Mutagenicity of Ochratoxin A: Role for a Carbon ... - ACS Publications

Nov 9, 2016 - Incorporation of model C-linked C8-aryl−dG adducts into the G3 site of the NarI sequence demonstrates a tendency to induce...
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Mutagenicity of Ochratoxin A: Role for a Carbon-Linked C8− Deoxyguanosine Adduct? Richard A. Manderville*,† and Stacey D. Wetmore*,§ †

Departments of Chemistry and Toxicology, University of Guelph, Guelph, Ontario, Canada N1G 2W1 Department of Chemistry & Biochemistry, University of Lethbridge, Lethbridge, Alberta, Canada T1K 3M4

§

ABSTRACT: Ochratoxin A (OTA) is a fungal toxin that is considered to be a potent kidney carcinogen in rodent models. The toxin produces double strand breaks and has a propensity for deletions, single-base substitutions, and insertions. The toxin reacts covalently with DNA to afford a C8−2′-deoxyguanosine carbon-linked adduct (OT−dG) as the major lesion in animal tissues. Incorporation of model C-linked C8-aryl−dG adducts into the G3 site of the NarI sequence demonstrates a tendency to induce base substitutions and deletion mutations in primer extension assays using model polymerases. The degree of misincorporation induced by the C-linked C8−dG adducts correlates with an ability to adopt the promutagenic syn conformation within the NarI duplex as predicted by molecular dynamics (MD) simulations. MD simulations of the OT−dG adduct within the NarI duplex predict an even greater degree of conformational flexibility, suggesting enhanced in vitro mutagenicity compared to the simpler model C-linked C8−dG adducts. Together these findings support the role of OT−dG in promoting OTA-mediated mutagenicity and carcinogenicity in animal studies. KEYWORDS: ochratoxin A (OTA), DNA damage, DNA adduct, DNA structure, mutagenicity, DNA replication, DNA polymerase, translesion synthesis

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been demonstrated in the kidney target site of rats16,17 and mice.18,19 In rats, OTA facilitates DNA double strand breaks (DSBs) to produce large deletion mutations,16 whereas singlebase pair deletions in repetitive GC sequences, base insertions, and base substitutions are predominant in mice.19 Although the reasons for differences in the mutation spectra between mice and rats treated with OTA remain unknown, it is speculated that DSBs mediated by OTA are the critical trigger for gene mutations.17 In general, DSBs can be induced by ionizing radiation, by reactive oxygen species (ROS), or during DNA replication when a polymerase encounters a single-strand lesion at the replication fork.20,21 However, analysis of damaged DNA from the kidney of rats exposed to OTA displayed no evidence for 8-oxo-2′-deoxyguanosine (8-oxo-dG) formation,16 the primary lesion produced by ROS.22 Gene expression analysis also showed a lack of evidence for changes in the genes related to oxidative stress.17 Thus, the overall conclusion from the rodent studies was that OTA induces DSBs and mutagenicity at the kidney target site by a mechanism that precludes oxidative stress16,17 and may be a consequence of the polymerase encountering an OTA lesion at the replication fork.19 The major nucleoside lesion produced by OTA is the carbon-linked C8−2′-deoxyguanosine (dG) adduct, OT−dG, 2 (Figure 1), which was originally produced from the photoreaction of OTA in the presence of excess dG.23 The photochemical mechanism for OT−dG formation likely

INTRODUCTION Ochratoxin A, 1 (OTA; N-[[(3R)-5-chloro-8-hydroxy-3-methyl-1-oxo-7-isochromanyl]carbonyl]-3-phenyl-L-phenylalanine; Figure 1), is a tetrasubstituted chlorophenolic mycotoxin

Figure 1. Structures of OTA (1) and OT−dG (2).

produced by species of Aspergillus and Penicillium.1 The toxin is detected in a wide variety of foodstuffs2−5 and is difficult to remove from the food supply under normal processing or cooking conditions.2 OTA is considered to be a potent renal carcinogen6 and is classified as a possible (group 2B) human carcinogen by the International Agency for Research on Cancer due to sufficient evidence of OTA-mediated carcinogenicity in laboratory animals.7 Consequently, the levels of OTA in foodstuffs are regulated in many countries. For example, the European Union has set current limits of 5 ng/g for OTA in raw cereal grains and 3 ng/g in processed cereal products.8,9 The mechanism of action of OTA-mediated renal carcinogenicity has generated considerable debate on whether the toxin exerts carcinogenicity in rodents through the formation of DNA adducts (addition products) or by indirect mechanism(s).10−15 However, over the past decade the ability of OTA to act as a mutagenic DNA-reactive carcinogen has © XXXX American Chemical Society

Special Issue: Public Health Perspectives of Mycotoxins in Food Received: Revised: Accepted: Published: A

September 1, 2016 November 4, 2016 November 9, 2016 November 9, 2016 DOI: 10.1021/acs.jafc.6b03897 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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Figure 2. Three major conformations produced by C8−dG adducts in duplex DNA and the NarI(12) and NarI(22) oligonucleotide sequences.

Figure 3. syn and anti structures of C-linked C8−dG nucleoside adducts. The dihedral angle χ (∠(O4′C1′N9C4)) defines the glycosidic bond orientation to be anti (χ = 180 ± 90°) or syn (χ = 0 ± 90°), and θ (∠(N9C8C14C15) for OT−dG) defines the degree of twist between the nucleobase and the C8-aryl substituent.

involves a reductive dehalogenation process to afford Cl− and a carbon-centered OTA phenyl radical, which reacts covalently at the C8 site of dG.12,13,15 The same adduct is also produced through activation of OTA/dG with Fe2+.23 The OTA photochemistry was then used to generate the corresponding 3′-monophosphate-OT−dG adduct that was treated with polynucleotide kinase T4 and [γ-32P]ATP for conversion into a 3′,5′-bisphospho-OT−dG adduct to serve as a cochromatographic standard for 32P-postlabeling detection of OTAmediated DNA adduction in kidney tissue from OTA-treated male rats or pigs24 and in human kidney (HK2) cells.25 Oral dosing of rats or pigs with OTA generates several lesions with the major adduct spot comigrating with the postlabeled OT− dG adduct standard.24 In HK2 cells, OTA affords three adduct spots with the major lesion being OT−dG at a level of ∼18 adducts/109 nucleotides, based on the labeling efficiency of the 3′-monophosphate-OT−dG adduct standard.25 Given that other researchers have questioned the ability of OTA to react covalently with DNA10,11,14 and adduct spots in 32Ppostlabeling lack evidence for the presence of the OTA moiety, MS experiments were used to confirm OT−dG formation from oral dosing of male rats with OTA.26 A plausible in vivo mechanism for OT−dG formation stems from initial oxidative stress mediated by OTA for a buildup of O2•− leading to the liberation of Fe2+ for reductive dehalogenation of OTA required for phenyl radical production and C8−dG adduct formation.12 This mechanism stresses the importance of the C5 chlorine atom for OT−dG formation and provides a rationale for the inability of the nonchlorinated OTB derivative to react covalently with dG following photoirradiation,27 to show a lack

of genotoxicity in HK2 cells,27 and to exhibit much lower in vivo toxicity compared to OTA.28,29 The OT−dG lesion containing a direct C8−C linkage (denoted C-linked C8−dG adduct) is a fairly common adduct type, with carcinogenic aryl hydrazines,30,31 polycyclic aromatic hydrocarbons (PAHs),32 estrogens,33,34 and nitroaromatics35 all producing C-linked C8−dG adducts following metabolic activation. Other C8−dG adducts include N-linked derivatives produced by arylamine carcinogens36−38 and O-linked C8−dG adducts produced by phenolic toxins.39−41 For the N-linked adducts, a variety of common conformational preferences of the associated damaged DNA duplex have been well characterized, that is, the major groove (B-type), the base-displaced stacked or intercalated (S-type), and the minor groove wedge (W-type) conformations (Figure 2).36,37 Furthermore, N-linked C8−dG adducts that exhibit potent mutagenicity possess polycyclic aromatic structures and, despite conformational heterogeneity, favor the syn conformation that generates S-type or W-type duplexes.36,37 On the other hand, the single-ringed N-linked C8-aniline−dG adduct favors the B-type conformation and lacks potent mutagenicity.42 The unsubstituted phenolic Olinked C8−dG adduct also favors the B-type conformation43 and is replicated as a natural G.41,44 Although the N-linked and O-linked C8−dG adducts both contain a flexible (amine or ether) tether separating the dG component from the aryl ring, the bulky moiety of C-linked adducts is closer to the damaged nucleobase, and the conformational preferences and biological outcomes are less understood. Although the C-linked OT−dG adduct (Figure 1) has yet to be incorporated into DNA, we recently chemically synthesized a number of model C-linked C8−dG adducts having the same B

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five distinct OTA conformations.51 The four lowest energy conformers, which fall within 5 kJ/mol, were subsequently used to build starting orientations of the OT−G nucleobase adduct. Because the nucleobase models primarily provide insight into the relative orientation of the OT moiety with respect to the damaged G, the potential energy surface (PES) for this smallest model was explored as a function of rotation about the OT− nucleobase linkage, the dihedral angle θ defined as ∠(N9C8C14C15) (Figure 3). In the absence of discrete interactions between the bulky moiety and nucleobase, all four OTA conformers result in similar OT−G PESs, with the lowest energy minima adopting a (18−22°) twist about the OT−G linkage.51 Although information about the model C8−dG adducts with various ring sizes (Figure 3) at the nucleobase level is limited, the degree of twist for OT−G is greater than reported for C-linked C8-phenyl−G adducts containing various ortho or para substituents, which are planar at the nucleobase level.55,56 Subsequently, OT−dG nucleoside models were developed by adding deoxyribose to the three OTA conformers associated with the most stable OT−G nucleobase conformers. The DFT (B3LYP/6-31G(d)) PES for the three OT−dG models were each mapped as a function of rotation about the bonds dictating the relative orientation of the OT moiety with respect to the nucleobase (θ) and the relative orientation of the nucleobase with respect to deoxyribose, that is, the dihedral angle χ defined as ∠(O4′C1′N9C4) (Figure 3). Specifically, θ and χ were constrained in 10° increments from 0 to 360°, and the remainder of the model was relaxed. The resulting energy contour plots were very similar for each OTA conformer, which primarily differ in the phenylalanine moiety that does not interact with the nucleobase. Subsequent optimization of the three anti and three syn minima, as well as six transition states, isolated from each surface revealed a consistently greater twist about the OT−nucleobase bond upon incorporation of deoxyribose, approximately 20° on average. Furthermore, deoxyribose restricts rotation about the OT−nucleobase linker (θ), precluding some conformations adopted by the nucleobase model. Both the twist and reduced rotation about the OT− nucleobase bond arise due to close contacts between the OT moiety and deoxyribose, reflecting the size of this bulky moiety and suggesting that interactions within DNA helices may dictate the final lesion conformation. The increased twist about the linker reported for OT−dG upon addition of deoxyribose also occurs for C-linked C8-phenyl−dG adducts bearing different ortho or para substituents, which exhibit a twist about the bulky moiety−nucleobase bond of 27.0−55.3°.56 Furthermore, the model C-linked C8−dG adducts with various aryl ring sizes (Figure 3) exhibit a twist about the linker in nucleoside models that ranges between 16.2 and 58.8° (Table 1). Perhaps more importantly, the nucleoside models revealed that the syn glycosidic conformations are lower in energy than the anti conformations. This observation is consistent with DFT calculations for the model C-linked adducts of differing aryl ring sizes (Figure 3 and Table 1). In general, the greater stability of the syn conformation arises in part due to repulsive steric interactions between the sugar moiety and the C8 substituent in the anti orientation versus stabilizing O5′−H··· N3 hydrogen bonds in the syn conformation. For the model Clinked C8−dG adducts, the adduct bearing the smallest furyl aryl ring exhibits the smallest syn preference (19.3 kJ/mol), whereas the C8-quinolyl−dG (Q−dG) derivative displays the

linkage type as OT−dG but differing in aryl ring size (Figure 3).45,46 The C-linked nucleoside adducts were synthesized using palladium-catalyzed (Suzuki−Miyaura) cross-coupling between 8-Br-dG and the arylboronic acid47 and were structurally characterized by NMR, which was combined with density functional theory (DFT) calculations to establish their degree of syn preference.45,48 The model C-linked C8−dG adducts were then converted into phosphoramidites and inserted into the G3 position (X) of the NarI GC-repeat sequence (Figure 2) using solid-phase DNA synthesis.45,46 The NarI sequence was chosen for study because it is a hotspot for deletion mutations induced by polycyclic N-linked C8−dG adducts49 via a two-base slippage mechanism50 and, therefore, the structural and biochemical impact of N-linked C8−dG adducts has previously been well-studied in this sequence context for comparison. The C-linked adducted NarI oligonucleotides were characterized using MS and the conformational preferences of the C-linked adducts in the NarI(12) DNA duplex (i.e., B-type vs S-type vs W-type) were studied using optical spectroscopies (UV/vis thermal melting assays, circular dichroism (CD), and fluorescence) combined with molecular dynamics (MD) simulations.45,46 Primer elongation assays were carried out using the NarI(22) template annealed to a 15-mer primer in the presence of model polymerases, including the Klenow fragment exo− (Kf−) and the Y-family DNA polymerase IV (Dpo4) from Sulfolobus solfataricus. These studies established a relationship between Clinked adduct structure and the levels of base misincorporation, polymerase blockage, and polymerase slippage to yield deletion mutations.45,46 From this perspective, we draw comparisons between the structural data for model C-linked C8−dG adducts and the OT−dG lesion. Specifically, structural insight into OT−dG has been obtained using DFT calculations on the nucleoside51 and MD simulations on NarI(12) DNA duplexes containing the OT−dG adduct at G3.51,52 These data are directly compared to the information obtained for model C-linked adducts using analogous computational approaches. Subsequently, the correlation between the conformation induced by the C-linked C8−dG adducts in the NarI duplex and their mutagenicity based on primer extension assays using model polymerases will be summarized. When combined, these data predict the OT− dG lesion to exhibit more potent in vitro mutagenicity than that established for the model C-linked adducts. This analysis suggests that OT−dG could make an important contribution to the observed mutagenicity of OTA in rat16,17 and especially mice kidney,18,19 where deletions in repetitive G/C sequences were observed.19

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NUCLEOBASE AND NUCLEOSIDE ADDUCT STRUCTURES In the absence of experimental structural data on the OT−dG adduct, a range of DFT calculations and MD simulations have been used in conjunction with models of various sizes to gain insight into the conformational flexibility of the lesion.51,52 Despite the likely deprotonation of the OT group under physiological conditions, initial focus was placed on the neutral OT−dG adduct due to the large number of models considered and the anticipated conformational complexity of the OT−dG lesion. Starting from the seven lowest energy conformations of isolated OTA (prior to addition to dG) determined from crystallographic, NMR, and molecular mechanics (MM) data,53,54 DFT (B3LYP/6-31G(d)) calculations characterized C

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is imperative to obtain structural information about C-linked C8−dG adducts within DNA duplexes.

Table 1. DFT Structural Properties of C-Linked C8−dG Nucleoside Adducts aryl

Eanti − Esyna

χb

θb

Fur Ph Q BTh Py OTc

19.3 25.1 29.5 24.5 20.9 11.7

50.1 66.8 52.2 66.8 61.1 45.7

343.8 41.2 305.6 211.1 58.8 322.8

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ADDUCT STRUCTURES IN NarI(12) Preliminary 20 ns MD (AMBER) simulations were performed with the lowest energy anti and syn conformations of the neutral OT−dG lesion isolated from nucleotide models (i.e., the 5′-phosphate added to the nucleoside models discussed above) incorporated at G3 (X) in the Nar(12) duplex (Figure 2) and paired opposite complementary cytosine.51 When the lesion adopts the anti conformation, the OT moiety is projected into the major groove of DNA, and Watson−Crick hydrogen bonding is maintained with the opposing C. In the syn orientation, a portion of the OT moiety intercalates into the DNA helix to stack with the flanking pairs, whereas the remainder is located in the minor groove, and the opposing C is displaced into the major groove. These OT−dG adducted DNA structures mirror the so-called B-type and S-type conformation adopted by well-studied aromatic amine C8− dG adducts, respectively (Figure 2).36,37 In the case of OT−dG, calculations predict that helix association is equally probable for the two adducted DNA conformations, because the anti lesion conformation is stabilized by interstrand hydrogen bonding, whereas the syn adduct orientation is stabilized by intrahelical stacking interactions. These were the first structural data suggesting that OT−dG adducted DNA may exhibit complex conformational heterogeneity.51 In a follow-up study, a more rigorous sampling approach was implemented to gain greater structural insight into the preferred conformations of DNA containing OT−dG.52 Specifically, over 2.1 μs of production simulations, excluding significant equilibration steps and preliminary trials, was used to explore all conformational themes previously characterized for the N-linked aromatic amine C8−dG lesions. Initial conformations of adducted DNA were considered with the OT moiety in the major groove (anti B-type conformation), in the minor groove (syn W-type conformation), and intercalated into the helix (syn S-type conformation) (Figure 2). Furthermore, two rotamers that deviate in the orientation about the OT− nucleobase bond (θ, Figure 3) were investigated, and each of three ionization states of OT−dG (neutral, carboxylic ionized (monoanionic), and carboxylic and phenolic ionized (dianionic)) were considered at G3 in the NarI(12) duplex. Representative MD (AMBER) structures of the monoanionic OT−dG adducted DNA duplex conformations are presented in Figure 5. Table 2 contains relative molecular mechanics/Poisson− Boltzmann surface area continuum solvation model (MMPBSA) free energies between the three distinct adducted DNA conformations for OT−dG, as well as the corresponding data obtained from an analogous modeling procedure for the model C-linked C8−dG adducts paired opposite C.45,46 Also included in Table 2 are the thermal melting parameters obtained from UV/vis spectroscopy for the model adducts at G3 within the NarI(12) duplexes. The melting parameters suggest that the model C-linked C8−dG adducts strongly decrease duplex stability, with the magnitude depending on aryl ring size. For example, the bulky fused pyrenyl adduct (Py−G) exhibits a ΔTm value of −20 °C,46 whereas the corresponding value for the smaller furyl adduct (Fur−G) is −9 °C.45 For all model adducts, MM-PBSA analysis suggests a preference for the (anti) B-type conformation in which the C8-aryl ring is exposed to solvent in the major groove of the duplex. However, the

In kJ/mol. bIn degrees. χ and θ are defined in Figure 3. cNeutral lesion.

a

largest (29.5 kJ/mol) (Table 1). Comparison of the data for the model adducts versus OT−dG (Table 1) demonstrates a lower anti/syn energy difference for OT−dG, with the most stable anti and syn OT−dG conformers (Figure 4) differing in energy

Figure 4. Most stable anti (A) and syn (B) conformations of the neutral OT−dG lesion obtained from DFT calculations on the nucleoside model.

by only 11.7 kJ/mol. The low-energy anti OT−dG structure possess an O−H···O (1.82 Å) hydrogen bond between the 5′hydroxyl group of the sugar and the carbonyl oxygen of the OTA moiety (Figure 4), whereas an analogous stabilizing interaction is not feasible in the model adducts. The nucleoside studies of OT−dG versus the model Clinked C8−dG adducts of various aryl ring sizes (Figure 3) highlight the inherent flexibility about the OT−nucleobase and nucleobase−deoxyribose bonds in the OTA lesion compared to the simpler models. Nevertheless, the intramolecular attractive and repulsive interactions observed in the nucleoside models may be disrupted in the DNA duplex environment. Furthermore, stacking interactions with flanking nucleobase pairs may become important in dictating the anti/syn conformational preference, and the helical environment may impose different conformational restrictions depending on the size or chemical composition of the C8-aryl group. Therefore, it D

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Figure 5. Representative MD (AMBER) structures of OT−dG adducted DNA conformations with the bulky moiety in the major groove (A) or minor groove (B) and intercalated into the helix (C). Structures are shown for the monoanionic adduct at the G3 position of NarI(12).

to steric hindrance between a flanking base and the bulky OTA moiety, enhanced stacking between the OTA moiety and neighboring base pairs, enhanced (unfavorable) electrostatics upon increased charge, and/or hydrogen-bonding interactions between the displaced opposing C and DNA phosphate backbone. Most importantly, the complicated conformational heterogeneity predicted by MD simulations suggests that different cellular environments may stabilize distinctive conformations, and therefore the biological outcomes of OT−dG will be vastly diverse.

Table 2. Thermal Melting Parameters and Relative Energies of Nar(12) Duplex Conformations Containing a C-Linked C8−dG Adduct X

Tma

ΔTmb

B-typec

Fur−G Ph−G Q−G BTh−G Py−G OT−G OT−G− OT−G2−

55 48 45 48 44

−9 −16 −19 −16 −20

0.0 0.0 0.0 0.0 0.0 0.0 29.3 0.0

S-typec

48.5 7.9 7.1 12.1 34.7

W-typec 14.2 26.4 25.1 18.4

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7.1 0.0 2.9

IN VITRO MUTAGENICITY OF MODEL C-LINKED C8−dG ADDUCTS To demonstrate the relationship between adduct conformation and in vitro mutagenicity, five model adducts with differing ring types (Figure 3) were incorporated into G3 (X) of NarI(22) to carry out primer elongation experiments.45,46 Two polymerases, namelym Escherichia coli DNA polymerase I Klenow fragment exo− (Kf−) and DNA polymerase IV from Sulfolobus solfataricus P2 (Dpo4), were used to model in vivo replication of C-linked C8−dG adducts. High-fidelity polymerases, such as Kf−, favor accurate replication when correct WC base pairing is established with the incoming deoxynucleotide triphosphate (dNTP).57 Bulky DNA adducts often block DNA replication by high-fidelity polymerases through distortion of the DNA duplex or polymerase active site. This is thought to trigger the recruitment of Y-family translesion (bypass) polymerases for potential error-free extension past the bulky adduct after polymerase switching.57−59 The Y-family polymerases have spacious active sites that can accommodate bulky DNA lesions while facilitating low-fidelity DNA replication. Dpo4 is regarded as a prototypical Y-family polymerase that serves as an excellent model to investigate how structural features of adducts determine lesion bypass efficiency and fidelity.38 To determine the miscoding potential of model C-linked C8−dG adducts, single-nucleotide and full-length insertion assays were performed with the NarI(22):15mer template:primer and Kf− or Dpo4 in the presence of dNTP(s). The single-nucleotide assays indicated that the greatest increased levels of misincorporation (A and G) by Kf− occur when the smallest Fur−G adduct is present in the template. This observation is consistent with the greater syn preference of the Fur−G nucleoside predicted by DFT (Table 1) and greater conformational flexibility of Fur−G adducted DNA predicted

a Tm values (°C) of duplexes (6 μM) measured in 50 mM sodium phosphate, pH 7, with 0.1 M NaCl; errors are ±1 °C. bΔTm = Tm (modified duplex) − Tm (unmodified duplex). cRelative MM-PBSA free energies (kJ mol−1) of B-type, S-type, and W-type duplex conformations.

smallest Fur−G adduct exhibits the smallest relative energy between the anti B-type and syn minor groove W-type conformations (14.2 kJ/mol),45 suggesting that the smallest ring is well accommodated in the minor groove, whereas the bulky Py−G adduct possesses the smallest relative energy between the anti B-type and syn S-type conformations (7.9 kJ/ mol) due to stronger intrahelical stacking interactions for this largest aryl group.46 In contrast to the model C-linked C8−dG adducts, OT−dG can acquire a number of different ionization states, and MD simulations suggest that the preferred conformation of the NarI(12) duplex containing OT−G is highly dependent on the adduct ionization state.52 Specifically, the neutral OT−G lesion preferentially induces the B-type conformation, but leads to small relative energies for the S-type and W-type conformations (7.1 kJ/mol). In contrast, the monoanionic OT−G− lesion preferentially results in the minor groove W-type conformation, with a high relative energy for the B-type conformation (29.3 kJ/mol) and the S-type conformer falling 12.1 kJ/mol above the W conformer. The dianionic OT−G2− adduct prefers to induce the groove duplex conformations (B-type and W-type, with an energy difference of only 2.9 kJ/mol), but is associated with a relatively high energy S-type conformation (34.7 kJ/ mol). This complex conformational profile arises due to differences in discrete interactions at the lesion site, including features such as the loss of Hoogsteen hydrogen bonding due E

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Figure 6. Proposed slippage mechanism induced by model C-linked C8−dG adducts within the NarI sequence, as well as representative structures from MD simulations of anti BTh-G and syn Py−G in the two-base bulged slipped intermediate.

by MD, that is, small free energy difference between B-type and W-type conformers (Table 2) compared to the other C-linked C8-aryl−dG adducts with larger C8-aryl substituents. With the adduct in the syn conformation, the Hoogsteen hydrogenbonding face can base pair with G and A, which provides a rational for the ability of C8-aryl−dG adducts to induce G → T and G → C transversion mutations.45 In the full-length primer extension assays, the smallest Fur−G adduct was extended most efficiently by Kf−,45 whereas bulkier lesions more significantly hinder extension one base past the adduct site than incorporation opposite the adduct.45,46 Primer extension of the adducted NarI(22) templates by Dpo4 demonstrated a strong tendency for template slippage, which led to the proposed mechanism outlined in Figure 6. In the starting NarI(22):15mer template:primer, the C-linked adducts are present in the favored syn conformation at the single strand-double strand junction. Insertion of C (step 1) forces the polymerase to flip the adduct from its stable syn conformation into the energetically destabilized anti conformation, which is expected to stall DNA replication compared to replication of the unmodified template. This stalling would allow for slippage and production of the two-base bulge slipped-mutagenic intermediate (step 2), as proposed for Nlinked C8−dG adducts.50 Following two-base bulge formation, the polymerase can insert a second C opposite G at position 4 in the template strand, included in step 2 (Figure 6). The resulting two-base bulged duplexes are anticipated to exist as an equilibrium mixture with the adduct in the anti or syn conformations. MD simulations suggest the bulkier Q−G and Py−G adducts favor the syn conformation within the two-base bulge by approximately 10−25 kJ/mol, whereas the smaller Fur−G, phenyl−G (Ph−G), and benzo[b]thienyl−G (BTh− G) adducts favor the anti geometry by approximately 15−30 kJ/mol. The anti structure is proposed to undergo a realignment process (step 3) to permit additional C incorporation by Dpo4 across from positions 3 and 4 in the template strand (step 4). This proposal rationalizes the incorporation of four C bases in the single nucleotide incorporation assays by Dpo4 for Fur−G, Ph−G, and BTh−

G, which generates a C:C mismatch at position 2 of the template strand.45,46 In contrast, realignment of the primer:template when Q−G or Py−G is present is disfavored because both adducts prefer the syn conformation within the slippedmutagenic intermediate (Figure 6). Thus, only two C bases were inserted by Dpo4 under single-nucleotide incorporation conditions for Q−G or Py−G, which is classical evidence for two-base slippage.60 Overall, the primer extension analysis of the model C-linked C8−dG adducts provides evidence of the ability of C-linked C8−dG aryl adducts to promote G → T and G → C transversion mutations due to a syn conformational preference. These adducts also effectively stall DNA replication to permit slippage of the primer at repetitive GC sequences, which will afford deletion and base substitution mutations.

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IMPLICATIONS FOR IN VITRO MUTAGENICITY BY OT−dG The primer extension assays for the model C-linked C8−dG adducts demonstrate their propensity to induce base substitution and deletion mutations in G/C repeat sequences.45,46 The smallest adduct tested (Fur−G) exhibited the greatest levels of A and G misincorporation and the most accessible W-type duplex structure when the lesion adopts the syn conformation. The MD simulations for the OT−dG adduct (Table 2) predict greater conformational flexibility, with the Wtype structure being energetically preferred for the monoanionic lesion and readily accessible for the dianionic adduct. Because hydrogen bonding between the damaged base and incoming mismatched dNTPs is more stable when the adduct assumes the syn conformation, increased conformational flexibility of bulky DNA adducts has been shown to enhance misincorporation by Y-family DNA polymerases.61 Thus, the OT−dG lesion is predicted to exhibit greater in vitro mutagenicity than the model C-linked C8−dG adducts experimentally studied to date. OTA in mouse kidney exhibits a propensity to lead to single-base deletions in repetitive G/C sequences, single-base substitutions, and insertions.19 The specificity for OTA-mediated deletions in repetitive G/C sequences is in keeping with the well-known tendency of F

DOI: 10.1021/acs.jafc.6b03897 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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ochratoxin A; PES, potential energy surface; Ph, phenyl; Pyr, pyrenyl; ROS, reactive oxygen species

C8−dG adducts to induce polymerase slippage and frameshift mutations in GC-repeat sequences, such as the G3 site of NarI.45,46,49,50,61 Thus, it is conceivable that the OT−dG adduct plays a contributing factor in promoting OTA-mediated mutagenicity and carcinogenicity.

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(1) Pfohl-Leszkowicz, A. An overview on toxicity and carcinogenicity in animals and humans. Mol. Nutr. Food Res. 2007, 51, 61−99. (2) Lee, H. J.; Ryu, D. Significance of ochratoxin A in breakfast cereals from the United States. J. Agric. Food Chem. 2015, 63, 9404− 9409. (3) Duarte, S.; Pena, A.; Lino, C. A review on ochratoxin A occurrence and effect of processing of cereal and cereal derived food products. Food Microbiol. 2010, 27, 187−198. (4) Lombaert, G. A.; Pellaers, P.; Neumann, G.; Kitchen, D.; Huzel, V.; Trelka, R.; Kotello, S.; Scott, P. M. Ochratoxin A in dried vine fruits on the Canadian retail market. Food Addit. Contam. 2004, 21, 578−585. (5) Jørgensen, K. Occurrence of ochratoxin A in commodities and processed fooda review of EU occurrence data. Food Addit. Contam. 2005, 22, 26−30. (6) Boorman, G. NTP Technical Report on the Toxicology and Carcinogenesis Studies of Ochratoxin A (CAS No. 303-47-9) in F344/N Rats (Gavage Studies); NIH Publication 89-2813; U.S. Department of Health and Human Services: Research Triangle Park, NC, USA, 1989. (7) IARC Monographs on the Evaluation of Carcinogenic Risks to Humans. Some Naturally Occurring Substances: Food Items and Constituents, Heterocyclic Aromatic Amines and Mycotoxins. IARC: Geneva, Switzerland, 1992; Vol. 58. (8) European Commission Regulation EC No. 466/2001setting maximum levels for certain contaminants in foodstuffs. Off. J. Eur. Communities 2002, 472/2002, 18−20. (9) European Commission Regulation EC No. 466/2001 as regards ochratoxin A. Off. J. Eur. Communities 2002, 123/2005, 3−5. (10) Mally, A.; Zepnik, H.; Wanek, P.; Eder, E.; Dingley, K.; Ihmels, H.; Völkel, W.; Dekant, W. Ochratoxin A: lack of formation of covalent DNA adducts. Chem. Res. Toxicol. 2004, 17, 234−242. (11) Turesky, R. J. Perspective: ochratoxin A is not a genotoxic carcinogen. Chem. Res. Toxicol. 2005, 18, 1082−1090. (12) Manderville, R. A. A case for the genotoxicity of ochratoxin A by bioactivation and covalent DNA adduction. Chem. Res. Toxicol. 2005, 18, 1091−1097. (13) Manderville, R. A.; Pfohl-Leszkowicz, A. Bioactivation and DNA adduction as a rationale for ochratoxin A carcinogenesis. World Mycotoxin J. 2008, 1, 357−367. (14) Adler, M.; Müller, K.; Rached, E.; Dekant, W.; Mally, A. Modulation of key regulators of mitosis linked to chromosomal instability is an early event in ochratoxin A carcinogenicity. Carcinogenesis 2009, 30, 711−719. (15) Pfohl-Leszkowicz, A.; Manderville, R. A. An update on direct genotoxicity as a molecular mechanism of ochratoxin A carcinogenicity. Chem. Res. Toxicol. 2012, 25, 252−262. (16) Hibi, D.; Suzuki, Y.; Ishii, Y.; Jin, M.; Watanabe, M.; SugitaKonishi, Y.; Yanai, T.; Nohmi, T.; Nishikawa, A.; Umemura, T. Sitespecific in vivo mutagenicity in the kidney of gpt delta rats given a carcinogenic dose of ochratoxin A. Toxicol. Sci. 2011, 122, 406−414. (17) Kuroda, K.; Hibi, D.; Ishii, Y.; Takasu, S.; Kijima, A.; Matsushita, K.; Masumura, K.-i.; Watanabe, M.; Sugita-Konishi, Y.; Sakai, H.; Yanai, T.; Nohmi, T.; Ogawa, K.; Umemura, T. Ochratoxin A induces DNA double-strand breaks and large deletion mutations in the carcinogenic target site of gpt delta rats. Mutagenesis 2014, 29, 27−36. (18) Hibi, D.; Kijima, A.; Suzuki, Y.; Ishii, Y.; Jin, M.; Sugita-Konishi, Y.; Yanai, T.; Nishikawa, A.; Umemura, T. Effects of p53 knockout on ochratoxin A-induced genotoxicity in p53-deficient gpt delta mice. Toxicology 2013, 304, 92−99. (19) Kuroda, K.; Hibi, D.; Ishii, Y.; Yokoo, Y.; Takasu, S.; Kijima, A.; Matsushita, K.; Masumura, K.; Kodama, Y.; Yanai, T.; Sakai, H.; Nohmi, T.; Ogawa, K.; Umemura, T. Role of p53 in the progression from ochratoxin A-induced DNA damage to gene mutations in the kidneys of mice. Toxicol. Sci. 2015, 144, 65−76.

FUTURE STUDIES So far, the site-specific incorporation of the OT−dG adduct into an oligonucleotide substrate has yet to be achieved. One issue with the chemical synthesis of the OT−dG nucleoside on a scale suitable for conversion into a phosphoramidite needed for solid-phase DNA synthesis is the sensitivity of the lactone moiety to the strongly basic conditions (promotes lactone ring opening) employed in Suzuki−Miyaura cross-coupling. Variation to the synthetic approach for incorporation of the model C-linked C8−dG adducts into the G3 site of the NarI sequence may be required for insertion of OT−dG so that a direct comparison to our results for the model C-linked C8−dG adducts can be established. Here, it will be important to determine the adduct ionization state, its preferred conformation(s) in duplex DNA, and its impact on DNA replication by model polymerases, such as Kf− and Dpo4. Generation of adducted oligonucleotide strands containing OT−dG would then pave the way for incorporation into DNA vectors for the determination of mutational frequency in cell-based assays. If a strong correlation between OTA-mediated mutagenicity in rodents and mutations induced by the OT−dG lesion is established, this would provide a chemical basis for the mechanism of action for OTA renal carcinogenicity in rodents, which is an unresolved issue.

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REFERENCES

AUTHOR INFORMATION

Corresponding Authors

*(R.A.M.) Phone: (519) 824-4120, x53963. Fax: (519) 7661499. E-mail: [email protected]. *(S.D.W.) Phone: (403) 329-2323. Fax: (403) 329-2057. Email: [email protected]. ORCID

Richard A. Manderville: 0000-0003-4035-8093 Funding

This project was supported by the Natural Sciences and Engineering Research Council of Canada (Discovery 3116002013 to R.A.M. and 249598-07 to S.D.W.), the Canada Research Chairs Program (950-228175 to S.D.W.), the Canadian Foundation of Innovation (10679 to R.A.M. and 22770 to S.D.W.) and the Ontario Research Fund (10679 to R.A.M.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS Computational resources provided by Westgrid and Compute/ Calcul Canada are greatly appreciated. R.A.M. thanks Lauren Jackson (FDA/CFSAN) and Dojin Ryu (University of Idaho) for the kind invitation to present this research at the ACS National Meeting in San Diego, CA, USA, March 2016.

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ABBREVIATIONS USED DFT, density functional theory; dG, 2′-deoxyguanosine; dNTP, deoxynucleotide triphosphate; Dpo4, DNA polymerase IV; DSB, double strand break; BTh, benzo[b]thienyl; Fur, furyl; Kf−, Klenow fragment exo−; MD, molecular simulations; OTA, G

DOI: 10.1021/acs.jafc.6b03897 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

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Journal of Agricultural and Food Chemistry (20) Jackson, S. P. Sensing and repairing DNA double-strand breaks. Carcinogenesis 2002, 23, 687−696. (21) White, R. R.; Milholland, B.; de Bruin, A.; Curran, S.; Laberge, R.-M.; van Steeg, H.; Campisi, J.; Maslov, A. Y.; Vijg, J. Controlled induction of DNA double-strand breaks in the mouse liver induces features of tissue ageing. Nat. Commun. 2015, 6, 6790. (22) Shigenaga, M. K.; Gimeno, C. J.; Ames, B. N. Urinary 8hydroxy-2′-deoxyguanosine as a biological marker of in vivo oxidative DNA damage. Proc. Natl. Acad. Sci. U. S. A. 1989, 86, 9697−9701. (23) Dai, J.; Wright, M. W.; Manderville, R. A. Ochratoxin A forms a carbon-bonded C8-deoxyguanosine nucleoside adduct: Implication for C8-reactivity by a phenolic radical. J. Am. Chem. Soc. 2003, 125, 3716− 3717. (24) Faucet, V.; Pfohl-Leszkowicz, A.; Dai, J.; Castegnaro, M.; Manderville, R. A. Evidence for covalent DNA adduction by ochratoxin A following chronic exposure to rat and subacute exposure to pig. Chem. Res. Toxicol. 2004, 17, 1289−1296. (25) Tozlovanu, M.; Faucet-Marquis, V.; Pfohl-Leszkowicz, A.; Manderville, R. A. Genotoxicity of the hydroquinone metabolite of ochratoxin A: structure-activity relationships for covalent DNA adduction. Chem. Res. Toxicol. 2006, 19, 1241−1247. (26) Mantle, P. G.; Faucet-Marquis, V.; Manderville, R. A.; Squillaci, B.; Pfohl-Leszkowicz, A. Structures of covalent adducts between DNA and ochratoxin A: a new factor in debate about genotoxicity and human risk assessment. Chem. Res. Toxicol. 2010, 23, 89−98. (27) Hadjeba-Medjdoub, K.; Tozlovanu, M.; Pfohl-Leszkowicz, A.; Frenette, C.; Paugh, R. J.; Manderville, R. A. Structure-activity relationships imply different mechanisms of action for ochratoxin Amediated cytotoxicity and genotoxicity. Chem. Res. Toxicol. 2012, 25, 181−190. (28) Heussner, A. H.; Bingle, L. E. H. Comparative ochratoxin toxicity: a review of the available data. Toxins 2015, 7, 4253−4282. (29) Mally, A.; Keim-Heusler, H.; Amberg, A.; Kurz, M.; Zepnik, H.; Mantle, P.; Völkel, W.; Hard, G. C.; Dekant, W. Biotransformation and nephrotoxicity of ochratoxin B in rats. Toxicol. Appl. Pharmacol. 2005, 206, 43−53. (30) Kohda, K.; Tsunomoto, H.; Kasamatsu, T.; Sawamura, F.; Terashima, I.; Shibutani, S. Synthesis and miscoding specificity of oligodeoxynucleotide containing 8-phenyl-2′-deoxyguanosine. Chem. Res. Toxicol. 1997, 10, 1351−1358. (31) Gannett, P. M.; Lawson, T.; Miller, M.; Thakkar, D. D.; Lord, J. W.; Yau, W. M.; Toth, B. 8-Arylguanine adducts from arenediazonium ions and DNA. Chem.−Biol. Interact. 1996, 101, 149−164. (32) Rogan, E. G.; Cavalieri, E. L.; Tibbels, S. R.; Cremonesi, P.; Warner, C. D.; Nagel, D. L.; Tomer, K. B.; Cerny, R. L.; Gross, M. L. Synthesis and identification of benzo[a]pyrene-guanine nucleoside adducts formed by electrochemical oxidation and by horseradish peroxidase catalyzed reaction of benzo[a]pyrene with DNA. J. Am. Chem. Soc. 1988, 110, 4023−4029. (33) Akanni, A.; Abul-Hajj, Y. J. Estrogen-nucleic acid adducts: dissection of the reaction of 3,4-estrone quinone and its radical anion and radical cation with deoxynucleosides and DNA. Chem. Res. Toxicol. 1999, 12, 1247−1253. (34) Akanni, A.; Abul-Hajj, Y. J. Estrogen-nucleic acid adducts: Reaction of 3,4-estrone-o-quinone radical anion with deoxyribonucleosides. Chem. Res. Toxicol. 1997, 10, 760−766. (35) Enya, T.; Kawanishi, M.; Suzuki, H.; Matsui, S.; Hisamatsu, Y. An unusual DNA adduct derived from the powerfully mutagenic environmental contaminant 3-nitrobenzanthrone. Chem. Res. Toxicol. 1998, 11, 1460−1467. (36) Patel, D. J.; Mao, B.; Gu, Z.; Hingerty, B. E.; Gorin, A.; Basu, A. K.; Broyde, S. Nuclear magnetic resonance solution structures of covalent aromatic amine−DNA adducts and their mutagenic relevance. Chem. Res. Toxicol. 1998, 11, 391−407. (37) Cho, B. Structure−function characteristics of aromatic amine− DNA adducts. In The Chemical Biology of DNA Damage; Wiley-VCH Verlag, 2010; pp 217−238. (38) Rechkoblit, O.; Kolbanovskiy, A.; Malinina, L.; Geacintov, N. E.; Broyde, S.; Patel, D. J. Mechanism of error-free and semitargeted

mutagenic bypass of an aromatic amine lesion by Y-family polymerase Dpo4. Nat. Struct. Mol. Biol. 2010, 17, 379−389. (39) Dai, J.; Wright, M. W.; Manderville, R. A. An oxygen-bonded C8-deoxyguanosine nucleoside adduct of pentachlorophenol by peroxidase activation: evidence for ambident C8 reactivity by phenoxyl radicals. Chem. Res. Toxicol. 2003, 16, 817−821. (40) Dai, J.; Sloat, A. L.; Wright, M. W.; Manderville, R. A. Role of phenoxyl radicals in DNA adduction by chlorophenol xenobiotics following peroxidase activation. Chem. Res. Toxicol. 2005, 18, 771− 779. (41) Witham, A. A.; Verwey, A. M. R.; Sproviero, M.; Manderville, R. A.; Sharma, P.; Wetmore, S. D. Chlorine functionalization of a model phenolic C8-guanine adduct increases conformational rigidity and blocks extension by a Y-family DNA polymerase. Chem. Res. Toxicol. 2015, 28, 1346−1356. (42) Shapiro, R.; Ellis, S.; Hingerty, B. E.; Broyde, S. Effect of ring size on conformations of aromatic amine-DNA adducts: the aniline− C8 guanine adduct resides in the B-DNA major groove. Chem. Res. Toxicol. 1998, 11, 335−341. (43) Kuska, M. S.; Witham, A. A.; Sproviero, M.; Manderville, R. A.; Majdi Yazdi, M.; Sharma, P.; Wetmore, S. D. Structural influence of C8-phenoxy-guanine in the NarI recognition DNA sequence. Chem. Res. Toxicol. 2013, 26, 1397−1408. (44) Verwey, A. M. R.; Witham, A. A.; Li, M.; Manderville, R. A. Mutagenicity analysis of C8-phenoxy-guanine in the NarI recognition DNA sequence. J. Toxins 2014, 1, 6. (45) Sproviero, M.; Verwey, A. M. R.; Rankin, K. M.; Witham, A. A.; Soldatov, D. V.; Manderville, R. A.; Fekry, M. I.; Sturla, S. J.; Sharma, P.; Wetmore, S. D. Structural and biochemical impact of C8-arylguanine adducts within the NarI recognition DNA sequence: influence of aryl ring size on targeted and semi-targeted mutagenicity. Nucleic Acids Res. 2014, 42, 13405−13421. (46) Sproviero, M.; Verwey, A. M. R.; Witham, A. A.; Manderville, R. A.; Sharma, P.; Wetmore, S. D. Enhancing bulge stabilization through linear extension of C8-aryl-guanine adducts to promote polymerase blockage or strand realignment to produce a C:C mismatch. Chem. Res. Toxicol. 2015, 28, 1647−1658. (47) Omumi, A.; Beach, D. G.; Baker, M.; Gabryelski, W.; Manderville, R. A. Postsynthetic guanine arylation of DNA by Suzuki-Miyaura cross-coupling. J. Am. Chem. Soc. 2011, 133, 42−50. (48) Blanchard, D. J. M.; Fadock, K. L.; Sproviero, M.; Deore, P.; Cservenyi, T. Z.; Manderville, R. A.; Sharma, P.; Wetmore, S. D. Photophysical properties of push-pull 8-aryl-deoxyguanosine probes within duplex and G-quadruplex structures. J. Mater. Chem. C 2016, 4, 2915−2924. (49) Fuchs, R. P. P.; Schwartz, N.; Daune, M. P. Hot spots of frameshift mutations induced by the ultimate carcinogen N-acetoxy-N2-acetylaminofluorene. Nature 1981, 294, 657−659. (50) Hoffmann, G. R.; Fuchs, R. P. P. Mechanisms of frameshift mutations: Insight from aromatic amines. Chem. Res. Toxicol. 1997, 10, 347−359. (51) Sharma, P.; Manderville, R. A.; Wetmore, S. D. Modeling the conformational preference of the carbon-bonded covalent adduct formed upon exposure of 2′-deoxyguanosine to ochratoxin A. Chem. Res. Toxicol. 2013, 26, 803−816. (52) Sharma, P.; Manderville, R. A.; Wetmore, S. D. Structural and energetic characterization of the major DNA adduct formed from the food mutagen ochratoxin A in the NarI hotspot sequence: influence of adduct ionization on the conformational preferences and implications for the NER propensity. Nucleic Acids Res. 2014, 42, 11831−11845. (53) Bredenkamp, M. W.; Dillen, J. L. M.; van Rooyen, P. H.; Steyn, P. S. Crystal structures and conformational analysis of ochratoxin A and B: probing the chemical structure causing toxicity. J. Chem. Soc., Perkin Trans. 2 1989, 1835−1839. (54) Dais, P.; Stefanaki, I.; Fragaki, G.; Mikros, E. Conformational analysis of ochratoxin A by NMR spectroscopy and computational molecular modeling. J. Phys. Chem. B 2005, 109, 16926−16936. (55) Schlitt, K. M.; Sun, K. M.; Paugh, R. J.; Millen, A. L.; NavarroWhyte, L.; Wetmore, S. D.; Manderville, R. A. Concerning the H

DOI: 10.1021/acs.jafc.6b03897 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Perspective

Journal of Agricultural and Food Chemistry hydrolytic stability of 8-aryl-2′-deoxyguanosine nucleoside adducts: implications for abasic site formation at physiological pH. J. Org. Chem. 2009, 74, 5793−5802. (56) Millen, A. L.; McLaughlin, C. K.; Sun, K. M.; Manderville, R. A.; Wetmore, S. D. Computational and experimental evidence for the structural preference of phenolic C-8 purine adducts. J. Phys. Chem. A 2008, 112, 3742−3753. (57) Broyde, S.; Wang, L.; Rechkoblit, O.; Geacintov, N. E.; Patel, D. J. Lesion processing: high-fidelity versus lesion-bypass DNA polymerases. Trends Biochem. Sci. 2008, 33, 209−219. (58) Ling, H.; Boudsocq, F.; Woodgate, R.; Yang, W. Crystal structure of a Y-family DNA polymerase in action: a mechanism for error-prone and lesion-bypass replication. Cell 2001, 107, 91−102. (59) Guengerich, F. P. Interactions of carcinogen-bound DNA with individual DNA polymerases. Chem. Rev. 2006, 106, 420−452. (60) Stover, J. S.; Chowdhury, G.; Zang, H.; Guengerich, F. P.; Rizzo, C. J. Translesion synthesis past the C8- and N2-deoxyguanosine adducts of the dietary mutagen 2-amino-3-methylimidazo[4,5-f ]quinoline in the NarI recognition sequence by prokaryotic DNA polymerases. Chem. Res. Toxicol. 2006, 19, 1506−1517. (61) Perlow-Poehnelt, R. A.; Likhterov, I.; Wang, L.; Scicchitano, D. A.; Geacintov, N. E.; Broyde, S. Increased flexibility enhances misincorporation. Temperature effects on nucleotide incorporation opposite a bulky carcinogen-DNA adduct by a Y-family DNA polymerase. J. Biol. Chem. 2007, 282, 1397−1408.

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DOI: 10.1021/acs.jafc.6b03897 J. Agric. Food Chem. XXXX, XXX, XXX−XXX