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Contents lists available at ScienceDirect

Archives of Biochemistry and Biophysics journal homepage: www.elsevier.com/locate/yabbi 5 6

Oxidatively generated base damage to cellular DNA by hydroxyl radical and one-electron oxidants: Similarities and differences

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a

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b

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Institut Nanosciences et Cryogénie, CEA/Grenoble, F-38054 Grenoble Cedex 9, France Département de Médecine Nucléaire et Radiobiologie, Faculté de Médecine des Sciences de la santé, Université de Sherbrooke, Sherbrooke, Québec J1H 5N4, Canada

a r t i c l e

1 2 3 7 14 15 16 17 18 19 20 21 22 23 24 25 26

Jean Cadet a,b,⇑, J. Richard Wagner b

i n f o

Article history: Received 24 February 2014 and in revised form 23 April 2014 Available online xxxx

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Keywords: Oxidized nucleobases Radical cations Tandem base lesions Intrastrand cross-links Interstrand cross-links DNA–protein cross-links Clustered DNA damage

a b s t r a c t Hydroxyl radical (OH) and one-electron oxidants that may be endogenously formed through oxidative metabolism, phagocytosis, inflammation and pathological conditions constitute the main sources of oxidatively generated damage to cellular DNA. It is worth mentioning that exposure of cells to exogenous physical agents (UV light, high intensity UV laser, ionizing radiation) and chemicals may also induce oxidatively generated damage to DNA. Emphasis is placed in this short review article on the mechanistic aspects of OH and one-electron oxidant-mediated formation of single and more complex damage (tandem lesions, intra- and interstrand cross-links, DNA–protein cross-links) in cellular DNA arising from one radical hit. This concerns DNA modifications that have been accurately measured using suitable analytical methods such as high performance liquid chromatography coupled with electrospray ionization tandem mass spectrometry. Evidence is provided that OH and one-electron oxidants after generating neutral radicals and base radical cations respectively may partly induce common degradation pathways. In addition, selective oxidative reactions giving rise to specific degradation products of OH and one-electron oxidation reactions that can be used as representative biomarkers of these oxidants have been identified. Ó 2014 Published by Elsevier Inc.

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Introduction

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Major attention has been given during the last four decades to the elucidation of the mechanisms of oxidation of nucleic acids because of its implication in aging [1,2] and several pathologies including cancers [3–5], atherosclerosis [6] and neurological diseases [7,8]. Comprehensive oxidative pathways are now available concerning the formation of several classes of DNA lesions including single oxidized nucleobases, tandem base modifications, intraand inter-strand cross-links [9–16] together with oligonucleotide single strand breaks and 2-deoxyribose oxidation products [17– 19]. Reliable information on both the qualitative and quantitative aspects of generation of several oxidatively produced lesions in cellular DNA has been gained from the development of accurate biochemical and chemical methods including the modified comet assay [20,21] and high performance liquid chromatography coupled with electrospray tandem mass spectrometry (HPLC/ESI-MS/ MS) or eventually MS3 detection [22–24]. The availability of these powerful analytical tools also allows for the assessment of the kinetics of repair of dedicated modifications [21,25] and studies

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toward the protection against oxidative reactions to DNA afforded by exogenous compounds [26]. Another relevant feature of HPLC– ESI-MS/MS measurements deals with the unambiguous identification of oxidatively generated base damage to cellular DNA. As striking findings, it was found that several DNA degradation products detected in cells [27–35] are identical to those previously characterized in model studies involving hydroxyl radical (OH),1 singlet oxygen (1O2) or one-electron oxidants in aerated aqueous solutions [9–13,16,36–38]. The latter conditions where molecular oxygen (O2) is able to react efficiently with most base radicals generated by either OH or one-electron oxidants appear to constitute suitable model systems for mimicking the cellular environment. This applies to the reactivity of oxidants towards DNA components and subsequent chemical reactions thus initiated that give rise to final decomposition products. The present review article is aimed at rationalizing the formation of oxidatively generated damage unambiguously detected in cellular DNA as the result of one initial radical hit. It appears that OH and one-electron oxidants exhibit similarities and also differences in the way they trigger the decomposition of nuclear DNA in cells. This survey does not include oxidation studies of cellular DNA by 1O2, a ROS produced by either

⇑ Corresponding author at: Institut Nanosciences et Cryogénie, CEA/Grenoble,

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F-38054 Grenoble Cedex 9, France. Fax: +33 4 38 78 50 90. E-mail address: [email protected] (J. Cadet).

1 Abbreviations used: OH, hydroxyl radical; 1O2, singlet oxygen; HOCl, hypochlorous acid; NO, nitric oxide.

http://dx.doi.org/10.1016/j.abb.2014.05.001 0003-9861/Ó 2014 Published by Elsevier Inc.

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type II photosensitization mechanism [39,40] or the reaction of hypochlorous acid (HOCl) reaction with H2O2 [41], both of which have been shown to predominantly generate 8-oxo-7,8-dihydroguanine (8-oxoGua) [12,27,42,43]. In addition, we will not discuss the reactions of HOCl, resulting from the oxidation of chloride ion with H2O2 catalyzed by myeloperoxidase in activated neutrophils [44], which leads to the highly specific chlorination of bases in cellular DNA with the following decreasing order of efficiency: cytosine > guanine > adenine [45,46]. The formation of oxidatively generated clustered lesions to DNA including double strand breaks are considered as hallmarks of radiation-induced damage that arise from several radical hits and excitation events within one or two DNA helix turns [47,48] are also excluded in the present review. Hydroxyl radical and one-electron oxidants in cells

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The two main oxidative pathways of cellular DNA considered in the present survey are those induced by OH and several exogenous one-electron oxidants including type I photosensitizers and high intensity UV laser pulses.

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Endogenous sources

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The two main endogenous sources of oxidative radical reactions having the ability to damage cellular DNA consist of OH and oneelectron oxidants. In contrast superoxide radical (O2) and H2O2 that are major ROS generated by respiratory burst and phagocytes [49] show very low reactivity with the nucleobases and 2-deoxyribose moiety [50]. The only known reaction mediated by O2 involves the one-electron reduction and addition to C5 of highly oxidizing guanine radical Gua(–H) that arises from deprotonation of the guanine radical cation [10,51]. It has also been reported that H2O2 is able to undergo electrophilic addition to N1 of adenine giving rise to the formation of adenine N1-oxide with however a very low efficiency [52]. Therefore H2O2 is likely involved in the formation of OH by Fenton type reactions with either ferrous ion and copper ions as the reducing agents, although in the latter case 1 O2 oxidation and one-electron oxidation of guanine have also been suggested to occur at least in model studies [53]. It may also be pointed out that OH reacts essentially at the site of its generation without any significant migration within the cells. Another relevant endogenous one-electron oxidant is inorganic carbonate radical anion (CO3) [54,55] that arises from the decomposition of nitrosoperoxycarbonate, the product of the reaction of peroxynitrite and carbonates [49,56]. It is also well-documented that ONOO is generated by radical coupling of nitric oxide (NO) and O 2 , two poorly reactive species, during inflammation processes in tissues [55].

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Exogenous radical oxidizing agents

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Ionizing radiation is one of the most commonly used exogenous oxidizing agents even if the molecular mechanism of action is complex involving several possible radical events in the formation of damage [57]. The indirect effect of X- and gamma-rays that predominates in aqueous solutions gives rise to OH through the radiolysis of water [17]. In addition, there is a contribution of ionization processes with the transient formation of organic radical cations through the direct interactions of high energy photons with target biomolecules such a DNA. A suitable way of generating DNA radical cations consists of exposing targets to high-intensity nanosecond UVC laser pulses leading to one-electron oxidation of purine and pyrimidine bases by a bi-photonic process [58,59]. One-electron oxidation of the guanine base is also possible upon incubation of cells with bromate, a renal carcinogen, subsequent

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to thiol-mediated reduction into Br2O [60]. A selective one-electron oxidation of the guanine base can also be achieved upon incubation of cells with 6-thioguanine and related analogs including azathioprine and 6-mercaptopurine that are incorporated into DNA before UVA excitation [61].

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Similarities between hydroxyl radical and one-electron oxidants-mediated DNA damage

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The two exogenous oxidizing systems which were applied to induce a significant increase in the yields of modified bases in cellular DNA with respect to steady-state levels include ionizing radiation through the predominant generation of OH and high intensity UV laser irradiation for one-electron oxidation of purine and pyrimidine bases.

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Single oxidized bases

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Several oxidized pyrimidine bases have been measured by HPLC–ESI-MS/MS as the corresponding 20 -deoxyribonucleosides after suitable extraction and enzymatic digestion of cellular DNA from either human monocytes [28,29] or Fischer F98 glioma cells [34] exposed to gamma rays of 137Cs sources. These include as degradation products of thymidine (dT) the four diastereomers of cis and trans 5,6-dihydroxy-5,6-dihydrothymidine (dTGly), 5(hydroxymethyl)-20 -deoxyuridine (HmdU) and 5-formyl-20 -deoxyuridine (5-FodU). More recently (5R⁄) and (5S⁄)-1-(2-deoxy-ß-Derythro-pentofuranosyl)-5-hydroxy-5-methylhydantoin (HydT), two diastereomeric pyrimidine ring rearrangement products have been detected as additional OH-mediated thymine oxidation products in cellular DNA [34]. In addition, 5-hydroxy-20 -deoxycytidine (5-OHdC), cis and trans diastereomers of 5,6-dihydroxy-5,6dihydro-20 -deoxyuridine (dUGly) and 3-(1-carbamoyl-4,5-dihydroxy-2-oxoimidazolidine (ImidC) together with the (5R⁄) and (5S⁄)-1-(2-deoxy-ß-D-erythro-pentofuranosyl)-5-hydroxyhydantoin (HydU) have been detected as cytosine decomposition products in the DNA of gamma-irradiated Fischer F98 glioma cells [34]. Several of these oxidized 20 -deoxyribonucleosides including dTGly, 5-HmdU, 5-FodU and 5-OHdC have also been shown to be generated in cellular DNA upon exposure to high intensity UVC nanosecond laser pulses [35]. The formation of dTGly may be rationalized in the initial steps by either OH addition across the 5,6ethylenic bond with a strong preference for the C5 position or hydration of the thymine radical cation 1 giving rise to the 6hydroxy-5,6-dihydrothymin-5-yl radical 2 [11,16,17]. Subsequently O2 is able to efficiently react with the carbon centered radicals thus generated at diffusion controlled rates of reaction giving rise to the formation of hydroperoxyl radicals and in turn to intermediate 5(6)-hydroxy 6(5)hydroperoxides [11,16]. The peroxyl radicals or hydroperoxides can decompose to give dTGly (Fig. 1). It may be concluded that there are strong similarities between the two types of oxidation reactions leading to dTGly particularly when OH addition takes place at the C6 position of thymine. This applies as well to the oxidative formation of 5-HmdU and 5-FodU that may be generated from either the deprotonation reaction of 1 or the OH-mediated H–atom abstraction from the methyl group of thymine, both resulting in the formation of 5-(uracilyl)methyl radical 3 [11]. Another relevant example of similarities in the  OH-mediated and one-electron oxidation of pyrimidine bases recently became apparent with the recent measurement of 5-OHdC in cellular DNA exposed to ionizing radiation and high intensity UVC irradiation [35]. The formation of 5-OHdC may be explained by the transient formation of cytosine radical cations upon one-electron oxidation followed by conversion of 6-hydroxy-5,

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O

O2

HN N dR

2

O

H

HN

N dR

-e

O

2 4

H2 N

HO

dThd

N dR

O HN 1 5

HydT

CH3

HN

HN

CH 3 OH

O

N dR

1

OH

dTGly

O

O

N

O

O

dR

OH CH3

H N

CH3 OH

HN

OH O

O

O

CH3

O

dR =

H 2N

-e

7

N N

dG

dR

OH

H 2N

CH 2

HN O

N

3

dR

O2

HN O

+e

O CH 2 OH

O

HN

N dR

5-FodU

5-HmdU

O

CHO

HN

N dR

O HN

OH O

+H2 O

8

N

HO

O

N

7

H2 N

O N

8-oxodG

• N

N

N

6

dR

OH H

-e O

H N N dR

N •+ N dR

HN H2 N

N

H N

CHO NH dR

FapydG



Fig. 1. One-electron oxidation and OH-mediated degradation of thymine.

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6-cytosyl radicals to 5,6-dihydroxy-5,6-dihydro-20 -deoxycytidine (dCGly), and finally dehydration of dCGly to 5-OHdC [12] (Fig. 2). Similarities have been also observed in the OH and one-electron oxidation degradation pathways of 20 -deoxyguanosine (dG) giving rise to 8-oxo-7,8-dihydro-20 -deoxyguanosine (8-oxodG) in cellular DNA [28,29,34,35]. Addition of OH to the C8 position of the purine moiety that is a relatively minor process with respect to recently reported OH-mediated hydrogen abstraction from the 2-amino group [62] leads to the transient formation of reducing 8-hydroxy-7,8-dihydroguanyl radical 6 that may also be generated by hydration of the purine radical cation 7, the one-electron oxidation product of dG (Fig. 3) [10,11]. In a subsequent step, one-electron oxidation of the latter radical that is likely to be mediated at least in part by O2 leads to 8-oxodGuo while competitive one-electron reduction yields the 20 -deoxyribonucleoside derivative of 2,6diamino-4-hydroxy-5-formamidopyrimidine (FapydG) [10,11].

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Tandem base lesion involving carbon centered pyrimidine radicals

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Another reaction of 5-(uracilyl)methyl radical 3 has been shown to give rise to interstrand cross-links through predominant covalent attachment to the C-8 of guanine preferentially on the 50 side [36,63–65]. The resulting G[8–5 m]T tandem lesion (Fig. 4) was subsequently found to be generated according to a predominant  OH-mediated pathway in the DNA of HeLa-S3 cells exposed to gamma rays [30]. The yield of G[8–5 m]T was much lower than that of either 5-FodU or 5-HmdU [66], which are also formed in

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NH2

N 4 dR

N O

OH

N

O

-e

O

N dR 5-OHdC

+H2O N dR

dC

NH 2

OH

H

N

-H2O NH 2

O2

N

OH O

N

5 dR

a competitive reaction of 5-(uracilyl)methyl radical 3 in the presence of oxygen. The presence of O2 is expected to efficiently scavenge the latter radical and therefore favor the formation of 5HmdU and 5-FodU through the transient formation of 5-hydroperoxymethyl radical at the expense of the tandem lesion. It may be added that G[8-5m]T has been shown to be endogenously formed in human brain tissues [67]. Interestingly, in the same study, G[85m]T was found to accumulate in the brain and liver of a rat model of Wilson’s disease that is characterized by an elevated production of ROS [67]. A second vicinal lesion whose formation involves the covalent addition of 6-hydroxy-5,6-dihydrocytosyl to 50 -guanine at C8 followed by dehydration of the resulting 6-hydroxy-5,6-dihydrocytosine residue has been characterized as a G[8-5]C intrastrand adduct and detected by HPLC-MS3 in the DNA of gamma irradiated HeLa-S3 cells [31]. One should be reminded that the formation of this G[8-5]C intrastrand adduct could be explained by either the addition of OH at C6 or the selective hydration of cytosine radical cation. The lack of detection of T[5m-8]G and C[5-8]G position isomers suggests that their formation is inefficient as reported in model studies [64,65]. This remark applies as well to the generation of tandem lesions involving adenine that exhibits a lower reactivity than guanine for the attachment reaction initiated by pyrimidine radicals. Other intrastrand crosslinks characterized in model studied derived from the initial generation of 5(cytosyl)methyl radical [68–70] and 5-hydroxy-5,6-dihydrothymin-6-yl radical [71,72] are awaiting detection in cellular DNA.

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Tandem base lesions involving peroxyl pyrimidine radicals

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Earlier evidence has been provided for the formation of tandem modifications consisting of 8-oxoGua and a formylamine residue, a well-characterized radical oxidation product of both thymine [11] and cytosine [12] when short oligonucleotides [73,74] and isolated DNA [75] are exposed to ionizing radiation in aerated aqueous solutions such that OH is the overwhelming reactive species. The formation of the tandem base modifications that is favored when the pyrimidine base is located at the 3’ position of the oligonucleotide strand was rationalized in terms of the addition reaction of 5(6)-hydroxy-6-hydroperoxy-5,6-dihydropyrimidyl radical [76,77]. This is accompanied by the transfer of an oxygen atom to the vicinal guanine base at C8 and the subsequent formation of two adjacent oxidized bases. This gives tandem lesions that are initiated by one initial radical hit. On the basis of [18O]-labeling

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NH 2

N

NH2

Fig. 3. One-electron oxidation and OH-mediated degradation of guanine.

H

O

N dR

H OH OH

dCGly

Fig. 2. One-electron oxidation and OH-mediated decomposition of cytosine.

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O

O

HN H 2N

CH2

N

N

O O

NH

N

N

O

O

HN H 2N

O

N

N

N

N

O

O O P O

NH 2 N

O

O

O

O O

O P O

OH

OH

G[8-5m]T

G[8-5]C

O

Fig. 4. Intrastrand cross-links involving 5-(uracilyl)methyl and 5-hydroxy-5,6-dihydrocytosyl radicals.

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experiments, it has been suggested been suggested that about half of 8-oxoGua lesions generated in duplex DNA by OH are partners in tandem base modifications whose formation is triggered by pyrimidine peroxyl radicals [78]. Other examples implicating peroxyl radicals in the formation of tandem lesions with thymine [79– 81] and the 2-deoxribose moiety [82,83] have not been detected so far in cellular DNA either upon exposure to OH or one-electron oxidants. A comprehensive study of the site-specific reactivity of 5-hydroxy-6-hydroperoxy-5,6-dihydrothymyl radical in DNA that leads to the formation of clustered damage has recently become available [84]. Efforts should be made in forthcoming studies to search for these biologically relevant vicinal modifications when the sensitivity of MS based methods is further improved.

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Selective hydroxyl radical-mediated DNA damage

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Oxidatively generated damage to the 2-deoxyribose backbone of DNA is mostly accounted for by initial H-atom abstraction reactions mediated by OH under various oxidative stress conditions [17,18,85,86]. However, several oxidants including 6-hydroperoxy-5,6-dihyropyrimidinyl radicals [77,81,82,87], 5-uracilyl radical generated by one-electron reduction of 5-halogenouracil [88,89] and activated antibiotics such as bleomycin [90,91] have the ability to oxidize, usually in a more selective way, the DNA sugar units. In addition, electron loss followed by deprotonation of the radical cation constitutes the steps leading to the formation of carbon centered 2-deoxyribosyl radicals through the direct effects of ionizing radiation [92]. It should be noted that most of biologically relevant one-electron oxidants including CO 3 radical, BrO2 and type I photosensitizers are able to oxidize guanine but not 2deoxyribose.

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Single strand breaks Formation of DNA strand breaks in cells is mostly explained by OH-mediated hydrogen abstraction from 2-deoxyribose at C3, C4 and C5 [17,18]. In addition 2-deoxyribonolactone (dL) and the C4 oxidized abasic site (C4-AP) have been shown to be generated through the transient formation of peroxyl radicals that arise from the addition of O2 to C1 and C4 centered radicals of the 2-deoxyribose backbone of DNA, respectively [18,87]. Several methods including GC–MS and HPLC have been developed for measuring dL and C4-AP residues in isolated DNA [93,94]. However no evidence has been provided so far toward the unambiguous formation of any of the two oxidized abasic sites in cellular DNA. In contrast more complex lesions including interstrand and intrastrand crosslinks that arise from further reactions of C4-AP and C5 radicals respectively, with nucleobases have been detected in the DNA of human cells as further discussed below. 

Interstrand cross-links

315

Abasic sites that result from the hydrolytic cleavage of the N-glycosidic bond are implicated in the formation of ICLs through the reaction of electrophilic aldehydes with either the 2-amino group of guanine [95] or the 6-amino group of adenine [96] on the opposite strand. However, no evidence has been provided so far for the occurrence of these ICLs in cells as well as for cross-links arising from the reaction of dL with several DNA repair glycosylases [97]. Interestingly C4-AP through a complex sequence of reactions has been shown to be implicated in the formation of ICLs when human monocyte cells were exposed to either radiation-induced OH or bleomycin mediated-specific H-atom abstraction from C4 of the 2-deoxyribose moiety [98]. This was supported by the HPLC-MS/ MS detection of the four diastereomers of 6-(2-deoxy-ß-D-erythropentofuranosyl)-2-hydroxy-3(3-hydroxy-2-oxopropyl)-2,6-dihydroimidazo[1,2-c]-pyrimidin-5(3H)-one 9 after suitable enzymatic hydrolysis of DNA [98,99]. The mechanism of formation of 9 may be rationalized in terms of the catalytic release of a highly reactive unsaturated keto-aldehyde 8 from C4-AP, which in a subsequent step undergoes efficient cycloaddition across the 5,6-ethylenic bond of cytosine on the opposite strand (Fig. 5). Further information on the selectivity and efficiency of ICLs was gained from subsequent detailed studies that are involved the site-specific generation of C4-AP using suitable photolabile precursors of sugar C4 radical [100,101]. It was found that adenine and to a lesser extent cytosine when located on the complementary strand opposite to CA-AP are prone to induce the b-elimination reaction in a sequence dependence manner giving rise to 8 with concomitant cleavage of the 30 phosphodiester bond of the C40 abasic site. It may be added that cycloaddition of 8 with adenine on the opposite strand has been shown to occur in DNA duplexes [97]. The search for the possible formation of the adenine cycloadducts in cellular DNA is awaiting further experiments. The radiation-induced formation of ICL in cellular DNA is a relatively minor oxidative pathway being about 2 orders of magnitude lower than the formation of 8-oxodG. However this oxidatively generated ICL whose severity is increased by the presence of a vicinal strand break is likely to represent a challenge for the cellular repair machinery as inferred from the observed very small removal of 9 from damaged DNA.

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Purine 50 ,8-cyclo-20 -deoxyribonucleosides

356

Earlier evidence for the OH-mediated formation of 50 ,8-cyclo20 -deoxyribonucleosides through the transient generation of 2deoxyribos-5-yl radical was gained from model studies that have involved purine 20 -deoxyribonucleosides and nucleotides [for a comprehensive review, see 38]. A regain of interest for this class of intrastrand cross-links appeared at the beginning of year 2000’

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OH O

O

HO

cytosine catalysis

O

OH

N

opposite C

N

O

O

O

N

O

O

O O

O

8

9

C4-AP O Fig. 5. Interstrand cross-link involving the 2-deoxyribos-4-yl radical.

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due to their structural features. The presence of a 50 -8 bond allow them to be substrates for the nucleotide excision repair pathway [102–104] but not substrates for the common base excision repair enzymes operating for most oxidatively generated base damage [102]. Therefore, emphasis was placed on the measurement of the 50 R and 50 S diastereomers of 50 ,8-cyclo-20 -deoxyadenosine (cyclodA) and 50 ,8-cyclo-20 -deoxyguanosine (cyclodG) that has led to inconsistent data reporting overestimated values of about 2–3 orders of magnitude. This was due to the use of inaccurate GC–MS and HPLC-MS methods [105,106] that are prone to several major drawbacks including artifactual oxidation and lack of detection specificity [32]. The recently developed HPLC-MS/MS method has led to much lower values for (50 R)-cyclodA, the only purine 50 ,8-cyclo-20 -deoxyribonucleoside detected in cellular DNA upon exposure to 2 kGy of gamma rays [32]. The yield of (50 R)-cyclodA that represents about 1% of that of 8-oxoGua is consistent with the low efficiency for OH to generate C5-yl radical on adenine nucleotide (less than 3%) and the efficient reaction of O2 to scavenge the latter radical as shown in model studies [32]. The four diastereomers of cyclodA and cyclodG have been detected in the DNA of brain and liver tissues of Long-Evans Cinnamon rats together with thymidine oxidation products including 5-FodU and 5-HmdU using a HPLC-MS3 method [23]. As expected the steady-state levels of both cyclodA and cyclodG were lower by at least two orders of magnitude with respect to those of thymidine oxidation products, which surprisingly are elevated in the DNA of cells, receiving a dose of about 500 Gy of ionizing radiation. The mechanism of generation of the purine 20 -deoxyribonucleosides is rationalized in terms of initial OH-mediated formation of 2deoxyribos-5-yl radicals, followed by intramolecular cyclization involving the C8 of the purine bases with rate constants measured k = (1.6 ± 0.2)  105 s1 and (6.9 ± 0.8)  105 s1 for adenine and guanine, respectively [107,108]. Subsequently the resulting transient radicals are oxidized by oxygen to yield cyclodA and cyclodG

(Fig. 6) with a predominance of 50 R diastereomer with respect to the 50 S derivative by factors varying between 3 and 4, respectively as recently confirmed in a recent model study in which a Fenton reagent was used as the oxidant [109].

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Selective one-electron oxidant-mediated DNA damage

401

As already discussed, most biologically relevant one-electron oxidation reactions to cellular DNA involve predominantly the guanine base, which exhibits the lowest ionization potential among DNA components. Furthermore, when stronger oxidants are applied, such as high intensity UV laser pulses and ionization radiation through its direct effect, the initially generated radical cations are redistributed through charge transfer along the DNA duplex with preferential trapping at guanine sites acting as sinks of hole migration. Therefore, the one-electron oxidation of purine and pyrimidine bases gives rise in large part to final damage located at guanine sites [24,35,110]. One of the main chemical features of guanine radical cations 7 is their ability to undergo reactions with nucleophiles including nucleobases and lysine residue of proteins.

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Guanine–Thymine intrastrand cross-links

416

A relevant example of nucleophilic addition reaction of thymine to guanine radical cations in DNA was recently reported using HPLC–ESI-MS/MS detection and HeLa cells exposed to high intensity nanosecond UVC laser pulses [35]. The yield of dG⁄-dT⁄ intrastrand cross-link involving guanine and thymine bases separated by one extra base was dependent on the intensity of the laser pulses and it was generated with a lower yield compared to 8oxodG (about 800-fold) as well as either 5-FodU or 5-HmdU (60fold). The mechanism of dG⁄-dT⁄ adduct formation involves initial

417

NH 2 N HO H HO H

B

N

NH 2 N

N H

N

HO

O

OH

(5'R)-cyclodA

N

N N

O

(5'S)-cyclodA

OH

O O N

OH

B= adenine, guanine

HO H

N

O

N

NH2

H HO

O

OH

N

NH

(5'R)-cyclodG

N

NH N

NH2

O

OH

(5'S)-cyclodG

Fig. 6. Formation of purine 50 ,8-cyclo-20 -deoxyribonucleosides.

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DNA-DNA crosslink dC

O N

HN H 2N

N dR

N

7

H2 N

dT

H N

H2 N N

N

O

O O

O

H 2N

O

O P O OH

N N

N dR

N

HO

N

O

O

N

KKK

HN

O

N

N

HO

H N

O

N H

O

N N

OH

O

HO

O

N

O

NH

dG*pdT*

OH

dG*-dT*

OH

Fig. 7. Formation of intrastrand and DNA protein cross-links involving the guanine radical cation.

435

generation of 7 followed by the creation of a covalent bond between C8 atom of guanine and N3 atom of thymine (Fig. 7). This was inferred from detailed model studies performed either on single-stranded oligonucleotides [37] or DNA duplexes [111] with the carbonate radical anion used as the guanine one-electron oxidant. The nucleophilic addition of thymine to 7 was found to be more efficient when the two intervening bases were separated by one cytosine base in a 50 -GCT-30 sequence context with respect to a vicinal location in 50 -GT-30 sequences giving rise to dG⁄pT⁄ after enzymatic digestion of DNA.

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Interstrand cross-links

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As reported above, OH has been shown to induce the formation of ICLs in cellular DNA following the initial generation of C4AP. A second example of oxidative formation of ICLs in which guanine radical cation 7 are key reactive intermediate has recently become available [13]. Thus, high intensity UVC nanosecond laser irradiation of DNA duplexes was shown to generate ICLs as inferred from polyacrylamide gel electrophoresis analysis [112]. The formation of ICLs was also observed upon UVA irradiation of either 6-thioguanine (TG) containing double-stranded DNA fragments or cells that were pre-incubated with TG in order to allow its incorporation into DNA [113,114]. A likely mechanism for the one-electron mediated formation of these ICLs that is still under investigation would involve the nucleophilic addition of either guanine or cytosine on the opposite strand to initially generated 7. This suggests the occurrence of a similar reaction pathway for explaining at least partly the radiationinduced formation of ICLs in cellular DNA through ionization of guanine bases.

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DNA–protein cross-links

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Ionizing radiation is able to induce the formation of DNA–protein cross-links, one of the main classes of deleterious oxidatively generated DNA damage, with an efficiency that is higher than that of reactions leading to ICLs and double strand breaks [115]. Several mechanisms including the radical–radical coupling of radiation-induced nucleobase and amino acid radicals have been proposed to explain the formation of DPCs [116,117]. However, the presence of a relatively high concentration of O2 in normoxic cells and tumor tissues with respect to anoxic conditions either did not or only partly inhibited the generation of DPCs upon exposure to gamma rays [118] and 12C6 (290 MeV/u) ions [119], respectively. This strongly suggests that at least part of the radiation-induced DPCs are not formed by radical–radical coupling

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because O2 would at least partly intercept these radicals. As an alternative mechanism that has received strong support from several model studies [120–124], it was proposed that free nucleophilic groups of bound proteins are able to covalent attach to the C8 of guanine leading to the formation of stable DPCs. Lysine, arginine and serine but not tyrosine were identified as reactive amino acid residues involved in the formation of DPCs. A comprehensive mechanism was provided from a detailed study of riboflavin photosensitization of TGT in which the guanine moiety was bound to trilysine (KKK) peptide, a model mimicking DNA– protein interactions [120]. In this reaction, one-electron oxidation of the guanine base was followed by efficient addition of the free e-amino group of the amino acid of KKK to the C8 position of 7 in a highly competitive way with the nucleophilic addition of water leading to 8-oxodG [125,126]. In a subsequent step, oxidation of the aminyl radical thus formed, gives rise to a stable DPC (Fig. 7) that was unambiguously characterized by extensive NMR and mass spectrometry analyses [120]. Interestingly, DPCs have been detected in UVA-irradiated cells that were incubated with 6-mercaptopurine, azathioprine or TG and subsequently incorporated in DNA as TG once metabolized. Unambiguous characterization of the latter DPCs is pending.

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Conclusions

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A large body of information in now available on the main oxidative degradation pathways of nucleic acid components mediated by OH and one-electron oxidants with emphasis on both purine and pyrimidine nucleobases. This is mostly inferred from the characterization of almost one hundred final oxidation products and detailed mechanistic studies that in some cases have received further confirmation from theoretical calculations. More recently significant progress has been made in the measurement of single and more complex oxidatively generated damage to cellular DNA thanks to the advent of accurate and quantitative methods including HPLC coupled with MS/MS and MS3 detection technique. Interestingly the formation of oxidation products in cellular DNA under well given conditions of oxidative stress has been mechanistically rationalized using information from model studies. This has allowed to identify similarities and differences in the oxidation reactions to cellular DNA triggered by OH and one-electron oxidants. Therefore the availability of dedicated biomarkers of specific degradation pathway could be of value for gaining mechanistic insights into radical reactions in cells. It remains however to improve the sensitivity of the detection methods and to extend the measurement to a larger number of oxidatively generated damage to DNA.

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References

514 515 516 517 518 519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556 557 558 559 560 561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589 590 591 592 593 594 595 596 597

Q4

[1] C. Canugovi, M. Misiak, L.K. Ferrarelli, D.L. Croteau, V.A. Bohr, DNA Repair (Amst) 12 (2013) 578–587. [2] R.S. Sohal, W.C. Orr, Free Radic. Biol. Med. 52 (2012) 539–555. [3] D. Ziech, R. Franco, A. Pappa, M.I. Panayiotidis, Mutat. Res. 711 (2011) 167– 173. [4] S. Toyokuni, IUBMB Life 60 (2008) 441–447. [5] S. Maynard, S.H. Schurman, C. Harboe, N.C. de Souza-Pinto, V.A. Bohr, Carcinogenesis 30 (2009) 2–10. [6] A. Borghini, T. Cervelli, A. Galli, M.G. Andreassi, Atherosclerosis 230 (2013) 202–209. [7] J. Wang, S. Xiong, C. Xie, W.R. Markesbery, M.A. Lovell, J. Neurochem. 93 (2005) 953–962. [8] Z. Sheng, S. Oka, D. Tsuchimoto, N. Abolhassani, H. Nomaru, K. Sakumi, H. Yamada, Y. Nakabeppu, J. Clin. Invest. 122 (2012) 4344–4361. [9] G. Pratviel, B. Meunier, Chemistry 12 (2006) 6018–6030. [10] J. Cadet, T. Douki, J.-L. Ravanat, Acc. Chem. Res. 41 (2008) 1075–1083. [11] J. Cadet, T. Douki, J.-L. Ravanat, Free Radic. Biol. Med. 40 (2010) 9–21. [12] J.R. Wagner, J. Cadet, Acc. Chem. Res. 43 (2010) 564–571. [13] J. Cadet, J.-L. Ravanat, M. TavernaPorro, H. Menoni, D. Angelov, Cancer Lett. 327 (2012) 5–15. [14] L. Cui, W. Ye, E.G. Prestwich, J.S. Wishnok, K. Taghizadeh, P.C. Dedon, S.R. Tannenbaum, Chem. Res. Toxicol. 26 (2013) 195–202. [15] A.M. Fleming, C.J. Burrows, Chem. Res. Toxicol. 26 (2013) 593–607. [16] J. Cadet, J.R. Wagner, Cold Spring Harb. Perspect. Biol. 5 (2013) a012559. [17] C. von Sonntag, Free-Radical-Induced DNA Damage and its Repair. A Chemical Perspective, Springer-Verlag, Berlin, Heidelberg, New-York, 2006. [18] P.C. Dedon, Chem. Res. Toxicol. 21 (2008) 206–219. [19] M. Pitié, G. Pratviel, Chem. Rev. 110 (2010) 1018–1059. [20] A. Azqueta, A.R. Collins, Arch. Toxicol. 87 (2013) 949–968. [21] A.R. Collins, Biochim. Biophys. Acta 1840 (2013) 794–800. [22] J. Cadet, T. Douki, J.-L. Ravanat, Mutat. Res. 711 (2011) 3–12. [23] J. Wang, B. Yuan, C. Guerrero, R. Bahde, S. Gupta, Y. Wang, Anal. Chem. 83 (2011) 2201–2209. [24] J. Cadet, T. Douki, J.-L. Ravanat, J.R. Wagner, Bioanal. Rev. 4 (2012) 55–74. [25] T. Bessho, K. Tano, S. Nishimura, H. Kasai, Carcinogenesis 14 (1993) 1069– 1071. [26] A.R. Collins, Adv. Food Nutr. Res. 68 (2013) 283–290. [27] J.-L. Ravanat, P. Di Mascio, G.R. Martinez, M.H. Medeiros, J. Cadet, J. Biol. Chem. 275 (2000) 40601–40604. [28] J.-P. Pouget, S. Frelon, J.-L. Ravanat, I. Testard, F. Odin, J. Cadet, Radiat. Res. 157 (2002) 589–595. [29] T. Douki, J.-L. Ravanat, J.-P. Pouget, I. Testard, J. Cadet, Int. J. Radiat. Biol. 82 (2006) 119–127. [30] Y. Jiang, H. Hong, H. Cao, Y. Wang, Biochemistry 46 (2007) 12757–12763. [31] H. Hong, H. Cao, Y. Wang, Nucleic Acids Res. 35 (2007) 7118–7127. [32] N. Belmadoui, F. Boussicault, M. Guerra, J.-L. Ravanat, C. Chatgilialoglu, J. Cadet, Org. Biomol. Chem. 8 (2010) 3211–3219. [33] J. Wang, C.L. Clauson, P.D. Robbins, L.J. Niedernhofer, Y. Wang, Aging Cell 11 (2012) 714–716. [34] F. Samson-Thibault, G.S. Madugundu, S. Gao, J. Cadet, J.R. Wagner, Chem. Res. Toxicol. 25 (2012) 1902–1911. [35] G.S. Madugundu, J.R. Wagner, J. Cadet, K. Kropachev, B.H. Yun, N.E. Geacintov, V. Shafirovich, Chem. Res. Toxicol. 26 (2013) 1031–1033. [36] Y. Wang, Chem. Res. Toxicol. 21 (2008) 276–281. [37] C. Crean, Y. Uvaydov, N.E. Geacintov, V. Shafirovich, Nucleic Acids Res. 36 (2008) 742–755. [38] C. Chatgilialoglu, C. Ferreri, M.A. Terzidis, Chem. Soc. Rev. 40 (2011) 1368– 1382. [39] J. Cadet, T. Douki, E. Sage, Mutat. Res. 571 (2005) 3–17. [40] B. Epe, Photochem. Photobiol. Sci. 11 (2011) 98–106. [41] G. Bauer, Anticancer Res. 33 (2013) 3589–3602. [42] J. Cadet, T. Douki, J.-L. Ravanat, P. Di Mascio, Photochem. Photobiol. Sci. 8 (2009) 903–911. [43] L.F. Agnez-Lima, A.H.S. Oliveira, A.R.S. Timoteo, K.M. Lima-Bessa, G.R. Martinez, M.H.G. Medeiros, P. Di Mascio, R.S. Galhardo, C.F.M. Menck, Mutat. Res. 751 (2012) 15–28. [44] H. Parker, C.C. Winterbourn, Front. Immunol. 3 (2013) 1–6. [45] T. Nakano, M. Masuda, T. Suzuki, H. Ohshima, Biosci. Biotechnol. Biochem. 76 (2012) 2208–2213. [46] C. Badouard, M. Masuda, H. Nishino, J. Cadet, A. Favier, J.-L. Ravanat, J. Chromatogr. B 827 (2005) 26–31. [47] H. Nikjoo, P. Girard, Int. J. Radiat. Biol. 88 (2012) 87–97. [48] A.G. Georgakilas, P. O’Neill, R.D. Stewart, Radiat. Res. 180 (2013) 100–109. [49] B. Kalyanaraman, Redox Biol. 1 (2013) 244–257. [50] J. Cadet, S. Loft, R. Olinski, M.D. Evans, K. Bialkowski, J.R. Wagner, P.C. Dedon, P. Moeller, M.M. Greenberg, M.S. Cooke, Free Radic. Res. 48 (2012) 367–381. [51] R. Misiaszek, C. Crean, A. Joffe, N.E. Geacintov, V. Shafirovich, J. Biol. Chem. 279 (2004) 32106–32115. [52] J. Cadet, M. Berger, T. Douki, J.-L. Ravanat, Rev. Physiol. Biochem. Pharmacol. 131 (1997) 1–87. [53] S. Frelon, T. Douki, A. Favier, J. Cadet, Chem. Res. Toxicol. 16 (2003) 191–197. [54] V. Shafirovich, A. Dourandin, W. Huang, N.E. Geacintov, J. Biol. Chem. 276 (2001) 24621–24626.

7

[55] C.C. Winterbourn, Nat. Chem. Biol. 4 (2008) 278–286. [56] D.B. Medinas, G. Cerchiaro, D.F. Trindade, O. Augusto, IUBMB Life 59 (2007) 255–262. [57] P. O’Neill, P. Wardman, Int. J. Radiat. Biol. 85 (2009) 9–25. [58] T. Douki, D. Angelov, J. Cadet, J. Am. Chem. Soc. 123 (2001) 11360–11366. [59] T. Douki, J.-L. Ravanat, D. Angelov, J.R. Wagner, J. Cadet, Top. Curr. Chem. 236 (2004) 1–25. [60] M. Murata, Y. Bansho, S. Inoue, K. Ito, S. Ohnishi, S. Kawanishi, Chem. Res. Toxicol. 14 (2001) 678–685. [61] R. Brem, P. Karran, Photochem. Photobiol. 88 (2012) 5–13. [62] C. Chatgilialoglu, M. D’Angelantonio, M. Guerra, P. Kaloudis, Q.G. Mulazzani, Angew. Chem. Int. Ed. 48 (2009) 2214–2217. [63] H.C. Box, E.E. Budzinski, J.B. Dawidzik, J.C. Wallace, M.S. Evans, J.S. Gobey, Radiat. Res. 145 (1996) 641–643. [64] S. Bellon, J.-L. Ravanat, D. Gasparutto, J. Cadet, Chem. Res. Toxicol. 15 (2002) 598–606. [65] V. Labet, C. Morell, A. Grand, J. Cadet, P. Cimino, V. Barone, Org. Biomol. Chem. 6 (2008) 3300–3305. [66] H. Hong, Y. Wang, Anal. Chem. 79 (2007) 322–326. [67] J. Wang, H. Cao, C. You, B. Yuan, R. Bahde, S. Gupta, C. Nishigori, L.J. Niedernhofer, P.J. Brooks, Y. Wang, Nucleic Acids Res. 40 (2012) 7368–7374. [68] Q. Zhang, Y. Wang, J. Am. Chem. Soc. 125 (2003) 12595–12802. [69] Q. Zhang, Y. Wang, Nucleic Acids Res. 33 (2005) 1593–1603. [70] H. Cao, Y. Wang, Nucleic Acids Res. 35 (2007) 4833–4844. [71] Q. Zhang, Y. Wang, J. Am. Chem. Soc. 126 (2004) 13287–13297. [72] Q. Zhang, Y. Wang, Chem. Res. Toxicol. 18 (2005) 1897–1906. [73] H.C. Box, H.G. Freund, E.E. Budzinski, J.C. Wallace, A.E. Maccubbin, Radiat. Res. 141 (1995) 91–94. [74] E.E. Budzinski, J.D. Dawidzik, J.C. Wallace, H.G. Freund, H.C. Box, Radiat. Res. 142 (1995) 107–109. [75] A.G. Bourdat, T. Douki, S. Frelon, D. Gasparutto, J. Cadet, J. Am. Chem. Soc. 122 (2000) 4549–4556. [76] T. Douki, J. Rivière, J. Cadet, Chem. Res. Toxicol. 15 (2002) 445–454. [77] C. Dupont, C. Patel, J.-L. Ravanat, E. Dumont, Org. Biomol. Chem. 11 (2013) 3038–3045. [78] F. Bergeron, F. Auvré, J.P. Radicella, J.-L. Ravanat, Proc. Natl. Acad. Sci. U.S.A. 107 (2010) 5528–5533. [79] S. Hong, K.N. Carter, K. Sato, M.M. Greenberg, J. Am. Chem. Soc. 129 (2007) 4089–4098. [80] A. Ghosh, A. Joy, G.B. Schuster, T. Douki, J. Cadet, Org. Biomol. Chem. 6 (2008) 916–928. [81] I.S. Hong, K.N. Carter, K. Sato, M.M. Greenberg, J. Am. Chem. Soc. 129 (2007) 4089–4098. [82] H. Huang, S. Imoto, M.M. Greenberg, Biochemistry 48 (2009) 7833–7841. [83] M.L. TavernaPorro, M.M. Greenberg, J. Am. Chem. Soc. 135 (2013) 16268– 16371. [84] J.M.N. San Pedro, M.M. Greenberg, J. Am. Chem. Soc. 136 (2014) 3928–3936. [85] B. Balasubramanian, W.K. Pogozelski, T.D. Tullius, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 9738–9743. [86] W.K. Pogozelski, T.D. Tullius, Chem. Rev. 98 (1998) 1089–1108. [87] C.S. Price, Y. Razskzovskiy, W.A. Bernhard, Radiat. Res. 174 (2010) 645–649. [88] T. Watanabe, T. Bando, Y. Xu, R. Tashiro, H. Sugiyama, J. Am. Chem. Soc. 127 (2005) 44–45. [89] Y. Xu, R. Tashiro, H. Sugiyama, Nat. Protoc. 2 (2007) 78–87. [90] S.M. Hecht, J. Nat. Prod. 63 (2000) 156–168. [91] J. Chen, J. Stubbe, Nat. Rev. Cancer 5 (2005) 102–112. [92] S. Purkayastha, J.R. Milligan, W.A. Bernhard, Radiat. Res. 168 (2007) 357– 366. [93] W. Chan, B. Chen, L. Wang, K. Taghizadeh, M.S. Demott, P.C. Dedon, J. Am. Chem. Soc. 132 (2010) 6145–6153. [94] M. Roginskaya, R. Mosheni, T.J. Moore, W.A. Bernhard, Y. Razskazovskiy, Radiat. Res. 181 (2014) 131–137. [95] S. Dutta, G. Chowdhury, K.S. Gates, J. Am. Chem. Soc. 129 (2007) 1852–1853. [96] N.E. Price, K.M. Johnson, J. Wang, M.I. Fekry, Y. Wang, H.S. Gates, J. Am. Chem. Soc. (2014) (Epub ahead of print). [97] M.M. Greenberg, Acc. Chem. Res. 47 (2014) 646–655. [98] P. Regulus, B. Duroux, P.A. Bayle, A. Favier, J. Cadet, J.-L. Ravanat, Proc. Natl. Acad. Sci. U.S.A. 104 (2006) 14032–14037. [99] P. Regulus, S. Spessotto, M. Gateau, J. Cadet, A. Favier, J.-L. Ravanat, Rapid Commun. Mass Spectrom. 18 (2004) 2223–2228. [100] J.T. Sczepanski, A.C. Jacobs, A. Majumdar, M.M. Greenberg, J. Am. Chem. Soc. 131 (2009) 11132–11139. [101] J.T. Sczepanski, C.N. Hiemstra, M.M. Greenberg, Biorg. Med. Chem. 19 (2011) 5788–5793. [102] I. Kuraoka, C. Bender, A. Romieu, J. Cadet, R.D. Wood, T. Lindhal, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 3832–3837. [103] P.J. Brooks, D.S. Wise, D.A. Berry, J.V. Kosmoski, M.J. Smerdon, R.L. Somers, H. Mackie, A.Y. Spoonde, E.J. Ackerman, K. Coleman, R.E. Tarone, J.H. Robbins, J. Biol. Chem. 275 (2000) 22355–22362. [104] K. Kropachev, S. Ding, M.A. Terzidis, A. Masi, Z. Liu, Y. Cai, M. Kolbanovskiy, C. Chatgilialoglu, S. Broyde, N.E. Geacintov, V. Shafirovich, Nucleic Acids Res. (2014) (Epub ahead of print). [105] M. D’Errico, E. Parlanti, M. Teson, B.M. Bernades de Jesus, P. Degan, A. Calcagnile, P. Jaruga, M. Bjøras, M. Crescenzi, A.M. Pedrini, J.M. Egly, G. Zambruno, M. Stefanini, M. Dizdaroglu, E. Dogliotti, EMBO J. 25 (2006) 4305– 4315.

Please cite this article in press as: J. Cadet, J.R. Wagner, Arch. Biochem. Biophys. (2014), http://dx.doi.org/10.1016/j.abb.2014.05.001

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YABBI 6676

No. of Pages 8, Model 5G

14 May 2014 8 684 685 686 687 688 689 690 691 692 693 694 695 696 697 698 699 700 701 702 703

J. Cadet, J.R. Wagner / Archives of Biochemistry and Biophysics xxx (2014) xxx–xxx

[106] M. D’Errico, E. Parlanti, M. Teson, P. Degan, T. Lemma, A. Calcagnile, I. Iavarone, P. Jaruga, M. Ropolo, A.M. Pedrini, D. Orioli, G. Frosina, G. Zambruno, M. Dizdaroglu, M. Stefanini, E. Dogliotti, Oncogene 26 (2007) 4336–4343. [107] F. Boussicault, P. Kaloudis, C. Caminal, Q.G. Mulazzani, C. Chatgilialoglu, J. Am. Chem. Soc. 130 (2008) 8377–8385. [108] C. Chatgilialoglu, M. D’Angelantonio, G. Kciuk, K. Bobrowski, Chem. Res. Toxicol. 24 (2011) 2200–2206. [109] C.R. Guerrero, J. Wang, Y. Wang, Chem. Res. Toxicol. 26 (2013) 1361–1368. [110] J. Cadet, J.R. Wagner, V. Shafirovich, N.E. Geacintov, Int. J. Radiat. Biol. (2014) (Epub ahead of print). [111] B.H. Yun, N.E. Geacintov, V. Shafirovich, Chem. Res. Toxicol. 24 (2011) 1144– 1152. [112] S. Cuesta-López, H. Menoni, D. Angelov, M. Peyrard, Nucleic Acids Res. 39 (2011) 5276–5283. [113] N.R. Attard, P. Karran, Photochem. Photobiol. Sci. 11 (2011) 62–68. [114] R. Brem, I. Daehn, P. Karran, DNA Repair (Amst) 10 (2011) 869–876. [115] S. Barker, M. Weinfeld, D. Murray, Mutat. Res. 580 (2005) 111–135. [116] R. Olinski, Z.Z. Nackerdien, M. Dizdaroglu, Arch. Biochem. Biophys. 297 (1992) 139–143. [117] M. Dizdaroglu, P. Jaruga, Free Radic. Res. 46 (2012) 387–419.

[118] S. Barker, M. Weinfeld, J. Zheng, L. Li, D. Murray, J. Biol. Chem. 280 (2005) 33826–33838. [119] M.I. Shoulkamy, T. Nakano, M. Ohshima, R. Hirayama, A. Uzawa, Y. Furusawa, H. Ide, Nucleic Acids Res. 40 (2012) e143. [120] S. Perrier, J. Hau, D. Gasparutto, J. Cadet, A. Favier, J.-L. Ravanat, J. Am. Chem. Soc. 128 (2006) 5703–5710. [121] K.L. Nguyen, M. Steryo, K. Kurbanyan, K.M. Nowitzki, S.M. Butterfield, S.R. Ward, E.D.A. Stemp, J. Am. Chem. Soc. 122 (2000) 3585–3594. [122] A.L. Madison, Z.A. Perez, P. To, T. Maisonet, E.V. Rios, Y. Trejo, C. OchoaPaniagua, A. Reno, E.D.A. Stemp, Biochemistry 51 (2012) 362–369. [123] K. Kurbanyan, K.L. Nguyen, P. To, E.V. Rivas, A.M.K. Lueras, C. Kosinski, M. Steryo, A. Gonzalez, D.A. Mah, E.D.A. Stemp, Biochemistry 42 (2003) 10269– 10281. [124] D. Angelov, M. Charra, C.W. Müller, J. Cadet, S. Dimitrov, Photochem. Photobiol. 77 (2003) 592–596. [125] H. Kasai, Z. Yamaizumi, M. Berger, J. Cadet, J. Am. Chem. Soc. 114 (1992) 9692–9694. [126] Y. Rokhlenko, J. Cadet, N.E. Geacintov, V. Shafirovich, J. Am. Chem. Soc. (2014) (Epub ahead of print).

Please cite this article in press as: J. Cadet, J.R. Wagner, Arch. Biochem. Biophys. (2014), http://dx.doi.org/10.1016/j.abb.2014.05.001

704 705 706 707 708 709 710 711 712 713 714 715 716 717 718 719 720 721 722 723