in cancer development and survival [687,688]. There are naturally-

occurring or synthesized inhibitors of the redox activity of APE1 that are being tested or are

in development [555].

Inhibiting Pol β may also be of importance for cancer therapy. Thus, greater levels of Pol

β have been found in breast, colon and prostate adenocarcinomas than in adjacent normal

tissues[689]. Overexpression of Pol β, which decreases the fidelity of BER[690], has been

52 associated with genetic instability, cellular transformation, hyperplasia and carcinogenesis as

well as resistance to therapy with DNA-damaging agents [690-695]. In fact, Pol β has been

shown to be upregulated in the presence of increased DNA damage [696,697]. Knocking

down Pol β has increased cellular sensitivity to DNA-damaging agents [698,699]. Efforts are

being underway to develop small molecule inhibitors of Pol β activities. One of the first

inhibitors exhibited inhibition of both lyse and polymerase activities of Pol β [700].

Numerous other inhibitors have been developed with a variety of potencies and specificities

[700-703]. Many of them lacked necessary characteristics to become cancer-specific drugs.

However, some of these compounds were promising and are in preclinical studies [555]. One

compound enhanced the ability of DNA-alkylating agent temozolomide (TMZ), which has

been used successfully for the treatment of some cancers [704,705], to impair the growth of

colon cancer cells [702]. Another study tested thousands of small molecules targeting Pol β

for chemotherapeutic intervention of colorectal cancer [706]. A compound with a low

molecular weight has been identified to be a potential inhibitor of Pol β activities. However,

it did not affect the activity of other BER proteins. Combining this small molecule inhibitor

with TMZ effectively blocked the growth of colon cancer cells in vitro and caused antitumor

activity in vivo. Taken together, more efforts may be needed to study mechanistic aspects of

Pol β activities and to develop high-throughput screening assays for the search of inhibitors

of this important BER protein to be used as anti-cancer drugs.

Flap endonuclease I (FEN1) is another BER protein, for which there are efforts to develop

inhibitors. This protein is involved in long-patch BER [707-712]. Following the action of

APE1, Pol δ/ε introduces two to eight deoxynucleotides past the AP site generating an

overhang polydeoxynucleotide with two to ten deoxynucleotides (5' flap) and the dRP. FEN1

removes this 5' flap and then DNA ligase I seals the remaining nick completing the repair.

There are other functions of FEN1 as well [711-714]. Efficient activities of FEN1 are

53 essential for the maintenance of the genomic integrity [715]. Overexpression of FEN1 has

been observed in numerous cancers, suggesting a role for FEN1 in tumor progression and

development, and therapy resistance [716-722]. Consequently, FEN1 might be a potential target for anti-cancer treatment. Assays have been developed to find inhibitors of FEN1

[723,724]. Several small molecule inhibitors with N-hydroxyurea-based compounds among them have been identified with the potential to serve as anti-cancer drugs [723-725]. FEN1 has also been found to be a potential target for synthetic lethality [726]. Some N-hydroxyurea series of FEN1 inhibitors are in early preclinical trials [555]. Developed assays may facilitate the developments of novel inhibitors of FEN1 to be used in cancer therapy.

Recent work suggested that targeting MTH1 of the nucleotide pool repair protein might

be beneficial in cancer therapy [204,727]. This protein dephosporylates modified 2'-

deoxynucleoside triphosphates in the nucleotide pool, preventing their incorporation into

DNA during DNA replication [197-202]. Modified 2'-deoxynucleoside triphosphates in the

nucleotide pool have been shown to be a significant contributor to genetic instability in

mismatch repair-deficient cells [728]. Overexpression of MTH1 has been observed in many cancers [200,203,729,730]. MTH1 activity has been found to be greater in tumors than surrounding normal tissues from non-small-cell lung cancer patients [203]. The level of 8-

OH-Gua was also lower in tumors than in surrounding normal tissues, suggesting DNA repair capacity of tumors may be greater that that of normal tissues. This is in agreement with reported lower levels of oxidatively induced DNA lesions and ethano-DNA adducts in cancerous tissues compared to surrounding non-cancerous tissues [123,481,482]. Cancer cells have been shown to require MTH1 for efficient survival by avoiding incorporation of modified 2'-deoxynucleoside triphosphates into their DNA during replication [204]. This finding suggested that MTH1 might be an excellent target for inhibition in cancer treatment.

Small molecule inhibitors of MTH1 have been found by screening compound libraries and

54 successfully applied to suppress tumor growth in mice with different cancers, validating

MTH1 as a novel anticancer target in vivo. Clinical trials of MTH1 inhibitors may be

performed in the future for drug development.

Despite the successes with other BER proteins, the development of inhibitors for DNA

glycosylases has been lagging. Recently, a study has been conducted to identify gene-specific

pathways that would serve as synthetic lethal partners with DNA glycosylases as the targets

for cancer therapy using chemotherapeutic agents that function through depletion of cellular

nucleotide pools [731]. Thymidylate synthetase (TS) plays the key role in the synthesis of 2'-

deoxythymidine (dT) by producing dTMP from dUMP [732,733]. Inhibitors that target the

TP pathway are widely used in the treatment of many cancers [734-737]. These are mainly

folate-based analogues with some nucleotide-based compounds and cause toxicity by

depletion of dTTP that inhibits DNA replication and increases incorporation of dUTP into

DNA. The combined siRNA-mediated reduction of NEIL1 and the treatment with four TS

inhibitors dramatically increased toxicity in an osteosarcoma cell line [731]. Depletion of

NEIL1 or OGG1 alone had no effect on toxicity. Moreover, loss of NEIL function has been shown to be synthetically lethal with the disruption of the Fanconi anemia DNA repair pathway. This work identified NEIL1 as the key BER protein in the repair of DNA following inhibition of TS pathway. This means that inhibition of NEIL1 may enhance clinical responses to TS pathway inhibitors. A recent work developed a strategy to discover inhibitors of DNA glycosylases with the goal of finding small molecules to be used in combination

therapy [738]. A high-throughput, fluorescence-based assay has been developed that uses incision of oligodeoxynucleotides with a single modified DNA base to detect small molecule inhibitors of DNA glycosylases with an associated AP lyase activity. NEIL1 has been used as the proof-of-principle glycosylase for this purpose. Oligodeoxynucleotides contained both Sp and Gh that are oxidation products of 8-OH-Gua (see above). As discussed above, NEIL1 is

55 specific for excision of FapyAde and FapyGua from DNA in vitro and in vivo, and also

efficiently excises Sp and Gh from oligodeoxynucleotides, but it does not act on 8-OH-Gua.

The high-throughput assay has been used to screen small molecule libraries with a large

number of compounds for inhibitors of the combined glycosylase/AP lyase activities. There

were a number of purine analogues among top hits of these screens. Since FapyAde and

FapyGua are physiological substrates of NEIL1, inhibition by purine analogues of NEIL1

activity for these compounds has been tested using DNA samples that contained multiple

oxidatively induced lesions. Fig. 9 illustrates the determination of the inhibition of NEIL1

activity for FapyAde and FapyGua by six purine analogues. An efficient activity of NEIL1 on

FapyAde and FapyGua has been observed in agreement with previous studies

[250,251,278,290,292]. Inhibitors P2, P6, P7 and P8 significantly (p < 0.005) inhibited the

excision of the two lesions, whereas P11 had a lesser but significant effect on FapyAde

excision and did not inhibit FapyGua excision. Taken together, this study may form a foundation for potential drug discovery for cancer therapy for the entire family of DNA glycosylases that are specific for a variety of oxidatively induced DNA base lesions. As was done in the study of NEIL1 inhibitors, simultaneous measurement of biological substrates of these enzymes using DNA samples with multiple DNA lesions will facilitate screening of

various potential inhibitors with the goal of discovering suitable drugs for inhibition of DNA

glycosylases in cancer therapy.

As Kelley has stated [663], knowledge of repair proteins’ overexpression or

underexpression in cancers may help predict and guide development of treatments,

potentially yielding the greatest therapeutic response. Thus far, the expression of DNA repair proteins in cells and tissues including clinical samples have been estimated by semi-

quantitative immunochemical methods. To be used as reliable biomarkers in cancer,

expression levels of DNA proteins must be accurately measured in tissues by proper chemical

56 and physical techniques. Mass spectrometry is the most suitable technique of choice for this

purpose and is being used worldwide for the measurement of proteins in the field of

proteomics. The application of this technique would be essential for positive identification

and accurate quantification of DNA repair proteins in human tissues. Recently, our laboratory

has developed methodologies that use mass spectrometry with isotope-dilution for the

measurements of DNA repair proteins [739-742]. Full length 15N-labeled analogues of human

OGG1, NEIL1, NTH1, Pol β and APE1 have been produced and purified to be used as internal standards for the accurate quantification of these proteins by LC-MS/MS. Thus,

APE1 has been identified and quantified in human tissues, cultured human cells and mouse liver. Efforts are now being made to extend this work to measure expression levels of DNA proteins in different types of human cancer tissues from patients and in tissues from disease- free individuals. We believe that such measurements will be of fundamental importance for the determination of DNA repair capacity, the use of DNA repair proteins as biomarkers and

the development of DNA repair inhibitors.

6. Conclusions

There is mounting evidence for an important role of oxidatively induced DNA damage

and its cellular repair in the etiology of cancer. Great strides have been made in the

understanding of various mechanisms of DNA repair since its discovery five decades ago.

Discovery of DNA repair proteins and elucidation of their functions paved the way for

understanding of the contribution of aberrant DNA repair, and mutations and polymorphisms

in DNA repair genes to carcinogenesis. The finding that increased DNA repair capacity in tumors causes resistance to therapy gave impetus to development of inhibitors of DNA repair to increase the efficacy of therapy. Success of the application of certain inhibitors as drugs in the therapy of some cancers is a very promising development. It will be important to continue

57 the development of inhibitors of proteins in numerous DNA repair pathways as drugs to be used to enhance the efficacy of current cancer therapy and to eradicate cancer. A thorough understanding of the DNA repair pathways in carcinogenesis and overexpression or

underexpression of DNA repair proteins in tumors in comparison to normal tissues will be of

utmost importance in improving therapy and achieving the best therapeutic response.

Conflict of interest statement

The author declares that there are no conflicts of interest.

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Figure Legends

Fig. 1. Structures of oxidatively induced DNA base lesions in DNA.

Fig. 2. Structures of the lesions derived from the sugar moiety of DNA.

Fig. 3. Tandem lesions identified in vivo [151,159,160,178-180].

Fig. 4. Levels of FapyAde and 8-OH-Gua in wt-, nth1‒/‒, ogg1‒/‒ and nth1‒/‒/ogg1‒/‒ mice

(data from [250]).

Fig. 5. Mispairings of oxidatively induced DNA lesions (redrawn from the data in

[297,349,356,358,367,370,399,430,434]).

Fig. 6. Left: Formation of C → T transition mutations from the mispairing of 5-OH-Cyt or 5-

OH-Ura with Ade [348,380,386,395-397]; Right: Formation of G → A transition mutations from the mispairing of S-cdG with Thy [297,430].

Fig. 7. Tumor incidence in neil1‒/‒, nth1‒/‒ and neil1‒/‒/nth1‒/‒ mice. Upper graphs: Lung

adenoma, adenocarcinoma. Lower graphs: Liver hepatocellular carcinoma, nodular

hyperplasia, severe dysplasia. Number (n) of animals: Males, neil1‒/‒, n = 25; nth1‒/‒, n = 52;

neil1‒/‒/nth1‒/‒, n = 43. Females, neil1‒/, n = 18; nth1‒/‒, n = 54; neil1‒/‒/nth1‒/‒, n = 29 (data

from [251]).

Fig. 8. Levels of R-cdA (A), S-cdA (B) and 8-OH-dG (C) in urine of control individuals (1)

and atherosclerosis patients (2) (data from [128]).

Fig. 9. Inhibition of the activity of human NEIL1 by small molecule inhibitors (data from

[738]).

107

guanine-derived products O O

N N NH NH O O N O O N N HN NHCHO N O N NH2 N NH2 HN H OH H2N HO H2N H O H O H2N N N H2N N NH2 H2N NH2 O H N NH2 N H H H H H H H H - , ------, , - - 8 hydroxyguanine 2 6 diamino 4 hydroxy 2,5 diamino 4H 2 2 4 triamino OH H OH H - - - - 5 formamidopyrimidine imidazol 4 one 5(2H) oxazolone (5'R)-8,5'-cyclo-2'-deoxyguanosine (5'S)-8,5'-cyclo-2'-deoxyguanosine

8-hydroxyguanine-derived products

H O N H O N O N NH N O H2N H N N H2N N H O H H H - spiroiminodihydantoin 5 guanidinohydantoin

adenine-derived products NH2 NH2

N N N N

NH2 NH2 NH2 N N N N N NHCHO N N N N HO H OH H H O HO O NH H H H H H N N H N 2 HO N N H H H H H H ------8 hydroxyadenine 4,6 diamino 5 form 2 hydroxyadenine OH H OH H ami o rimi ine d py d (5'R)-8,5'-cyclo-2'-deoxyadenosine (5'S)-8,5'-cyclo-2'-deoxyadenosine

cytosine-derived products

NH2 NH2 NH2 NH2 NH2 H H H OH N OH N OH N N OH N H OH H H OH H O N H O N O N O N H O N H H H H H H ------cytosine glycol 5 hydroxycytosine 5,6 dihydroxy 5 hydroxy 6 5,6 dihydrocytosine cytosine hydrocytosine

O O O O O O O H O H H OH H O O HN OH HN HN OH HN HN HN HN OH HN H OH H OH H O H H O N H N O N H O N OH O N O O N O N O O N H H H H H H H H H H ------, - uracil glycol 5 hydroxyuracil 5 hydroxy 6 5 hydroxy alloxan dialuric acid isodialuric acid 5 6 dihydrouracil hydrouracil hydantoin

thymine-derived products

O O O O O O CH3 CH3 CH OH CHO CH HN 2 HN HN 3 HN OH HN OH HN H H OH CH3 H O O N H O N H O N OH O N H N H H O N H H H H H H

- rox - - ro- m ne co - rox me - - orm urac - rox - - , - ro m ne 5 h y d y 6 hyd thy i gly l 5 ( h y d y thyl) 5 f yl il 5 hyd y 5 5 6 dihyd thy i thymine uracil methylhydantoin

Figure 1 sugar products released from DNA

CH HO CH HO CH 3 O 2 O 2 O O O O H H H O H H H O H H H H H H H

OH H H H OH H OH H

2,5-dideoxypentos-4-ulose 2,3-dideoxypentos-4-ulose 2-deoxypentos-4-ulose 2-deoxytetradialdose

sugar products bound to DNA as end group

CH3 ~ HO CH2 O O P O CH CH O H H 2 O 2 O H H H O O H O H H ~ O H H O ~H H P H O ~H P OH H O H P 2,5-dideoxypentos-4-ulose 2-deoxypentos-4-ulose 2-deoxypentos-4-ulose 2-deoxypentos-4-ulose

H ~

~ O C base O O O P O CH2 H H P O CH2 H H O H H O H H H O H H O ~H O O ~ H H H P P 2,3-dideoxypentos-4-ulose 2-deoxytetradialdose 3'-phosphoglycolate 5'-aldehyde

sugar products within DNA ~ ~ ~

P O CH2 P O CH2 P O CH2 O OH OH O H H H H COOH H H H H H O ~H O ~H O ~ O P P P

2-deoxypentos-4-ulose 2-deoxypentonic acid erythrose

Figure 2 O N O HN N HN O OH DNA H2N N N ~ DNA H2N N N ~ HN CH2 P O CH2 O O P O CH2 O NH2 H H H H2C O N NH H H H N H H CH2 O H H DNA H N O O H - - H N O HN CH CHO P O CH2 P O CH O 2 O H H H H thymine-tyrosine cross-link H H H H O ~ H O ~ H P DNA P DNA

Gua[8,5-Me]Thy Gua[8,5]Cyt

Figure 3 8-OH-Gua 4.0 FapyAde

3.0 DNA bases) 6 2.0

1.0 level (lesions/10

0.0

Figure 4 H N H H H N N O N H2N A N N N D H N N H N O D NA 8-OH-Gua•Ade

H H H N N H H H H O N O C N N N O H2N N H A N H N H N N N N D HN N H H N A C NH N NH D D O H N N N H 2 DNA A FapyGua•Ade FapyGua•Ade

H N N H H H O H O C N N H NH2 N H N N N N N H N N H N H D A N N A O N D NH N H N H N H N DNA DNA H

8-OH-Ade•Gua FapyAde•Ade

H H O N H N N H H N H N N N A N N D D NA O H

5-OH-Cyt•Ade (anionic imino tautomer)

D N A O H H CH3 D NA CH O H N H O 3 N N O H N O H H H H O N N N O N N H H N D O N H O H N H N A DNA O H A N D O N N H D H H A NH2 H H

S-cdG•Thy S-cdA•Thy

Figure 5 O

N NH

O NH2 N N NH2 OH OH H HN N C HO O H H O H H N O N H H H H OH H - - ' - , '- - '- 5 hydroxyuracil 5 hydroxycytosine (5 S) 8 5 cyclo 2 deoxyguanosine

G = C* *G=C

replication replication

A = C* *G =T

replication replication

A = T A=C* *G=T A = T

C → T G → A transition transition

Figure 6 males females 80 74.4% 50 41.4% 40 60

30 40 20

20 12% 10 cancer incidence (%) cancer incidence (%) cancer incidence 3.7% 1.9% 0% 0 0 -/- -/- -/- -/- -/- -/-

nth1 nth1 neil1 -/- /nth1 neil1 -/- /nth1

neil1 neil1

males females 50 46.5% 20 17%

40 15 13% 30 11.1% 10 20 16% 15.4% 5 10 12% 12% cancer incidence (%) cancer incidence (%) cancer incidence

0 0 -/- -/- -/- -/- -/- -/-

nth1 nth1 neil1 -/- /nth1 neil1 -/- /nth1

neil1 neil1

Figure 7 A 0.06 p < 0.0001 B 0.06 p < 0.0001 C 6 p = 0.0008

0.04 0.04 4 (nmol/mmol) (nmol/mmol) 0.02 (nmol/mmol) 0.02 2 R -cdA/creatinine S -cdA/creatinine 8-OH-dG/creatinine

0.00 0.00 0 1 2 1 2 1 2

Figure 8 NH2 O NHCHO NHCHO 75 N 160 HN NH H N 2 H2N N NH2 FapyAde FapyGua 60 120 DNA bases) DNA bases) 45 6 6 80 30

40 15 level (lesions/10 level (lesions/10

0 0

NEIL1 NEIL1 control control NEIL1NEIL1 + P2NEIL1 + P6NEIL1 + P7 + P8 NEIL1NEIL1 + P2NEIL1 + P6NEIL1 + P7 + P8 NEIL1NEIL1 + P11 + P19 NEIL1NEIL1 + P11 + P19 NEIL1 + DMSO NEIL1 + DMSO

S N

HN N HN S HN N N N N N N N N N

N N N N N N N N N H H H H H H

P2 (NCGC00188618) P6 (NCGC00188616) P7 (NCGC00188617) F

O N HN F NH HN O HN N N N N N N N N N F N N S N N N O O N N N H H H F H

P8 (NCGC00188619) P11 (NCGC00182914) P19 (MLS001126460)

Figure 9