Molecular Modeling of the Major DNA Adduct Formed from Food

Subscriber access provided by UNIV OF NEWCASTLE

Article

Molecular Modeling of the Major DNA Adduct Formed from the Food Mutagen Ochratoxin A in NarI 2-base Deletion Duplexes: Impact of Sequence Context and Adduct Ionization on Conformational Preference and Mutagenicity Preetleen Kathuria, Purshotam Sharma, Richard A. Manderville, and Stacey D Wetmore Chem. Res. Toxicol., Just Accepted Manuscript • DOI: 10.1021/acs.chemrestox.7b00103 • Publication Date (Web): 18 Jul 2017 Downloaded from http://pubs.acs.org on July 23, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Chemical Research in Toxicology is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Chemical Research in Toxicology

Molecular Modeling of the Major DNA Adduct Formed from the Food Mutagen Ochratoxin A in NarI 2-base Deletion Duplexes: Impact of Sequence Context and Adduct Ionization on Conformational Preference and Mutagenicity Preetleen Kathuria,1 Purshotam Sharma,1 Richard A. Manderville2 and Stacey D. Wetmore3* 1Department

of Chemistry and Centre for Advanced Studies in Chemistry, Panjab

University, Chandigarh, India 160014.

2Departments

of Chemistry and Toxicology,

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

*Address: Department of Chemistry & Biochemistry University of Lethbridge E850, University Hall 4401 University Drive Lethbridge, Alberta, Canada T1K 3M4 E-mail: [email protected] Telephone: (403) 329-2323. Fax: (403) 329-2057

ACS Paragon Plus Environment

1


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 35

TOC-Graphic


2

Page 3 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


ABSTRACT: Exposure to ochratoxin A (OTA), a possible human carcinogen, leads to many different DNA mutations. As a first step toward understanding the structural basis of OTA-induced mutagenicity, the present work uses a robust computational approach and a slipped mutagenic intermediate model previously studied for C8-dG aromatic amine adducts to analyze the conformational features of post-replication 2-base deletion DNA duplexes containing OT-dG, the major OTA lesion at the C8 position of guanine. Specifically, a total of 960 ns of molecular dynamics simulations (excluding trial simulations) were carried out on four OT-dG ionization states in three sequence contexts within oligomers containing the NarI recognition sequence, a known hotspot for deletion mutations induced by related adducts formed from known carcinogens. Our results indicate that the structural properties and relative stability of the competing ‘major groove’ and ‘stacked’ conformations of OTA adducted 2-base deletion duplexes depend on both the OTA ionization state and the sequence context, mainly due to conformation-dependent deviations in discrete local (hydrogen-bonding and stacking) interactions at the lesion site, as well as DNA bending. When the structural characteristics of the OT-dG adducted 2-base deletion duplexes are compared to those associated with previously studied C8-dG adducts, a greater understanding of the effects of the nucleobase−carcinogen linkage and size of the carcinogenic moiety on the conformational preferences of damaged DNA is obtained. Most importantly, our work predicts key structural features for OT-dG-adducted deletion DNA duplexes, which in turn allows us to develop hypotheses regarding OT-dG replication outcomes. Thus, our computational results are valuable for the design and interpretation of future biochemical studies on the potentially carcinogenic OT-dG lesion.


3


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 35

INTRODUCTION Ochratoxin A (OTA) is a secondary metabolite of the Aspergillus1,

2

and

Penicillium3 species of fungi, which grow on a wide range of food crops, such as those associated with legumes, cereals, coffee, wines and fruit juices.4, 5 Exposure to OTA can cause serious conditions including renal carcinogenesis in rodents,6 and urinary tract,7 esophageal8 and testicular cancer9 in humans. Consequently, OTA has been categorized as a possible human carcinogen (Group 2B) by the International Agency for Research on Cancer (IARC).10 Despite significant progress in understanding OTA-mediated carcinogenesis,11 the mechanism of action (MoA) of OTA induced toxicity remains elusive. Specifically, OTA-associated carcinogenesis has been argued to be caused either by a direct mechanism that involves OTA bioactivation and subsequent reaction with DNA to form adducts (addition products),12 or by an indirect mechanism that involves participation of OTA in events such as oxidative DNA damage or disruption of mitosis.11-13 This current ambiguity in the MoA has hampered consensus among health agencies around the world regarding the tolerable daily intake of OTA.14 Despite the controversy surrounding the MoA of OTA, recent studies have ruled out the involvement of oxidative DNA damage,15 and support the role of DNA adducts in OTA-mediated carcinogenesis.12, 15, 16 As a result, considerable effort has been devoted over the past decade to identify and characterize OTA−DNA adducts. Most notably, in

vitro studies on animal tissues treated with OTA used

32P−postlabelling

of the DNA

nucleotides to provide evidence for the formation of bulky OTA−DNA adducts, specifically with 2′−deoxyguanosine monophosphate.17-19 Further spectroscopic studies characterized three distinct OTA-dG covalent adducts, viz. C-linked C8-OT-dG,20 O-linked C8-OTA-dG21 and N,N-linked N1,N2-OTHQ-dG.22 Among these lesions, the OT-dG adduct


4

Page 5 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(Figure 1a) was found to be the most prevalent.23, 24 More recent studies have focused on understanding the mutagenicity associated with OTA. For example, an in vitro study on the SupF gene of the mutation reporter plasmid PS189 in human Ad293 cells25 and the red/gam gene (Spi−) at the renal outer medulla of rodents15 revealed increased mutation frequencies upon OTA exposure. Further analysis of the mutational spectrum in Spi− mutants reported changes in specific mutation frequencies for all types of mutations observed in control cells upon OTA exposure.16 Site-specific incorporation of adducts into DNA duplexes is widely considered to be a critical first step toward gaining structural insights into the role of adduct formation

in

the

associated

mutagenicity.

However,

despite

experimental

characterization of the structural features of DNA containing a range of potentially carcinogenic adducts,26-29 the site-specific incorporation of OTA adducts into DNA has yet to be reported. Consequently, no experimental NMR or X-ray studies on OT-dG adducts in DNA are presently available. Previous studies on DNA damaged by a variety of genotoxins, including aristolochic acids,30-32 phenols,33-35 peroxynitrosocarbonates36 and benzo[a]pyrene,37 have shown that molecular dynamics (MD) simulations provide an approach for predicting solution conformations. In this context, previous work from our group used state-of-the-art computational methods to analyze the conformational themes of DNA duplexes containing OT-dG paired opposite complementary cytosine at the G1, G2 or G3 site in a 12-mer oligonucleotide (5′–CTCG1G2CG3CCATC–3′) containing the NarI sequence (underlined),32,33 which is the most vulnerable hotspot for deletion mutations induced by the acetylaminofluorene adduct of dG (AAF-dG) adduct in bacteria.38 Rigorous MD simulations predicted a mixture of OTA-adducted DNA conformations, irrespective of the lesion site sequence context.39 Each of these conformations was proposed to interact differently with DNA polymerases and repair


5


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 35

enzymes, and thereby may contribute to the varied mutagenic outcomes observed in OTA treated cells.15, 16, 25 Although the duplex structures provide clues about the cellular processing of damaged DNA, analysis of post-replication structures is critical for understanding lesion-induced mutagenicity. In general, 2-base deletion mutations are important factors that may contribute to the mutational profile of C8-dG DNA adducts. For example, experiments on the carcinogenic aromatic amines AAF and aminofluorene (AF), which have extensively been used as models for studying chemical carcinogenesis, have revealed that the associated C8-dG adducts (AAF-dG and AF-dG, respectively) have the propensity to cause 2-base deletions within the NarI sequence context.40-42 In this context, previous studies on the post-replication duplexes of DNA adducted at the C8position of dG by AAF and AF have provided significant biophysical insights into factors that control the onset of deletion mutations.43-49 Specifically, the widely accepted formation mechanism involves lesion-induced stalling of the polymerase and subsequent strand realignment, which leads to a bulge or slipped mutagenic intermediate (SMI) at the lesion site (Figure S1).40 Furthermore, the lesion conformation within a SMI and the ability of a DNA adduct to stabilize the SMI in a postreplication deletion duplex are believed to play critical roles in the formation of deletion mutations.40, 41 The present contribution uses molecular modeling to systematically analyze the possible conformations of 2-base deletion duplexes that may form following replication of OTA-damaged DNA. Although three OTA derived adducts have been previously identified,12 the present work focuses on 2-base deletion duplexes containing the most prevalent OTA–DNA adduct (namely OT-dG, Figure 1a). The 2-base deletion mutations were chosen as a first step since these have been well studied in the literature for DNA


6

Page 7 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


damaged by AA C8-dG adducts,40-42 which allows meaningful comparisons to be made with OT-dG. Since the chemical structure of OT-dG is different from AA adducts in terms of the nucleobase−carcinogen linker (i.e., the C–N–C linker in AA adducts versus the C–C linker in OT-dG) and the ability of OT-dG to adopt multiple ionization states arising from the carboxylic and phenolic functional groups (pKa values of ~ 4.4 and 7.1, respectively;50 Figure 1a), comparison of the newly identified structural characteristics of OTA-adducted 2-base deletion duplexes to those of 2-base deletion duplexes containing other C8-dG adducts can provide unique insight into the effects of adduct chemical composition, including ionization state, on the occurrence of frameshift mutagenesis. This may in turn help develop meaningful hypotheses regarding possible mutagenic outcomes of OTA exposure. In the present contribution, MD simulations and post-processing free energy calculations were carried out with OT-dG reiteratively incorporated into three sequence contexts within a 12-mer DNA strand paired against a 10-mer strand, such that the bulge in the 12-mer contains the adduct and the 5′−base with respect to lesion (Figure 1b). Within each sequence context, four OT-dG ionization states were considered, including three charged states relevant to cellular conditions and the (control) neutral form to monitor the effects of charge. Overall, detailed analysis of lesion site noncovalent (hydrogen–bonding and stacking) interactions and global structural features demonstrate the effect of sequence context and adduct ionization state on the conformations adopted by OTA-adducted 2-base deletion duplexes. MATERIALS AND METHODS MD simulations were carried out on a 12-mer oligonucleotide containing the NarI sequence (5′−G1G2CG3CC) paired against a 10-mer strand, where the adduct was incorporated at G1, G2 or G3 and paired opposite the 2-base deletion site (Figure 1b).


7


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 35

The adduct was considered in the carboxylic ionized (COO–), phenolic ionized (ArO–) and dianionic (COO– and ArO–) OT-dG ionization states, as well as the (control) neutral state (Figure 1a). The initial structure for the stacked conformer of the deletion duplex was built based on an NMR structure of DNA containing the NarI sequence adducted at C8 of syn (χ=0±90°) G3 by aminofluorene (AF), a AA carcinogen (PDB ID: 1AX6).42 The major groove conformer of the OT-dG adducted 2-base deletion duplex was built by adjusting the lesion glycosidic torsion angle to the anti orientation. The corresponding starting structures with the lesion at G1 or G2 were obtained by maintaining the adduct position (i.e., the seventh base from the 5′−end of the damaged 12-mer), modifying the strand sequence, and replacing any resulting A:T terminal pairs with G:C pairs to help prevent duplex unraveling during the MD simulations (Figure 1b). Each duplex strand conformer was energy minimized and heated to 300 K using a 20 ps constant volume simulation. The structure thus obtained was simulated for 40 ns in explicit (TIP3P) water using the AMBER 12 program. Thus, a total of 960 ns of final production simulation (i.e., excluding trial simulations) was analyzed for 24 unique duplexes. The structural analysis, including identifying representative structures, was conducted using the cptraj and ptraj modules of AMBER. Post-processing free energy calculations were performed to evaluate the conformational preferences of the deletion duplexes using the molecular mechanics-Poisson Boltzmann surface area (MM-PBSA) approach.51 Additional details of the simulation protocol, including the selection of initial structures, force field parameters, structural analysis and free energy calculations are provided in the Supporting Information (Section S1). RESULTS Lesion Site Structural Features of OTA-adducted 2-base Deletion DNA Duplex.


8

Page 9 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


The structural features of the OTA-adducted 2-base deletion duplex are in part dictated by the orientation of OT-dG. The glycosidic conformation of the damaged nucleotide is characterized by the χ dihedral angle (∠(O4′–C1′–N9–C4), Figure 1a), which lies on average between 212–218ᵒ (anti) and 37–50ᵒ (syn) for the major groove and stacked conformations, respectively (Table S1). The relative orientation of the OT moiety with respect to the damaged G is characterized by the θ dihedral angle (∠(N9–C8–C10–C11), Figure 1a), which lies on average between 11–18ᵒ and 342–350ᵒ for the major groove and stacked conformations, respectively (Table S1). A similar twist about the nucleobase–OT linkage has been reported for the major groove and stacked conformers of the OT-dG adducted full DNA duplex.39 Thus, there is a preference for slight twist about the nucleobase−OT linkage regardless of the duplex type, conformation, sequence context or adduct ionization state. However, OT-dG is up to 110° less twisted in the deletion or full duplexes39 compared to the isolated OT-dG nucleotide,52 primarily because of constraints imposed by the flanking nucleotides in the DNA environment. In general, the complete OT moiety is solvent exposed in the major groove conformation, while the isocoumarin ring is intercalated within the helix and only the amide-linked phenylalaninic moiety (Figure 1a) is solvent exposed in the minor groove in the stacked conformation (Figures 2 and S2−S4). The guanine moiety of OT-dG remains intrahelical in the major groove conformation, while dG is partially displaced towards the DNA major groove in the stacked conformation (Figures 2 and S2−S4). Although the basic structural features of each deletion duplex are similar, the discrete interactions at the lesion site are affected by the identity of the bases surrounding the lesion and the adduct ionization state, as described in detail in the subsequent sections.

Sequence context most significantly affects the lesion site interactions for the major groove conformation. (i) Effects of sequence on lesion-site hydrogen-bonding interactions: For the


9


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 35

major groove conformation with OT-dG at G1 or G2, the Watson-Crick (WC) hydrogen bonding within the base pairs on the 5′– and 3′–side of the lesion-containing bulge remain intact irrespective of the adduct ionization state (occupancies > 80%; Table S2 and Figures 3, S2 and S3, left). However, when the lesion is at G2, N1 of the damaged G forms additional transient interactions with N2 of the 5′–interstrand G with respect to the bulge (~ 40−50% occupancy; Table S2), and the O6 of the 3′−interstrand G with respect to the bulge (~ 25−30% occupancy; Table S2). In contrast, when the lesion is at G3, the WC hydrogen bonding in the 3′−flanking base pair remains intact (Figure S4, left), while the WC hydrogen bonding in the 5′−base pair either completely or partially disappears (occupancies ~ 0−45%; Table S2). Although disruption in the 5′–base pair distorts the lesion site, this allows the damaged G to WC hydrogen bond with the 5′– interstrand C of the disrupted base pair (occupancies ~ 25−100%; Figures 3 and S5a, and Table S2). This disruption only occurs when the adduct is at G3 since this is the only sequence context with a 5′−interstrand C that can WC hydrogen bond with the OT-dG lesion. In contrast, the 3′−interstrand C within the G1 sequence context does not interact with OT-dG, mainly since its 3′−position hinders accommodation within the bulge formed by OT-dG and its 5′–flanking base.

(ii) Effects of sequence on lesion-site stacking interactions: Similar to the lesion-site hydrogen bonding, lesion-site stacking interactions in the major groove conformation exhibit a marked sequence dependence (Figure 4 and Table S3). For example, when OTdG is at G1, the adduct exhibits maximum stacking with the 3′–intrastrand G (up to −10.4 kcal mol–1), and minimum stacking with the 5′–interstrand A (−0.9 to −1.6 kcal mol–1) regardless of the ionization state. On the other hand, at both G2 and G3, the adduct exhibits minimum stacking with the interstrand bases in both the 5′– and 3′–base pairs.


10

Page 11 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Maximum stacking of the adduct at G2 is observed with the bulged G (up to −6.8 kcal mol–1) since the extrahelical G interacts with the bulky OT moiety in the major groove (Figures 3 and S3, left). In contrast, maximum stacking occurs with the 3′−intrastrand C in the G3 context (up to −6.6 kcal mol–1). Nevertheless, regardless of the sequence context, OT-dG exhibits significant stacking (−4.1 to −6.8 kcal mol–1) with the bulged base present 5′ with respect to the lesion.

(iii) Effect of OTA ionization state on lesion-site interactions: The OTA ionization state has much smaller effects than the sequence context on the lesion-site structural features and therefore discrete lesion-site interactions in the 2-base deletion duplex. Indeed, the adduct ionization state does not significantly affect the lesion-site hydrogen-bonding and stacking interactions when OT-dG is at G2 and G3 (Figure 4, and Tables S2 and S3). However, noticeable differences occur for damage at G1. For example, although no specific hydrogen-bonding interactions occur between the adduct, and the neighboring 5′– and 3′–base pairs for any ionized form of OT-dG at G1, N1 of neutral OT-dG partially interacts with N4 of the 3′–interstrand C (occupancy 22%; Table S2). Similarly, stacking interactions of dianionic OT-dG are significantly weaker (by ~9.6 kcal mol–1) compared to other ionization states at G1, predominantly because greater charge increases repulsive interactions between the bulky moiety and the sugar−phosphate backbone of the flanking bases, which in turn repositions the damaged G in the lesion site such that the stacking interactions are reduced. Thus, although adduct ionization state can alter lesion site interactions in the major groove conformer, this only occurs in select cases and the effects are not as large as discussed for sequence context.

Ionization state and sequence context equally influence the lesion-site interactions for the stacked conformation. (i) Effects on lesion-site hydrogen-bonding interactions: For the


11


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 35

stacked conformation of the 2-base deletion duplex with the adduct at G1 (Figures 5 and S2, right), WC hydrogen bonding in both the 5′– and 3′–neighboring base pairs remains intact for COO– and dianionic OT-dG (occupancy > 90%, Table S2 and Figure S2, right). However, although the 3′−base pair remains intact for neutral OT-dG, the 5′−base pair is disrupted such that the 5′−T is stacked with the bulged C. Additionally, for ArO– OT-dG, the bulged C is intrahelical and stacks with the lesion, which disrupts the 5′−base pair. Nevertheless, irrespective of the adduct ionization state, the adduct does not hydrogen bond with the neighboring 5′– or 3′−base pairs. In contrast to G1, hydrogen bonding occurs between OT-dG at G2 and the neighboring base pairs. Although the WC hydrogen bonds within the 5′− and 3′−base pairs remain mostly intact (occupancy > 80%; Table S2 and Figure S3, right) for the neutral, COO– and dianionic OT-dG duplexes, only the 3′–base pair remains intact for the ArO– duplex. The disruption of the 5′–base pair in the ArO– duplex allows the phenolic oxygen and the lactone group of the OT moiety to hydrogen bond with N1 and N2 of the 5′–interstrand G in the (disrupted) base pair (99 to 37% occupancy; Table S2 and Figure S5b). Furthermore, despite retention of WC hydrogen bonding within the 3′–base pair, transient hydrogen bonds (up to 25% occupancy; Table S2) form between OT-dG and the 3′–interstrand G in deletion duplexes containing neutral OT-dG (i.e., between O6 of OT-dG and N2 of the 3′–interstrand G) and COO– OT-dG (i.e., between N of the peptide linkage of OT-dG and O4′ of the 3′–interstrand G sugar moiety). Although the 5′– and 3′–base pairs are disrupted in at least one OTA ionization state for damage at G1 and G2, WC hydrogen bonding in the 5′– and 3′–base pairs is maintained for damage at G3 regardless of the OTA ionization state (Figure S4, right). Nevertheless, the COO– form of OT-dG uniquely exhibits a transient interaction between


12

Page 13 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


the phenolic group of the OT moiety and O2 of the 5′–interstrand C (24.3% occupancy, Table S2). Thus, overall, both the sequence context and adduct ionization state affect discrete hydrogen-bonding interactions at the lesion site in the stacked conformer, including contacts between the bulky moiety and surrounding nucleobases.

(ii) Effects on lesion-site stacking interactions: Similar to hydrogen-bonding interactions, lesion-site stacking interactions in the stacked conformation are affected by both the sequence and adduct ionization state (Figure 4 and Table S3). At G1, all OTA ionization states exhibit strong stacking with the 3′–intrastrand G (−11.6 to –12.6 kcal mol−1, Table S3). However, due to favorable alignment (Figure S2, right), COO– OT-dG stacks well with the 5′–interstrand A (−7.4 kcal mol−1), while the neutral, ArO– and dianionic adduct forms exhibit weaker lesion-site stacking (~ −3 kcal mol−1; Table S3). Nevertheless, the stacking interactions with other neighboring bases are similar regardless of the adduct ionization state. In the case of damage at G2, the stacking interactions exhibit a marked dependence on the adduct ionization state (Figure 4). For neutral and dianionic OT-dG, the 5′–intrastrand bulged G is intrahelical (Figure S3, right) and strongly stacks with the adduct (−8.7 to −11.3 kcal mol−1; Table S3). Additionally, in both ionization states, the adduct stacks more strongly with the interstrand G in the 3′– than 5′−base pair since intercalation of the bulged G increases the distance between the adduct and the 5′−base pair (Figure S3, right). In contrast, COO– and ArO– OT-dG exhibit negligible (0.0 to −2.5 kcal mol−1) stacking with the (bulged) 5′−G, which remains extrahelical. Although the ArO– lesion exhibits poor stacking (−1.2 to −2.1 kcal mol−1) with the interstrand G of the 5′– and 3′–base pairs, the COO– form exhibits stronger stacking with these bases (−11.1


13


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 35

to −12.1 kcal mol−1), largely due to enhanced intercalation of the OT moiety (Figure S3, right). At the G3 position, the stacked conformer containing ArO– or dianionic OT-dG exhibits decreased lesion site stacking with the 5′–interstrand G compared to neutral and COO– OT-dG (Table S3). This may be because the ArO– and dianionic forms place the isocoumarin ring containing the negatively charged phenolic group inside the DNA helix (Figure S4, right). Additionally, although dianionic OT-dG stacks strongest with the 3′– interstrand G, the other ionization states stack strongest with the 3′–intrastrand C. This is due to more favorable alignment of the π-systems of the OT moiety and the 3′– interstrand G in the deletion duplex containing dianioinic OT-dG compared to other ionization states (Figure S4, right). Overall, the lesion-site structure, and therefore the lesion-site hydrogen-bonding and stacking interactions, for the stacked deletion duplex conformer are more greatly affected by the adduct ionization state compared to the major groove conformer, primarily due to the intrahelical location of the (ionizable) bulky moiety. However, the sequence context can significantly affect the lesion-site non-covalent interactions for both the major groove and stacked conformers, which is mainly due the changes in possible discrete interactions of the lesion with the identity of the surrounding bases. Major Groove and Stacked Conformers of OTA-adducted 2-base Deletion DNA Duplex are Nearly Equally Stable. The previous sections suggest that there are conformation-dependent deviations in discrete local (hydrogen-bonding and stacking) interactions at the lesion site, as well as DNA bending, with sequence context and adduct ionization state. Therefore, it is not surprising that the relative stability of the major groove and stacked conformers also varies with both properties. When OT-dG is at G1, with the exception of the COO– OT-dG


14

Page 15 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


adducted strand, the major groove conformer is more stable than the stacked conformer by ~ 1 – 7 kcal mol–1 (Table 1). For the COO– ionization state, the stacked conformation is ~ 12 kcal mol–1 more stable than the major groove conformation, which may be in part due to stronger stacking between OT-dG and the 5′−interstrand A in the stacked conformer associated with this ionization state (Table S3). Thus, although both the major groove and the stacked conformers lie close in energy for deletion duplexes containing neutral, dianionic or ArO– OT-dG at G1, the major groove conformer may be energetically inaccessible to the COO– duplex. When the adduct is incorporated at G2, the most stable duplex conformation depends on the OTA ionization state. Specifically, the major groove and stacked conformers lie very close in energy (i.e., within ~ 1 kcal mol–1) for the COO– and ArO– adduct forms. In contrast, the stacked conformer is ~ 5–8 kcal mol–1 more stable than the major groove conformer for helices containing neutral or dianionic OT-dG (Table 1). Although, the partial occupancies of the hydrogen bonds in the terminal base pairs may affect the calculated free energies for the 2-base deletion duplexes with the lesion at G2 (see Supporting Information), both conformations may be accessible for the various OTdG ionization states. Finally, when the lesion is at G3, the major groove and stacked conformers fall within ~ 4 kcal mol–1 for the COO– and ArO– adduct, indicating that both conformers may be accessible to adducted DNA (Table 1). In contrast, the stacked conformer is stabilized for neutral OT-dG (by 12.4 kcal mol–1), but destabilized for dianionic OT-dG (by 11.7 kcal mol–1), compared to the major groove conformation. This difference arises in part due to greater lesion site stacking for neutral OT-dG in the stacked conformer compared to the major groove conformer (Table S3) and in part due to a large DNA bend for the stacked conformer containing dianionic OT-dG (Table S4). In summary, the


15


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 35

trends in the relative energies of the major groove and stacked conformers are affected by both the sequence context and the adduct ionization state. However, the effect of adduct ionization state is most pronounced in the G1 and G3 sequence contexts, where one conformer is energetically inaccessible for certain ionization states. DISCUSSION Although previous studies structurally analyzed various conformational themes of full DNA duplexes containing OT-dG to predict the repair propensity of the lesion,39, 52 analysis of post-replication DNA structures is crucial to explain the replication outcomes associated with OTA-adducted DNA. In this context, the present work uses extensive molecular dynamics simulations to provide the first structural insights into the post-replication 2-base deletion duplex containing OT-dG. Our study characterizes two conformations of the deletion duplex (denoted the major groove and stacked conformers) and reveals that the structural features of both conformers are markedly influenced by the identity of the bases surrounding the lesion site. This effect is reflected in both the lesion-site hydrogen-bonding and stacking interactions, as well as the helical bend (see Supporting Information). Furthermore, although OTA ionization state plays a less significant role in dictating the structure of the major groove conformer, the structural features of the stacked conformer are determined by the synergistic effects of sequence and OT-dG ionization state. This difference arises because the extrahelical position of the OT moiety in the major groove conformer precludes significant interactions with the neighboring nucleobases, while the intercalated OT moiety in the stacked conformation permits discrete interactions with the bases surrounding the lesion site. Thus, our simulations demonstrate the significant effect of sequence context and adduct ionization state on the structural features of the OT-dG containing deletion duplex. In turn, these distinct structural features lead to


16

Page 17 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


variations in the relative stability of the deletion duplex conformers with sequence context and ionization state. Nevertheless, for the majority of deletion duplexes considered, the major groove and stacked conformations are likely both energetically accessible. In the subsequent sections, our structural data is compared to that previously collected for other C8-dG adducts, and predictions are made regarding the effects of lesion composition on the frequency of frameshift mutagenesis and OT-dG replication outcomes. Comparison of 2-base Deletion Duplexes Formed by OT-dG and Other C-linked C8dG Adducts within the G3 position of the NarI Sequence.

Effects of nucleobase–carcinogen linker and substituents: To understand the relationship between the chemical composition of the adduct and the induction of deletion mutations, previous studies have structurally characterized 2-base deletion duplexes containing AAF-dG41 or AF-dG42, 53 (Figure 6a) at G3 in the NarI sequence. NMR data revealed that the 2-base deletion duplex containing AF-dG exclusively adopts a stacked conformation in the oligonucleotide sequence considered in the present study.42 However, when the next nearest intrastrand C located 3′ with respect to G3 is replaced by T, the 2-base deletion duplex adopts a mixture of the stacked and major groove conformers.53 Although no sequence dependent conformational preferences have been reported for AAF-dG, circular dichroism and

19F-NMR

reveal that AAF-dG adducted

deletion duplexes adopt a mixture of stacked and major groove conformations within the G3 sequence context.41 The conformational heterogeneity reported for AAF-dG and the sequence dependence of the conformational preference reported for AF-dG parallel our conclusions that more than one conformer of the OT-dG deletion helix may be stabilized and that the preferred conformation depends on sequence context. However, the


17


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 35

relative stability of the competing conformers of the deletion duplex containing OT-dG can also be affected by the adduct ionization state. Furthermore, OT-dG is inherently distinct from these AA adducts in terms of the nucleobase−carcinogen linkage. Specifically, in the stacked conformer, the longer and bent C−N−C linker of the AA adducts only permits the intercalated bulky moiety to stack with the intrastrand bases in the 5′– and 3′−flanking base pairs for AF-dG42 or the 5′−intrastrand base for AAFdG.41 In contrast, the smaller and linear C−C linker of OT-dG allows the OT moiety to stack with each base in the 5′– and 3′−flanking base pairs. As a result, changes in the lesion site sequence may cause greater structural perturbations to the stacked conformer containing OT-dG compared to AF-dG or AAF-dG. Although both AAF-dG and OT-dG deletion duplexes adopt a mixture of major groove and stacked conformers at G3, the lesion site within the stacked conformer is more disrupted for AAF-dG41 compared to OT-dG. Specifically, whereas the WC pairing within the 5′− and 3′−base pairs with respect to the lesion largely remain intact in the stacked conformer for all OT-dG ionization states, the 3′−base pair is disrupted in the case of AAF-dG. This may be due to the position of the AAF-dG acetyl substituent on the nucleobase–carcinogen linker, which exhibits steric repulsion within the helix. In contrast, the phenylalanilic substituent of OT-dG is solvent exposed in the minor groove and therefore causes less distortion (Figure S4, right). Furthermore, discrete interactions between substituents on the OT bulky moiety and the surrounding nucleobases can stabilize the stacked deletion duplex conformer. On the other hand, in the major groove conformer, OT-dG hydrogen bonds with the 5′−interstrand C (Figures S4, left and S5a), while the position of the acetyl substituent of AAF-dG repositions the damaged base and thereby prevents such hydrogen bonding with the surrounding


18

Page 19 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


bases.41 As a result, the major groove deletion duplex conformation is more distorted when containing OT-dG than AAF-dG at G3. Hence, deviations in the structural features of both the major groove (AAF-dG and OT-dG) and the stacked conformers (AF-dG, AAFdG and OT-dG) for different C-linked and N-linked C8-dG adducts suggest that the linker type and the location of substituents within the bulky moiety may dictate the structural features, and therefore the stability, of 2-base deletion duplexes.

Effect of bulky moiety ring size and directionality of ring extension: As mentioned in the Introduction, unlike the AA adducts, OT-dG belongs to a relatively less studied class of adducts that possess a C−C linkage at the guanine–carcinogen junction. Five model Clinked C8-dG adducts (Figure 6b) have been previously studied in 2-base deletion duplexes at the G3 site of the NarI sequence using both experimental and computational techniques.34, 54 Similar to OT-dG, the major groove and stacked conformations of 2-base deletion duplexes containing all five model adducts are close in energy.34, 54 However, deletion duplexes containing a single−ringed system (such as Fur-dG and Ph-dG), as well as a double ring system with a linear extension of the C8-moiety (such as BTh-dG), prefer the major groove conformation (by ~ 1.1−3.6 kcal mol−1),34, 54 which is stabilized by hydrogen bonding between the damaged G and the 5′−interstrand C with respect to the bulge. Although similar hydrogen-bonding interactions are also observed in the major groove conformation of deletion duplexes containing Q-dG34 or Py-dG,54 favorable alignment of the bicyclic ring system of Q-dG and the larger size of the Py-dG bulky moiety allow these adducts to preferentially stabilize the stacked conformer (by ~ 3 to 6 kcal mol−1).54 For adduct formation at G3, the stacked conformer of deletion duplexes containing neutral or COO− OT-dG is more stable than the major groove conformer (Table 1). Therefore, neutral or COO− OT-dG possesses a conformational preference


19


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 35

similar to Q-dG and Py-dG.34, 54 This comparable conformational preference likely occurs because the alignment of the neutral isocoumarin moiety of OT-dG relative to the damaged dG is similar to the bulky moieties of Q-dG and Py-dG (i.e., C8-dG substituent extension is perpendicular to dG; Figures 1a and 6b). On the other hand, the Fur-dG and Ph-dG (i.e, single-ringed adducts), and BTh-dG (i.e, linear extension of the C8 substituent relative to the damaged G or extension in a direction perpendicular to the isocoumarin moiety of OT-dG; Figure 6b) lead to weaker stacking between the bulky moiety and the surrounding bases, which results in a less stable stacked adducted deletion duplex conformation. However, we note that the dianionic and ArO− forms of OT-dG at G3 destabilize the stacked conformer (Table 1), probably due to enhanced repulsion when the negatively charged isocoumarin moiety adopts an intrahelical position. Therefore, the adduct ionization state also plays a role in dictating the preferred conformation of OT-dG adducted deletion duplexes. Regardless, comparison of the previously studied Clinked C8-dG adducts and OT-dG further highlights that the size and extension direction of the C8-dG substituent have significant impact on the conformational preferences of damaged DNA, including deletion duplexes. Implications to Mutagenesis. The mutagenic outcome of bulky DNA lesions is the consequence of the replication of damaged DNA by a DNA polymerase. High-fidelity (replicative) polymerases follow an induced-fit mechanism to ensure proper WC base pairing between the incoming dNTP and the nucleotide on the template strand. Due to the high selectivity of the tight active site, these polymerases typically do not replicate bulky DNA lesions, which instead stall DNA replication. One way to overcome stalling is through recruitment of translesion synthesis (TLS) polymerases, which possess more flexible active sites and thus can replicate past bulky DNA lesions, albeit leading to error prone replication.55 As


20

Page 21 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


discussed in the Introduction, many bulky lesions have been shown to trigger base slippage and stabilize a SMI, which is the accepted precursor for a frameshift mutation.34, 40 For damage at G3 in the NarI sequence, the prevalent mechanism for 2-base deletion mutations induced by AA C8-dG adducts involves initial insertion of C opposite the lesion, followed by a 2-base slippage, which forms a bulge in the template strand (Figure S1).40 A syn lesion conformation allows the C8-moiety of the AA adducts to intercalate within the bulge and stabilize the SMI.41, 56-58 The polymerase then further extends the SMI to produce the 2-base deletion product. Along similar lines, combined experimental and computational studies of the replication of model C-linked C8-dG adducts (i.e., Fur-dG, Ph-dG, Q-dG, BTh-dG and Py-dG, Figure 6b) at G3 proposed replication past these lesions proceeds via a slippage mechanism (Figure S6).34,

54

Specifically, since these adducts prefer a syn conformation in the isolated nucleoside, they have been proposed to adopt the syn conformation at the single-strand–doublestrand junction during TLS.34, 40 However, each adduct is proposed to then flip to a less favored anti conformation to pair with the incoming dCTP, which allows the damaged base and its 5′–base to slip to form a two-base bulge (Figure S6). However, since the

anti conformation of the smallest lesions (i.e., Fur-dG, Ph-dG and BTh-dG) is energetically preferred within the 2-base bulged DNA, the SMI is not significantly stabilized, and can re-align to form a C:C mismatch. In contrast, Q-dG and Py-dG are stabilized in the syn conformation within the 2-base bulge due to the direction of the ring expansion and enhanced stacking arising from the larger bulky moieties, and therefore both adducts stabilize 2-base deletion mutations.34, 54 Similar to the model C-linked adducts, our previous quantum mechanical study revealed OT-dG intrinsically prefers the syn conformation, and has a small anti/syn


21


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 35

inter-conversion barrier at the nucleotide level.52 Thus, OT-dG may undergo a syn to

anti conformational switch at the single-strand–double-strand junction to permit WC hydrogen bonding with an incoming dCTP. However, with the exception of dianionic OT-dG, the present work reveals that the stacked conformation of the OT-dG adducted 2-base deletion duplexes at G3 is either strongly preferred or both the major groove and stacked conformations are energetically accessible. The accessibility of the stacked conformation in at least three OT-dG ionization states suggests that the lesion will likely lead to 2-base deletion mutations upon replication. This proposal parallels the observed behavior of the Q-dG34 and Py-dG54 C-linked adducts, which contain similar extension of the bulky moiety in a perpendicular direction with respect to the damaged dG. Interestingly, our predicted sequence and ionization state dependent conformational outcomes of OT-dG adducted 2-base deletion duplexes correlates with the complicated mutagenic profile observed upon OTA exposure.15, 16 For example, an in

vivo study of the site-specific exposure of OTA to the kidneys of gpt delta rats reveals that only the DNA of cells derived from the renal outer medulla is affected by OTA.15 Although a number of factors may lead to the differential expression of OTA-induced mutagenicity, it is possible that the stabilization of alternate conformations of 2-base deletion duplexes due to changes in the cellular environment may contribute to this effect. For example, perhaps the variation in the pKa of the ionizable (−COOH and −OH) groups in the OT moiety with local cellular environments, such as when the damaged DNA binds to proteins,59 may dictate the OT-dG ionization state. The present work illustrates that this shift in ionization state may in turn affect the relative stability of the stacked conformer, and therefore the propensity of OT-dG to cause deletion mutations. In summary, the present work uses extensive molecular dynamics simulations on 2-base deletion duplexes to reveal conformational-dependent deviations in discrete


22

Page 23 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


local (hydrogen-bonding and stacking) interactions at the lesion site, as well as DNA bending, with OT-dG sequence context and ionization state, which in turn affects the stability of the slipped mutagenic intermediate associated with the formation of deletion mutations. Although our present work predicts the conformational outcomes and stability of 2-base deletion duplexes containing OT-dG adduct, further experimental input is required to verify the key hypotheses put forward in the present work regarding OTA-mediated mutations. Indeed, there is a current lack of explicit biochemical data on the replication of OT-dG adducted DNA by a DNA polymerase. Furthermore, other adducts associated with OTA21, 22 and different types of frameshift mutations15, 16 may also play key roles in the harmful effects of OTA exposure. Thus, future studies are needed to verify the role of all key players, including 2-base deletions, in OTA induced mutagenesis. ASSOCIATED CONTENT SUPPORTING INFORMATION

Additional details of the computational protocol; Global Structural Features of OTAadducted 2-base Deletion DNA Duplex; proposed 2-base slippage mechanism for the formation of deletion mutations arising from aromatic amines C8-dG adducts; Lesion site structures of DNA deletion duplexes containing OT-dG at G1, G2 or G3; interstrand hydrogen bonds in deletion duplexes involving dianionic OT-dG at G3 or ArO– OT-dG at G2; previously proposed mechanisms for mutations depicting the fate of the SMI formed by model C-linked C8-dG adducts; average and standard deviations in the χ and θ dihedral angles for the adduct over the last 10 ns of the production simulation; hydrogen-bond occupancies at the lesion site for the last 10 ns of the production simulation; average and standard deviation in the van der Waals stacking energy of OTdG and the flanking base pairs; average and standard deviation in the bending angle for


23


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 35

each OT-dG adducted DNA deletion duplex, hydrogen-bond occupancies for the terminal G:C pairs over the last 10 ns of the production simulation; average and standard deviation of the rmsd over the last 10 ns and full 40 ns of the production simulations. The Supporting Information is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. Telephone: (403) 329-2323. Fax: (403) 329-2057. ORCID Stacey D. Wetmore: 0000-0002-5801-3942 Purshotam Sharma: 0000-0002-5164-9833 Funding SDW thanks the Canada Foundation for Innovation (CFI) [22770], the Natural Sciences and Engineering Research Council (NSERC) of Canada [249598-07]; and the Canada Research Chair (CRC) program [950-228175] for supporting this research, RAM thanks the Natural Sciences and Engineering Research Council (NSERC) of Canada [3116002013]; PS thanks the Department of Science and Technology (DST), and University

Grants Commission (UGC), New Delhi, for financial support through the DST INSPIRE (IFA14-CH162) and the UGC FRP (F.4-5(176-FRP/2015(BSR)) programs, respectively. Notes The authors declare no competing financial interest. ACKNOWLEDGMENTS Calculations were conducted on the New Upscale Cluster for Lethbridge to Enable Innovative Chemistry (NUCLEIC), as well as additional resources provided by WestGrid and Compute/Calcul Canada.


24

Page 25 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


ABBREVIATIONS A, adenine; AAF, acetylaminoflourene; AF, aminoflourene; BTh, benzothiophene; C, cytosine; dCTP, deoxycytidine triphosphate; dG, 2′-deoxyguanosine; Fur, furan; G, guanine; AAs, aromatic amines; IARC, International Agency of Research on Cancer; MD, molecular dynamics; MM-PBSA, molecular-mechanics-Poisson-Boltzmann surface area; MoA, mechanism of action; OTA, ochratoxin A; Ph, phenyl; Py, pyrene; Q, quinolone; SMI, slipped mutagenic intermediate; TLS, translesion synthesis; T, thymine; WC, Watson Crick. REFERENCES (1)

(2)

(3) (4) (5) (6) (7)

(8)

(9) (10)

(11) (12)

Bayman, P., Baker, J. L., Doster, M. A., Michailides, T. J., and Mahoney, N. E. (2002) Ochratoxin production by the aspergillus ochraceus group and aspergillus alliaceus. Appl. Environ. Microbiol. 68, 2326-2329. Abarca, M. L., Bragulat, M. R., Castellá, G., and Cabañes, F. J. (1994) Ochratoxin A production by strains of aspergillus niger var. niger. Appl. Environ. Microbiol. 60, 26502652. Cabañes, F. J., Bragulat, M. R., and Castellá, G. (2010) Ochratoxin A producing species in the genus Penicillium. Toxins 2, 1111-1120. Engelhardt, G., Barthel, J., and Sparrer, D. (2006) Fusarium mycotoxins and ochratoxin A in cereals and cereal products. Mol. Nutr. Food Res. 50, 401-405. Jörgensen, K. (1998) Survey of pork, poultry, coffee, beer and pulses for ochratoxin A. Food Addit. Contam. 15, 550-554. Mally, A., and Dekant, W. (2009) Mycotoxins and the kidney: modes of action for renal tumor formation by ochratoxin A in rodents. Mol. Nutr. Food Res. 53, 467-478. Gazinska, P., Herman, D., Gillett, C., Pinder, S., and Mantle, P. (2012) Comparative immunohistochemical analysis of ochratoxin A tumourigenesis in rats and urinary tract carcinoma in humans; mechanistic significance of p-S6 ribosomal protein expression. Toxins 4, 643-662. Liu, J., Wu, S., Shen, H., Cui, J., Wang, Y., Xing, L., Wang, J., Yan, X., and Zhang, X. (2015) Ochratoxin A induces DNA damage and G2 phase arrest in human esophageal epithelium Het-1A cells in vitro. J. Toxicol. Sci. 40, 657-665. Schwartz, G. G. (2002) Hypothesis: does ochratoxin A cause testicular cancer? Cancer Causes Control 13, 91-100. Kujawa, M. (1994) Some naturally occurring substances: Food items and constituents, heterocyclic aromatic amines and mycotoxins. IARC monographs on the evaluation of carcinogenic risks to humans, Vol. 56. Herausgegeben von der International Agency for Research on Cancer, World Health Organization. 599 Seiten, zahlr. Abb. und Tab. World Health Organization. Geneva 1993. Preis: 95,—Sw. fr; 95, 50 US$. Food/Nahrung 38, 351-351. Kőszegi, T., and Poór, M. (2016) Ochratoxin A: Molecular interactions, mechanisms of toxicity and prevention at the molecular Level. Toxins 8, 111. Pfohl-Leszkowicz, A., and Manderville, R. A. (2011) An update on direct genotoxicity as a molecular mechanism of ochratoxin a carcinogenicity. Chem. Res. Toxicol. 25, 252-262.


25


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(13) (14)

(15)

(16)

(17)

(18)

(19) (20)

(21)

(22)

(23)

(24)

(25)

(26)

(27)

(28)

(29) (30)

Page 26 of 35

Malir, F., Ostry, V., Pfohl-Leszkowicz, A., Malir, J., and Toman, J. (2016) Ochratoxin A: 50 Years of research. Toxins 8, 191. Kuiper-Goodman, T., Hilts, C., Billiard, S. M., Kiparissis, Y., Richard, I. D. K., and Hayward, S. (2010) Health risk assessment of ochratoxin A for all age-sex strata in a market economy. Food Addit. Contam. Part A, Chemistry, Analysis, Control, Exposure & Risk Assessment 27, 212-240. Hibi, D., Suzuki, Y., Ishii, Y., Jin, M., Watanabe, M., Sugita-Konishi, Y., Yanai, T., Nohmi, T., Nishikawa, A., and Umemura, T. (2011) Site-specific in vivo mutagenicity in the kidney of gpt delta rats given a carcinogenic dose of ochratoxin A. Toxicol. Sci., kfr139. Kuroda, K., Hibi, D., Ishii, Y., Takasu, S., Kijima, A., Matsushita, K., Masumura, K.-i., Watanabe, M., Sugita-Konishi, Y., and Sakai, H. (2013) Ochratoxin A induces DNA doublestrand breaks and large deletion mutations in the carcinogenic target site of gpt delta rats. Mutagenesis, get054. Faucet, V., Pfohl-Leszkowicz, A., Dai, J., Castegnaro, M., and Manderville, R. A. (2004) Evidence for covalent DNA adduction by Ochratoxin A following chronic exposure to rat and subacute exposure to pig. Chem. Res. Toxicol. 17, 1289-1296. Obrecht-Pflumio, S., and Dirheimer, G. (2001) Horseradish peroxidase mediates DNA and deoxyguanosine 3'-monophosphate adduct formation in the presence of ochratoxin A. Arch. Toxicol. 75, 583-590. Obrecht-Pflumio, S., and Dirheimer, G. (2000) In vitro DNA and dGMP adducts formation caused by ochratoxin A. Chem. Biol. Interact. 127, 29-44. Dai, J., Wright, M. W., and Manderville, R. A. (2003) Ochratoxin A forms a carbon-bonded C8-deoxyguanosine nucleoside adduct: implications for C8 reactivity by a phenolic radical. J. Am. Chem. Soc. 125, 3716-3717. Dai, J., Wright, M. W., and Manderville, R. A. (2003) An oxygen-bonded C8deoxyguanosine nucleoside adduct of pentachlorophenol by peroxidase activation: evidence for ambident c8 reactivity by phenoxyl radicals. Chem. Res. Toxicol. 16, 817821. Tozlovanu, M., Faucet-Marquis, V., Pfohl-Leszkowicz, A., and Manderville, R. A. (2006) Genotoxicity of the hydroquinone metabolite of ochratoxin A: structure-activity relationships for covalent DNA adduction. Chem. Res. Toxicol. 19, 1241-1247. Manderville, R. A., and Wetmore, S. D. (2016) Understanding the mutagenicity of Olinked and C-linked guanine DNA Adducts: A combined experimental and computational approach. Chem. Res. Toxicol. 30, 177-188. Mantle, P. G., Faucet-Marquis, V., Manderville, R. A., Squillaci, B., and Pfohl-Leszkowicz, A. (2010) Structures of covalent adducts between DNA and ochratoxin A: A new factor in debate about genotoxicity and human risk assessment. Chem. Res. Toxicol. 23, 89-98. Akman, S. A., Adams, M., Case, D., Park, G., and Manderville, R. A. (2012) Mutagenicity of ochratoxin A and its hydroquinone metabolite in the SupF gene of the mutation reporter plasmid Ps189. Toxins 4, 267-280. Kozack, R., Seo, K.-Y., Jelinsky, S. A., and Loechler, E. L. (2000) Toward an understanding of the role of DNA adduct conformation in defining mutagenic mechanism based on studies of the major adduct (formed at N2-dG) of the potent environmental carcinogen, benzo [a] pyrene. Mutat. Res. Fund. Mol. Mech. Mut. 450, 41-59. Patel, D. J., Mao, B., Gu, Z., Hingerty, B. E., Gorin, A., Basu, A. K., and Broyde, S. (1998) Nuclear magnetic resonance solution structures of covalent aromatic amine-DNA adducts and their mutagenic relevance. Chem. Res. Toxicol. 11, 391-407. Millen, A. L., Sharma, P., and Wetmore, S. D. (2012) C8-linked bulky guanosine DNA adducts: experimental and computational insights into adduct conformational preferences and resulting mutagenicity. Future Med. Chem. 4, 1981-2007. Cho, B. P. (2004) Dynamic conformational heterogeneities of carcinogen-DNA adducts and their mutagenic relevance. J. Environ. Sci. Health C 22, 57-90. Kathuria, P., Sharma, P., Abendong, M. N., and Wetmore, S. D. (2015) Conformational preferences of DNA following damage by aristolochic acids: structural and energetic


26

Page 27 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38)

(39)

(40)

(41)

(42)

(43)

(44)

Insights into the different mutagenic potential of the ALI and ALII-N6-dA Adducts. Biochemistry 54, 2414-2428. Kathuria, P., Sharma, P., and Wetmore, S. D. (2015) Adenine versus guanine DNA adducts of aristolochic acids: role of the carcinogen–purine linkage in the differential global genomic repair propensity. Nucleic Acids Res. 43, 7388-7397. Kathuria, P., Sharma, P., and Wetmore, S. D. (2016) Effect of base sequence context on the conformational heterogeneity of aristolactam-I adducted DNA: structural and energetic insights into sequence-dependent repair and mutagenicity. Toxicol. Res. 5, 197209. Kuska, M. S., Witham, A. A., Sproviero, M., Manderville, R. A., Majdi Yazdi, M., Sharma, P., and Wetmore, S. D. (2013) Structural influence of C8-phenoxy-guanine in the NarI recognition DNA sequence. Chem. Res. Toxicol. 26, 1397-1408. 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., and Wetmore, S. D. (2014) Structural and biochemical impact of C8-aryl-guanine adducts within the NarI recognition DNA sequence: influence of aryl ring size on targeted and semi-targeted mutagenicity. Nucleic Acids Res. 42, 13405-13421. Witham, A. A., Verwey, A. M. R., Sproviero, M., Manderville, R. A., Sharma, P., and Wetmore, S. D. (2015) Chlorine functionalization of a model phenolic C8-guanine adduct increases conformational rigidity and Blocks Extension by a Y-Family DNA Polymerase. Chem. Res. Toxicol. 28, 1346-1356. Ding, S., Kropachev, K., Cai, Y., Kolbanovskiy, M., Durandina, S. A., Liu, Z., Shafirovich, V., Broyde, S., and Geacintov, N. E. (2012) Structural, energetic and dynamic properties of guanine (C8)–thymine (N3) cross-links in DNA provide insights on susceptibility to nucleotide excision repair. Nucleic Acids Res. 40, 2506-2517. Mu, H., Kropachev, K., Chen, Y., Zhang, H., Cai, Y., Geacintov, N. E., and Broyde, S. (2013) Role of structural and energetic factors in regulating repair of a bulky DNA lesion with different opposite partner bases. Biochemistry 52, 5517-5521. Fuchs, R. P. P., Schwartz, N., and Daune, M. P. (1981) Hot spots of frameshift mutations induced by the ultimate carcinogen N-acetoxy-N-2-acetylaminofluorene. Nature 294, 657-659. Sharma, P., Manderville, R. A., and Wetmore, S. D. (2014) 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. 42, 1183111845. Roy, D., Hingerty, B. E., Shapiro, R., and Broyde, S. (1998) A slipped replication intermediate model is stabilized by the syn orientation of N-2-aminofluorene- and N-2(acetylaminofluorene-modified guanine at a mutational hotspot. Chem. Res. Toxicol. 11, 1301-1311. Sandineni, A., Lin, B., MacKerell Jr, A. D., and Cho, B. P. (2013) Structure and thermodynamic insights on acetylaminofluorene-modified deletion DNA duplexes as models for frameshift mutagenesis. Chem. Res. Toxicol. 26, 937-951. Mao, B., Gorin, A., Gu, Z., Hingerty, B. E., Broyde, S., and Patel, D. J. (1997) Solution structure of the Aminofluorene-Intercalated Conformer of the syn [AF]-C8-dG Adduct Opposite a− 2 Deletion Site in the NarI Hot Spot Sequence Context. Biochemistry 36, 14479-14490. Bichara, M., and Fuchs, R. P. (1985) DNA binding and mutation spectra of the carcinogen N-2-aminofluorene in Escherichia coli: A correlation between the conformation of the premutagenic lesion and the mutation specificity. J. Mol. Biol. 183, 341-351. Ames, B. N., Gurney, E. G., Miller, J. A., and Bartsch, H. (1972) Carcinogens as frameshift mutagens: Metabolites and derivatives of 2-Acetylaminofluorene and other aromatic amine carcinogens. Proc. Natl. Acad. Sci. 69, 3128-3132.


27


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

(45)

(46) (47)

(48) (49)

(50)

(51)

(52)

(53)

(54)

(55) (56)

(57)

(58)

(59)

Page 28 of 35

Masumura, K.-i., Matsui, K., Yamada, M., Horiguchi, M., Ishida, K., Watanabe, M., Wakabayashi, K., and Nohmi, T. (2000) Characterization of mutations induced by 2amino-1-methyl-6-phenylimidazo [4, 5-b] pyridine in the colon of gpt delta transgenic mouse: novel G: C deletions beside runs of identical bases. Carcinogenesis 21, 20492056. Shibutani, S., and Grollman, A. P. (1993) On the mechanism of frameshift (deletion) mutagenesis in vitro. J. Biol. Chem. 268, 11703-11710. Gupta, P. K., Johnson, D. L., Reid, T. M., Lee, M. S., Romano, L. J., and King, C. M. (1989) Mutagenesis by single site-specific arylamine-DNA adducts. Induction of mutations at multiple sites. J. Biol. Chem. 264, 20120-20130. Schorr, S., and Carell, T. (2010) Mechanism of acetylaminofluorene-dG induced frameshifting by Polymerase η. ChemBioChem 11, 2534-2537. Burnouf, D., Koehl, P., and Fuchs, R. (1989) Single adduct mutagenesis: strong effect of the position of a single acetylaminofluorene adduct within a mutation hot spot. Proc. Natl. Acad. Sci. 86, 4147-4151. Chiodini, A. M., Scherpenisse, P., and Bergwerff, A. A. (2006) Ochratoxin A contents in wine: comparison of organically and conventionally produced products. J. Agric. Food Chem. 54, 7399-7404. Miller III, B. R., McGee Jr, T. D., Swails, J. M., Homeyer, N., Gohlke, H., and Roitberg, A. E. (2012) MMPBSA. py: an efficient program for end-state free energy calculations. J. Chem. Theory Comput. 8, 3314-3321. Sharma, P., Manderville, R. A., and Wetmore, S. D. (2013) Modeling the conformational preference of the carbon-bonded covalent adduct formed upon exposure of 2′deoxyguanosine to ochratoxin A. Chem. Res. Toxicol. 26, 803-816. Jain, N., Li, Y., Zhang, L., Meneni, S. R., and Cho, B. P. (2007) Probing the sequence effects on Nar I-induced−2 frameshift mutagenesis by dynamic 19F NMR, UV, and CD spectroscopy. Biochemistry 46, 13310-13321. Sproviero, M., Verwey, A. M. R., Witham, A. A., Manderville, R. A., Sharma, P., and Wetmore, S. D. (2015) Enhancing bulge stabilization through linear extension of C8-arylguanine adducts to promote polymerase blockage or strand realignment to produce a C:C mismatch. Chem. Res. Toxicol. 28, 1647-1658. Stallons, L. J., and McGregor, W. G. (2010) Translesion synthesis polymerases in the prevention and promotion of carcinogenesis. J. Nucleic Acids 2010, 643857. Jain, N., Li, Y., Zhang, L., Meneni, S. R., and Cho, B. P. (2007) Probing the sequence effects on NarI-Induced -2 frameshift mutagenesis by dynamic 19F NMR, UV, and CD spectroscopy. Biochemistry 46, 13310-13321. Xu, L., and Cho, B. P. (2016) Conformational insights into the mechanism of acetylaminofluorene-dG-induced frameshift mutations in the NarI mutational hotspot. Chem. Res. Toxicol. 29, 213-226. Milhé, C., Fuchs, R. P. P., and Lefèvre, J.-F. (1996) NMR data show that the carcinogen N2-acetylaminofluorene stabilises an intermediate of −2 frameshift mutagenesis in a region of high mutation frequency. Eur. J. Biochem. 235, 120-127. Il'ichev, Y. V., Perry, J. L., and Simon, J. D. (2002) Interaction of ochratoxin A with human serum albumin. Preferential binding of the dianion and pH Effects. J. Phys. Chem. B 106, 452-459.


28

Page 29 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Table 1: Relative free energies (kcal mol–1) of the major groove and stacked conformers of the 2-base deletion duplex containing OT-dG in different ionization states at G1, G2 or G3. sequence

ionization state

G1

neutral COO− ArO− dianionic neutral COO− ArO− dianionic neutral COO− ArO− dianionic

G2

G3

conformer major groove stacked 0.0 6.6 11.9 0.0 0.0 3.2 0.0 0.8 5.2 0.0 0.0 1.0 0.0 1.1 8.1 0.0 12.4 0.0 4.3 0.0 0.0 3.6 0.0 11.7


29


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 30 of 35

Figure 1. (a) Chemical structure of the OT-dG adduct indicating the constituent structural moieties and atomic numbering required to define important dihedral angles. Wavy bonds represent the 3′ and 5′–sites where the adduct is covalently linked within a DNA strand. The OT-dG adduct was considered in the neutral (non-ionized form; R1 = OH, R2 = OH), carboxylic group ionized (COO–; R1 = O–, R2 = OH), phenolic group ionized (ArO–; R1 = OH, R2 = O–) and dianionic (R1 = R2 = O–) forms. (b) The oligonucleotide sequence used to simulate the deletion duplex containing OT-dG in the G1, G2 or G3 sequence context, with the lesion position highlighted in red.


30

Page 31 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 2. Representative structures highlighting the lesion site for the two conformations of the OT-dG adducted DNA 2-base deletion duplex considered in the present work. Examples are provided for the COO− ionization state of OT-dG at G3. The OT moiety is shown in red, the damaged G moiety in blue, the bulged base in orange, and the base pairs flanking the lesion in green.


31


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 32 of 35

Figure 3. Lesion site representative structures of the OT-dG adducted deletion duplexes obtained from MD simulations highlighting the effect of sequence context on the hydrogen-bonding pattern in the major groove conformer. Examples are provided for the COO– form of OT-dG at G1, G2 or G3. The OT moiety is shown in red, the damaged G moiety in blue, the bulged base in orange, and the base pairs flanking the lesion in green. Disrupted hydrogen bonds are encircled in red.


32

Page 33 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 4. Variation in the van der Waals interaction energy (kcal mol–1) for the (a) major groove and (b) stacked conformers of deletion duplexes with OT-dG at the G1 (red), G2 (blue) or G3 (green) position.


33


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 34 of 35

Figure 5. Lesion site representative structures of the OT-dG adducted deletion duplexes obtained from MD simulations highlighting the effect of adduct ionization state on the hydrogen-bonding pattern in the stacked conformer. Examples are provided for the OTdG lesion at the G1 position in the NarI recognition sequence. The OT moiety is shown in red, the damaged G in blue, the bulged base in orange, and the base pairs flanking the lesion in green. Disrupted hydrogen bonds are encircled in red.


34

Page 35 of 35

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 6. (a) N-linked and (b) model C-linked C8-dG adducts previously studied in 2base deletion duplexes.


35

Molecular Modeling of the Major DNA Adduct Formed from Food

Recommend Documents