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. 2023 Nov 21;36(12):1947–1960. doi: 10.1021/acs.chemrestox.3c00226

Base-Displaced Intercalated Structure of the 3-(2-Deoxy-β-D-erythropentofuranosyl)-pyrimido[1,2-f]purine-6,10(3H,5H)-dione (6-oxo-M1dG) Lesion in DNA

Yizhi Fu 1, Plamen P Christov 1, Philip J Kingsley 1, Robyn M Richie-Jannetta 1, Lawrence J Marnett 1, Michael P Stone 1,*
PMCID: PMC10731638  PMID: 37989274

Abstract

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The genotoxic 3-(2-deoxy-β-D-erythro-pentofuranosyl)pyrimido[1,2-α]purin-10(3H)-one (M1dG) DNA lesion arises from endogenous exposures to base propenals generated by oxidative damage and from exposures to malondialdehyde (MDA), produced by lipid peroxidation. Once formed, M1dG may oxidize, in vivo, to 3-(2-deoxy-β-D-erythropentofuranosyl)-pyrimido[1,2-f]purine-6,10(3H,5H)-dione (6-oxo-M1dG). The latter blocks DNA replication and is a substrate for error-prone mutagenic bypass by the Y-family DNA polymerase hpol η. To examine structural consequences of 6-oxo-M1dG damage in DNA, we conducted NMR studies of 6-oxo-M1dG incorporated site-specifically into 5′ -d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG). NMR spectra afforded detailed resonance assignments. Chemical shift analyses revealed that nucleobase C21, complementary to 6-oxo-M1dG, was deshielded compared with the unmodified duplex. Sequential NOEs between 6-oxo-M1dG and A5 were disrupted, as well as NOEs between T20 and C21 in the complementary strand. The structure of the 6-oxo-M1dG modified DNA duplex was refined by using molecular dynamics (rMD) calculations restrained by NOE data. It revealed that 6-oxo-M1dG intercalated into the duplex and remained in the anti-conformation about the glycosyl bond. The complementary cytosine C21 extruded into the major groove, accommodating the intercalated 6-oxo-M1dG. The 6-oxo-M1dG H7 and H8 protons faced toward the major groove, while the 6-oxo-M1dG imidazole proton H2 faced into the major groove. Structural perturbations to dsDNA were limited to the 6-oxo-M1dG damaged base pair and the flanking T3:A22 and A5:T20 base pairs. Both neighboring base pairs remained within the Watson–Crick hydrogen bonding contact. The 6-oxo-M1dG did not stack well with the 5′-neighboring base pair T3:A22 but showed improved stacking with the 3′-neighboring base pair A5:T20. Overall, the base-displaced intercalated structure was consistent with thermal destabilization of the 6-oxo-M1dG damaged DNA duplex; thermal melting temperature data showed a 15 °C decrease in Tm compared to the unmodified duplex. The structural consequences of 6-oxo-M1dG formation in DNA are evaluated in the context of the chemical biology of this lesion.

Introduction

The M1dG [3-(2′-deoxy-α-D-erythro-pentofuranosyl)pyrimido[1,2-a]-purin-10(3H)-one] lesion arises as a consequence of oxidative damage to DNA (Chart 1), from formation of base propenals that may transfer the oxopropenyl moiety to dG.1−3 (Scheme 1). M1dG also forms in DNA following endogenous cellular exposure to malondialdehyde (MDA), a toxic metabolite produced by lipid peroxidation and prostaglandin biosynthesis.4−6 The latter exists in solution primarily as β-hydroxyacrolein and reacts with dG as a bis-electrophile to form M1dG (Scheme 1).7,8 The chemical and structural biology of M1dG is of interest9−12 since it is the most abundant exocyclic DNA lesion present endogenously in human DNA.13−17 Significantly, M1dG undergoes further chemistry in DNA (Scheme 1). When placed opposite dC, it rearranges to N2-oxopropenyl-dG (OpdG).18 It also may be oxidized to 3-(2-deoxy-β-D-erythropentofuranosyl)-pyrimido[1,2-f]purine-6,10(3H,5H)-dione (6-oxo-M1dG).19−21 The Y-family polymerase hPol η preferentially inserts dAMP or dGMP across from 6-oxo-M1dG, suggesting that like M1dG,22,23 6-oxo-M1dG is genotoxic. Crystal structures of hPol η forming both insertion and extension complexes in which template-primer oligodeoxynucleotide duplexes contained a site-specific 6-oxo-M1dG lesion suggested that 6-oxo-M1dG blocks DNA replication and is mutagenic.24

Chart 1. A. Structure of 6-oxo-M1dG;a B. Oligodeoxynucleotide Used in This Work, Showing Numbering of Individual Nucleotides.

Chart 1

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a Note the atom numbering of 6-oxo-M1dG differs from the conventional atom numbering of dG.

Scheme 1. Formation of 6-oxo-M1dG in DNA.

Scheme 1

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Endogenous cellular exposures to base propenals arising due to DNA oxidation (shown is dA-Propenal) or malondialdehyde (MDA), a lipid peroxidation product, produce M1dG lesions, which exist in equilibrium with OpdG. When M1dG is opposite dC in dsDNA it exists primarily as OpdG. PdG has been employed as a stable structural analog of M1dG. M1dG may also undergo further oxidation to form 6-oxo-M1dG.

Here, we inserted 6-oxo-M1dG site-specifically into 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) (Chart 1).21 NMR analyses afforded detailed resonance assignments, and the structure of the 6-oxo-M1dG modified DNA duplex was refined by using molecular dynamics (rMD) calculations restrained by NOE data. This revealed that 6-oxo-M1dG intercalated into the duplex and remained in the anti-conformation about the glycosyl bond, while the complementary nucleotide C21 extruded from the helix, forming a base-displaced intercalation structure. While the presence of 6-oxo-M1dG thermally destabilized the DNA, the lesion formed stronger stacking interactions with the 3′-neighbor A5:T20 base pair than with the 5′-neighbor T3:A22 base pair. The results are evaluated in the context of the chemical biology of the 6-oxo-M1dG lesion.

Materials and Methods

Sample Preparation

The Vanderbilt University Molecular Design and Synthesis Center synthesized 5′-dCATXATGACGCT-3′ (X = 6-oxo-M1dG) using phosphoramidite solid phase chemistry21 at a 1-μmol scale. The synthesis was conducted using a Perceptive Biosystems Model 8909 DNA synthesizer and Expedite reagents with the standard synthetic protocol for the coupling of the unmodified bases. The 6-oxo-M1dG phosphoramidite was coupled offline for 30 min.25 The 6-oxo-M1dG-modified oligodeoxynucleotide was cleaved from the solid support and the exocyclic amino groups were deprotected overnight in a single step using K2CO3 (0.5 M, methanol) at room temperature. Its complement 5′-dAGCGTCATCATG-3′ was purchased from Integrated DNA Technologies (Coralville, IA). Both oligodeoxynucleotides were further purified by reverse-phase HPLC. The 5′-dCATXATGACGCT-3′:5′-dAGCGTCATCATG-3′ (X = 6-oxo-M1dG) was annealed at 25 °C in 100 mM NaCl, 50 μM Na2EDTA in 50 mM HEPES (pH 7.0). It was lyophilized, dissolved in 1 mL of H2O, and isolated by passing through a hydroxylapatite (HAP) column equilibrated with 200 mM NaH2PO4 (pH 7) to remove single-stranded oligodeoxynucleotide. Buffer A contained 100 mM NaCl, 50 μM Na2EDTA in 10 mM NaH2PO4; buffer B contained 100 mM NaCl, 50 μM Na2EDTA in 200 mM NaH2PO4. A linear gradient from 0% to 25% Buffer B was run for 1 h, followed by a linear gradient from 25% to 100% buffer B for 40 min, and the HAP column was maintained at 100% B for 20 min. The duplex was desalted by passing it through a Sephadex G-25 column.

Melting Temperature (Tm) Experiments

The absorbances of 5 μM 5′-dCATXATGACGCT-3′:5′-dAGCGTCATCATG-3′ (X = 6-oxo-M1dG) and of the corresponding unmodified duplex were measured in 100 mM NaCl, 50 μM Na2EDTA, 10 mM NaH2PO4 (pH 7), at 260 nm, using a 1 cm path length cuvette. Thermal scans were performed from 15 to 85 °C in 1 °C/min intervals. Tm values were determined using the first derivatives of absorbances vs. temperature plots that were obtained experimentally. All experiments were performed on a Cary 100 Bio UV/vis spectrometer.

NMR Spectroscopy

The 5′-dCATXATGACGCT-3′:5′-dAGCGTCATCATG-3′ (X = 6-oxo-M1dG) and the corresponding unmodified duplex were prepared in 100 mM NaCl, 50 μM Na2EDTA, and 10 mM NaH2PO4 (pH 7.0). The concentrations of oligodeoxynucleotide duplexes were 300 μM. The sample volumes were maintained at 600 μL. Data were collected in 5 mm NMR tubes with cryogenic probes (Bruker Biospin, Inc., Billerica, MA). NOESY26,27 spectra of nonexchangeable protons were obtained in 99.996% D2O at 298 K, while NOESY spectra of exchangeable protons were obtained in 95:5 H2O:D2O at 274 K. NOESY and COSY28 spectra for nonexchangeable protons were collected at 900 MHz at 298 K, and at 274 K for exchangeable protons. NOESY data were collected with 2048 real data points in the t2 dimension and 512 real data points in the t1 dimension. NOESY data for nonexchangeable protons were collected at 300, 250 and 100 ms mixing times and a relaxation delay of 1.5 s. The data were zero-filled to obtain final matrices of 2048 × 1024 data points. Data were analyzed using the program TOPSPIN (Bruker Biospin, Billerica, MA) and the program SPARKY.29

NMR Distance Restraints

The program SPARKY was used to integrate the volume of NOESY cross-peaks from spectra obtained with 100, 250 and 300 ms mixing times at 298 K. The intrinsic error of the integrations was given to be one-half of the lowest intensity cross-peak volume. Five levels of value in error were assigned to each cross-peaks. A 10% error was assigned to well-resolved and nonoverlapping cross-peaks. A 20% error was assigned to strong slightly broadened cross-peaks, overlapped cross-peaks, or peaks with a moderate S/N. A 30% error was assigned to strong, but medially broadened, or overlapped cross-peaks. A 40% error was assigned to cross-peaks with weak S/N or that were slightly overlapped. A 50% error value was assigned to cross-peaks that were highly broadened, near the diagonal or water- suppression, or possessed moderate S/N and medial overlap or broadening.30−32 Distance restraints were generated through using the program MARDIGRAS.33 Additional distance restraints were created using canonical Watson–Crick hydrogen bonding distances for base pairs in B-DNA.34 Watson–Crick hydrogen bonding restraints were not used for the T3:A22 base pair, and for the C21 base, which was complementary to X4. The square potential energy wells between T3 H1′ and X4 H8, and C21 H1′ and A22 H8 were set from a lower bound of 5 Å to an upper bound of 7 Å because weak NOEs were observed.

Restrained Molecular Dynamics

Canonical B-DNA values for deoxyribose pseudorotation and phosphodiester backbone torsion angles34 were used as restraints in the rMD calculations. Backbone torsion angle restraints for nucleotides were assigned a potential energy well window of ±30°, except for nucleotides T3, X4, C21, and A22, which were assigned potential energy well windows of ±60°. The backbone torsion angle restraints were centered at the B-DNA backbone torsions α = −60°, β = 180°, γ = 60°, ε = 195°, and ζ = −105°. The partial charges and bond lengths for 6-oxo-M1dG were calculated using the program GAUSSIAN.35 Calculations used the B3LYP/6–13G* basis set. 6-oxo-M1dG was constructed using the program MOE.36 The unmodified duplex containing base G4 was built first, then base G4 was modified to 6-oxo-M1dG using the Builder function in the program MOE. The resulting structure was further processed in the program xLEAP37,38 by adding information on partial charges and bond lengths of 6-oxo-M1dG, which were obtained from GAUSSIAN. The .top and .inp files were generated in xLEAP using the AMBER suite of programs.38

The rMD calculations were performed using the simulated annealing protocol in AMBER and the parm99 force field.39 The generalized Born model40 was used for solvation. The salt concentration was set at 0.1 M. All restraints had an applied force constant of 30 kcal mol–1 A–2. Initial calculations were performed for 100 ps over 100,000 steps. The system was heated from 0 to 600 K for the first 3000 steps with a 0.5 ps coupling. The system was then held at 600 K for 3001–100000 steps. NOE generated distances were compared to intensities calculated from emergent structures using complete relaxation matrix analysis by the program CORMA.33 The 20 structures with the lowest deviations from experimental distance restraints were used to generate an average refined structure. The structural renderings were created in the program CHIMERA.41

Results

Melting Temperature Analyses

To determine the thermodynamic consequences of incorporating the 6-oxo-M1dG lesion into 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ (X = 6-oxo-M1dG), thermal melting experiments were conducted of the modified and the corresponding unmodified duplexes, each at the same concentration and in buffer containing 0.1 M NaCl (Figure 1). In both instances, monophasic melting transitions were observed, indicating that the presence of the 6-oxo-M1dG modification did not alter the concerted conversion of dsDNA into ssDNA, which would be anticipated for oligodeoxynucleotides of this length. The respective Tm values were determined based on the first derivative curves of DNA absorbance at 260 nm collected as a function of temperature. The Tm value of the 6-oxo-M1dG modified duplex was 48.3 °C. In comparison, the Tm value of the unmodified duplex was 62.8 °C. The incorporation of the 6-oxo-M1dG lesion into this DNA duplex destabilized the DNA duplex and resulted in a 14.5 °C decrease in the Tm value, as compared to the corresponding unmodified duplex.

Figure 1.

Figure 1

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Thermal melting curves as recorded by UV 260 absorbance. The data for the modified 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′ (X = 6-oxo-M1dG) duplex is shown in panel A; the data for the unmodified 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′ duplex is shown in panel B. The dotted lines represent heating curves; the solid lines represent corresponding first derivative curves.

NMR Spectroscopy

The 1H NMR spectrum of the 6-oxo-M1dG modified duplex 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ (X = 6-oxo-M1dG) was examined as a function of temperature over the range of 274 to 308 K, which remained below the Tm value for the duplex. The sample was well-behaved and exhibited sharp signals over this temperature range. The optimal temperature for collecting nonexchangeable 1H data was determined to be 298 K. In contrast, the optimal temperature for collecting spectra of the Watson–Crick hydrogen bonded imino- and amino exchangeable protons was determined to be 274 K, as at higher temperatures these resonances experienced increased exchange broadening.

Assignment of Nonexchangeable DNA Protons

COSY NMR data revealed the presence of six 3J COSY cross-peaks corresponding to vicinal couplings of the six cytosine H5 and H6 protons, in both the unmodified duplex 5′-d(CATGATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ and the modified duplex 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ (X = 6-oxo-M1dG) (Figure 2). The presence of the 6-oxo-M1dG lesion in the modified duplex resulted in a significant chemical shift perturbation for the cytosine H5–H6 COSY cross-peak arising from C21 (Figure 2B), the cytosine complementary to the 6-oxo-M1dG lesion (Chart 1). This reflected a downfield shift of the cytosine H5 and H6 protons in the presence of 6-oxo-M1dG.

Figure 2.

Figure 2

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A. Expanded COSY spectrum of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) showing scalar couplings between cytosine H5 and H6 protons; cross-peak labeled X4 shows the scalar coupling between the 6-oxo-M1dG H7 and H8 protons. B. Expanded COSY spectrum of the unmodified duplex 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′. The spectra were acquired at 900 MHz and 298 K.

The sequential NOE assignments for 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ (X = 6-oxo-M1dG) and the unmodified duplex 5′-d(CATGATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ were accomplished using standard protocols42,43 (Figure 3). In the modified duplex, the imidazole proton of 6-oxo-M1dG is numbered as H2, as opposed to the numbering of the corresponding deoxyguanosine imidazole proton as H8 (Chart 1). For the 6-oxo-M1dG modified strand, a continuous set of sequential NOEs was observed. The sequential NOEs between the T3 H1′ and X4 H2 and X4 H1′ protons and between the X4 H1′ and A5 H8 protons were weak, where the intensity of the NOE between the X4 H1′ and A5 H8 protons was the lowest. The remainder of the sequential NOEs from the A5 H8 proton to the T12 H1′ proton were of normal intensities. For the complementary strand, a continuous set of sequential NOEs was observed. The sequential NOE between the T20 H1′ proton and the C21 H6 proton was weak. It was not observed at the 250 ms mixing time. This NOE was observed as a low intensity peak in the 300 ms mixing time data (Figure S1 in the Supporting Information). The NOE between the C21 H6 proton and the A22 H1′ proton was observed but with low intensity. The remaining sequential NOEs from the A22 H6 proton to the G24 H1′ proton were of normal intensities. The complete assignments of the deoxyribose H2′, H2″, and H3′ proton resonances were achieved. Most of the H4′ proton resonances were assigned. Partial assignments were made for the deoxyribose H5′ and H5′′ proton resonances; these were in many instances overlapped precluding unequivocal assignments. The assignments of the DNA nonexchangeable protons are tabulated in Table S1 of the Supporting Information.

Figure 3.

Figure 3

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Expanded plot of the NOESY spectrum of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′ d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG), showing the sequential NOEs between aromatic H6/H8 protons and deoxyribosyl H1′ protons. A. The strand containing X4, 6-oxo-M1dG, showing nucleotides C1 to T12. The X4 H1′ → A5 H8 cross-peak is weak due to the incorporation of 6-oxo-M1dG. B. The complementary strand, showing nucleotides A13 to G24. The NOE connectivity is broken at the T20 H1′→ C21 H6 cross-peak; C21 is the nucleotide opposite to X4. Expanded plot of the NOESY spectrum of the unmodified duplex 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′. C. Nucleotides C1 to T12. D. The complementary strand, showing nucleotides A13 to G24. The spectra were acquired at 900 MHz and 298 K.

Exchangeable DNA Protons

Figure 4 shows expansions of 1H experiments collected in a buffer containing 95:5 H2O:D2O, for the 6-oxo-M1dG-modified and unmodified duplexes. For the 6-oxo-M1dG- modified 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ duplex the deoxyguanosine N1H and deoxythymine N3H imino proton resonances were sharp in 1H spectra collected at 274 K but broadened at higher temperatures. Because of spectral broadening, they were not observed at 298 K. The G10 N1H, G14 N1H, and G16 N1H imino proton resonances were partially resolved. The imino proton resonances were assigned using standard protocols;44 a 1H NOESY spectrum at 274 K is shown in Figure 4. Base pair G7:C18 is flanked by base pairs T6:A19 and A8:T17 (Chart 1); the G7 N1H resonance was assigned based on NOEs observed to both the T6 N3H proton and the T17 N3H proton. The T17 N3H proton exhibited an NOE to the G16 N1H proton, allowing its assignment at δ 12.4 ppm, the farthest downfield resonance in this region of the 1H spectrum. In the NOESY spectrum, the serial G16 N1H → G10 N1H → G14 N1H NOEs were close to the diagonal since the G16 N1H, G10 N1H and G14 N1H resonances were only partially resolved. The T6 N3H proton was assigned at δ 13.3 ppm from its NOE to the T20 N3H proton. On the 5′-side of the 6-oxo-M1dG lesion, a weak NOE between the T3 N3H proton and the T23 N3H proton was observed, likely due to fraying of the thermodynamically destabilized duplex (Figure 1). Nevertheless, in the amino region of the 1H spectrum, NOEs between the T3 N3H imino proton and the A22N6H amino proton, as well as the A22 H2 proton, and between the T23 N3H imino proton and the A2N6H amino proton and A2 H2 proton were observed. For base pairs G7:C18, G10:C15, and C11:G14 (Chart 1), NOEs between the guanine N1H imino protons and the cytosine N4H protons were evident. The NOEs between the thymine N3H and adenine H2 protons were evident for each A:T base pair except for the terminal base pair T12:A13. The guanine N1H imino resonance from the terminal C1:G24 base pair was not observed, which underwent significant exchange broadening even at 274 K. For the unmodified duplex, the imino and amino proton resonances were fully assigned and showed no unusual effects. The chemical shift assignments for the exchangeable DNA protons are provided in Table S2 of the Supporting Information.

Figure 4.

Figure 4

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A. 1H spectrum showing the thymine N3H and guanine N1H imino protons of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG). B. Expanded plot of the NOESY spectrum of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) showing sequential NOE connectivities for imino to amino protons. The cross-peaks are assigned as a, T17 N3H → C18N4Hn; b, T17 N3H → A8 H2; c, T17 N3H → C9N4Hb; d, T23 N3H → A2 H2; e, T6 N3H → C18N4Hb; f, T6 N3H → A19 H2; g, T20 N3H → C21N4Hn; h, T20 N3H → C21N4Hb; i, T20 N3H → A5 H2; j, T3 N3H → A5 H2; k, G14 N1H → C11 H5; l, G14 N1H → C11N4Hn; m, G14 N1H → C11N4Hb; n, G10 N1H → C15 H5; o, G10 N1H → C15N4Hn; p, G10 N1H → C15N4Hb; q, G16 N1H → C9 H5; r, G16 N1H → C9N4Hn; s, G16 N1H → C9N4Hb; t, G7 N1H → C18 H5; u, G10 N1H → C18N4Hb; v, G7 N1H → C18N4Hn; w, G7 N1H → A8 H2; x, G7 N1H→ C18N4Hb. C. Expanded plot of the NOESY spectrum of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) showing sequential NOE connectivities for the imino protons. D. 1H spectrum showing the thymine N3H and guanine N1H imino protons of 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′. E. Expanded plot of the NOESY spectrum of 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ showing sequential NOE connectivities for imino to amino protons. The cross-peaks are assigned as a, T17 N3H → C18N4Hn; b, T17 N3H → A8 H2; c, T17 N3H → C9N4Hb; d, T23 N3H → A2 H2; e, T6 N3H → C18N4Hb; f, T6 N3H → A19 H2; g, T20 N3H → C21N4Hn; h, T20 N3H → C21N4Hb; i, T20 N3H → A5 H2; j, T3 N3H → A5 H2; k, G14 N1H → C11 H5; l, G14 N1H → C11N4Hn; m, G14 N1H → C11N4Hb; n, G10 N1H → C15 H5; o, G10 N1H → C15N4Hn; p, G10 N1H → C15N4Hb; q, G16 N1H → C9 H5; r, G16 N1H → C9N4Hn; s, G16 N1H → C9N4Hb; t, G7 N1H → C18 H5; u, G10 N1H → C18N4Hb; v, G7 N1H → C18N4Hn; w, G7 N1H → A8 H2; x, G7 N1H→ C18N4Hb. F. Expanded plot of the NOESY spectrum of 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′:5′- d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ showing sequential NOE connectivities for the imino protons. The NOESY data were collected at 900 MHz at 250 ms mixing time, at 274 K.

Assignment of 6-oxo-M1dG Resonances

For the 6-oxo-M1dG- modified 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ duplex, the X4 H2 imidazole proton resonance was assigned at δ 8.05 ppm from analysis of 1H NOESY NMR data (Figure 3). The 6-oxo-M1dG lesion was characterized by the characteristic vicinal coupling of the H7 and H8 resonances in the exocyclic ring (Chart 1). The 6-oxo-M1dG 3J COSY cross-peak between X4 H7 and H8 was observed in the COSY spectrum at 298 K, affording the assignment of X4 H7 at δ 7.81 ppm and X4 H8 proton at δ 5.31 ppm (Figure 2). The deoxyribose protons of the 6-oxo-M1dG lesion were identified from the 1H NOESY NMR data. The X4 H1′ resonance was observed at δ 5.40 ppm, the H2′ and H2″ resonances were overlapped at δ 2.48 ppm, the H3′ resonance was observed at δ 4.92 ppm, the H4′ resonance was observed at δ 4.23 ppm, and the H5′ and H5′′ resonances overlapped at δ 4.09 ppm. These were all in the normally anticipated chemical shift ranges. Due to the presence of the exocyclic ring between the N1 and N6 positions of the purine ring, the 6-oxo-M1dG lesion does not have an imino proton. The assignments of 6-oxo-M1dG modification specifically are tabulated in Table S3 of the Supporting Information.

NOEs Between 6-oxo-M1dG and DNA Protons

The observation of NOEs between the 6-oxo-M1dG protons and DNA protons (Figure 5) ultimately allowed the structure of the 6-oxo-M1dG- modified 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ duplex to be determined. The intranucleotide NOE cross-peak between the X4 H1′ resonance (δ 5.40 ppm) and the X4 H2 resonance (δ 8.05 ppm) was of normal intensity. As noted above, the internucleotide NOEs between the T3 H1′ proton and the X4 H2 proton (δ 8.05 ppm), and between the X4 H1′ proton and the A5 H8 proton were weak. The internucleotide cross-peaks between the X4 H2 proton and the other deoxyribose H2′, H2″, protons and T3 H6 proton were observed (Figure S2 of the Supporting Information). They all showed low intensity compared with the corresponding NOEs in the unmodified duplex. The aromatic X4 H7 and X4 H8 protons of 6-oxo-M1dG showed internucleotide connectivities with the T3 and T20 methyl protons (Figure S3 of the Supporting Information), as well as the T3 and T20 deoxyribose H2′, H2″ protons (Figure S4 of the Supporting Information).

Figure 5.

Figure 5

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Internucleotide sequential NOE cross-peaks of X4 and C21 in the aromatic H2/H6/H8 and deoxyribose H1′ region as observed in a 0.25 s mixing time NOESY spectrum of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG). A. The sequential connectivity NOE between T3 H1′ and X4 H8. B. The sequential connectivity NOE between X4 H1′ and A5 H8. C. The sequential connectivity NOE between T20 H1′ and C21 H6. D. The sequential connectivity NOE between C21 H1′ and A22 H8.

Chemical Shift Perturbations

Figure 6 summarizes chemical shift perturbations induced in the 6-oxo-M1dG- modified 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ duplex as compared to the corresponding unmodified duplex. These were clustered at the damage site X4:C21 and the two flanking base pairs 5′ and 3′ to the 6-oxo-M1dG lesion. The greatest perturbation was observed for the C21 nucleobase, which was complementary to the 6-oxo-M1dG modified X4 nucleotide. The C21 H6 proton resonance shifted downfield by 0.7 ppm compared to the corresponding unmodified duplex. As well, the C21 H5 proton resonance shifted downfield by 0.25 ppm compared to the unmodified duplex. This may be observed in the 1H COSY spectrum shown in Figure 2. In contrast, the imidazole ring H2 proton of the X4 6-oxo-M1dG-modified nucleotide shifted upfield by 0.25 ppm. Significant downfield chemical shifts were also observed for the X4 H1′, X5 H1′, and C21 H1′ anomeric proton resonances. Of the Watson–Crick hydrogen bonded imino protons, at base pair T3:A22, the T3 N3H imino proton shifted downfield.

Figure 6.

Figure 6

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Chemical shift changes of protons H2/H6/H8 (green) and H1′ (blue) of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) relative to 5′-d(C1A2T3G4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′. A. Chemical shift changes of protons of each nucleotide of template strands. B. Chemical shift changes of protons of each nucleotide of complementary strands.

Structural Refinement

Distance Restraints

A total 320 NOE-derived distance restraints were obtained from the 1H NOESY data for nonexchangeable protons (Table S4 in the Supporting Information). These distances were calculated using the program MARDIGRAS.33 Of these, there were 169 intranucleotide distance restraints, 151 internucleotide distance restraints, and 22 distance restraints involving the 6-oxo-M1dG lesion, which were critical to determining the structure of the 6-oxo-M1dG- modified 5′-d(CATXATGACGCT)-3′:5′-d(AGCGTCATCATG)-3′ duplex. The experimental distance restraints were combined with 35 additional empirical Watson–Crick hydrogen bonding restraints, 95 backbone torsion angle restraints, and 70 deoxyribose pseudorotation restraints based on the canonical B-DNA structure.34 The complete list of pseudorotation restraints is provided in Table S5 of the Supporting Information.

Restrained Molecular Dynamics (rMD) Calculations

The 6-oxo-M1dG was constructed using the Builder function in the program MOE.36 The resulting structure was further processed using the program xLEAP;37,38 the partial charges and bond lengths were calculated using the B3LYP/6–13G* basis set in the program Gaussian.35 (Tables S6, and S7 in the Supporting Information). A series of 20 100 ps rMD calculations were performed, using a simulated annealing protocol.39 The generalized Born model40 was used for solvation at 0.1 M salt concentration. A total of 520 experimental and canonical restraints with applied force constants of 30 kcal mol–1 A–2 were applied. These are collected in Table 1. Because the 6-oxo-M1dG lesion created structural perturbation at base pairs T3:A22 and X4:C21, canonical restraints were not employed for X4, A5, T20, C21, and A22 in the 6-oxo-M1dG modified region, nor were they employed at the terminal nucleotides C1, T12, A13, and G24. Each of the 20 emergent structures was further subjected to potential energy minimization using the conjugate gradients algorithm. After potential energy minimization, the 20 structures emergent from the rMD calculations exhibited a maximum rms pairwise difference of 1.59 Å, indicating that the restraints employed in the rMD calculations allowed satisfactory convergence of the individual calculations toward a common structure. From these 20 energy-minimized structures, an average structure was also calculated, using the program SUPPOSE. Its potential energy was minimized using the conjugate gradients’ algorithm.73 The average structure had a rms pairwise difference of 1.09 Å as compared to the 20 individual structures. An overlay of 20 energy minimized emergent structures from the rMD calculations with the average minimized structure is shown in Figure 7. These structures revealed the intercalation of the 6-oxo-M1dG lesion into the duplex, and the extrusion of the complementary C21 nucleotide out of the duplex and into the major groove. As anticipated, the convergence was not as good at the two ends of the modified oligodeoxynucleotide duplex, which were presumed to undergo base pair fraying at 298 K.

Table 1. NMR Restraints Used for the 6-oxo-M1dG Modified Duplex Structural Refinement.
internucleotide restraints 151
intranucleotide restraints 169
6-oxo-M1dG restraints 22
total NOE restraints 320
backbone torsion angle restraints 95
hydrogen bonding restraints 35
deoxyribose pseudorotation restraints 70
total number of restraints 520
number of distance restraints violations 50
number of torsion angle restraint violations 11
total distance penalty/maximum penalty (kcal/mol) 129.712/7.800
total torsion angle penalty/maximum penalty (kcal/mol) 0.995/0.073
rms distances (Å) 0.0228
rms angles (deg) 2.305
distance restraint force field (kcal/mol/Å2) 30
torsion angle restraint force field (kcal/mol/deg2) 30
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Figure 7.

Figure 7

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Overview of the 20 lowest energy violation structures generated from rMD calculations of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) carried out using a simulated annealing protocol and experimental NOEs generated distance restraints.

Complete Relaxation Matrix Analysis

The average energy minimized structure was evaluated as to its accuracy by complete relaxation matrix analysis using the program CORMA.33 Each of the individual intra- and internucleotide sixth root residuals RX1 were equal or less than 0.1 (Figure 8). The average RX1 values of all intranucleotides and internucleotides were 0.0695 and 0.0679, respectively; the overall RX1 value of the duplex containing 6-oxo-M1dG was 0.0689 (Table 2). The complete relaxation matrix analysis suggested that the average refined structure of the 6-oxo-M1dG duplex was in reasonable accordance with the experimental NOE data.

Figure 8.

Figure 8

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Calculation of sixth root residual values (RX1) between theoretical NOEs predicted by complete relaxation matrix calculations and experimental NOEs for the averaged refined structure of 5′-d(C1A2T3X4A5T6G7A8C9G10C11T12)-3′:5′-d(A13G14C15G16T17C18A19T20C21A22T23G24)-3′ (X = 6-oxo-M1dG) emergent from the rMD calculations, using CORMA. A. The RX1 values of intranucleotide and internucleotide NOEs in the 6-oxo-M1dG modified strand. B. The RX1 values of intranucleotide and internucleotide NOEs in the unmodified complementary strand. The intranucleotide NOE residuals are colored in black. The internucleotide NOE residuals are colored in gray.

Table 2. RMS Differences and Sixth Root Residual (RX1) Values for the Average Structure of the 6-oxo-M1dG Modified Duplex Emergent From rMD Calculations.
greatest RMS pairwise difference between 20 structures 1.59 Å
greatest RMS difference from the average structures 1.09 Å
sixth root residual RX1a calculated for the intranucleotide distances in the average structure, using CORMAb 0.069
sixth root residual RX1a calculated for the internucleotide distances in the average structure, using CORMAb 0.068
sixth root residual RX1a calculated for all distances 0.069
in the average structure, using CORMAb
average errorc 0.00102
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a

RX1 is the sixth root R factor Σ[((Io)i1/6)–((Ic)i1/6Σ((Io)i1/6].

b

Mixing time was 250 ms.

c

Average error: Σ(Ic – In)/n, where Ic is NOE intensities calculated from the refined structure, and In is experimental NOE intensities.

d. Unmodified DNA Duplex

The structure of the unmodified DNA duplex was also determined from NOE data using the same strategy as the 6-oxo-M1dG modified duplex, described above. For the unmodified duplex, a total of 202 NOE distance restraints were obtained from the 1H NOESY spectrum, which combined additional 42 empirical Watson–Crick hydrogen bonding restraints, 99 backbone torsion angle restraints, and 100 deoxyribose pseudorotation restraints based on the canonical B-DNA structure. Overall, a total of 446 experimental and canonical restraints were employed in the rMD calculations (Table S8 in the Supporting Information). Figure S5 in the Supporting Information shows the overlay of 20 potential energy-minimized structures emergent from the rMD calculations for the unmodified duplex, along with the average minimized structure. These results indicated excellent convergence.

Structure of the 6-oxo-M1dG Modified Duplex

Figure 9 shows views of the 6-oxo-M1dG modified duplex from the major and minor grooves, respectively, as well as views of the same region of the unmodified duplex from the major and minor grooves. In comparison, the unmodified duplex remained in a B-DNA type conformation. The structure of the modified duplex revealed that 6-oxo-M1dG intercalated into the duplex. It remained in the anti-conformation around the glycosyl bond. This placed the imidazole ring proton X4 H2 into the major groove, and the cytosine O4 oxygen atom faced into the major groove. The 6-oxo-M1dG H7 and H8 protons faced toward the major groove. The complementary cytosine C21 was extruded out of the duplex and toward the major groove, which allowed the bulky 6-oxo-M1dG lesion to be accommodated within the helix. The localization of the structural perturbations to the damaged base pair X4:C21 and its 5′- and 3′-neighboring base pairs T3:A22 and A5:T20 was evident; the flanking base pairs remained within the Watson–Crick hydrogen bonding contact. Figure 10 shows views along the helical axis showing base stacking interactions. The intercalation of 6-oxo-M1dG placed it between flanking base pairs T3:A22 and A5:T20. It showed more favorable stacking with the 3′-neighbor base pair A5:T20.

Figure 9.

Figure 9

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Conformations of the average structures from the 20 lowest energy violation structures of 6-oxo-M1dG modified region and the same region of unmodified duplex from the rMD calculations. A. Base pairs T3:A22, X4:C21 and A5:T20 of 6-oxo-M1dG modified duplex DNA as seen from the major groove. B. Base pairs of T3:A22, X4:C21 and A5:T20 6-oxo-M1dG modified duplex DNA as seen from the minor groove. C. Base pairs of T3:A22, G4:C21 and A5:T20 of unmodified duplex DNA as seen from the major groove. D. Base pairs of T3:A22, G4:C21 and A5:T20 of unmodified duplex DNA as seen from the minor groove. The base pairs T3:A22, X4/G4:C21, and A5:T20 are colored in purple, red, and green, respectively.

Figure 10.

Figure 10

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Base stacking interactions of the average structures of 6-oxo-M1dG region and the same region of unmodified duplex DNA from the 20 lowest energy violation structures from the rMD calculations. A. View showing the T3:A22 of 6-oxo-M1dG modified duplex DNA base pair in purple and X4:C21 in red. B. View showing X4:C21 in red and the A5:T20 of 6-oxo-M1dG modified duplex DNA in green. C. View showing the T3:A22 of unmodified duplex DNA base pair in purple and G4:C21 in red. D. View showing G4:C21 in red and the A5:T20 of the modified duplex DNA in green.

Discussion

The M1dG lesion, produced endogenously in DNA by base propenals1,3,45 and lipid peroxidation,2,4,6,7 has been identified in DNA from rodent46 and human47,48 tissue samples, as have other exocyclic purine lesions49,50 by mass spectroscopic,47,51,52 postlabeling,53,54 and immunochemical55 techniques. It also forms in mitochondrial DNA.56 M1dG is genotoxic in human cells,57 producing both base pair substitutions and frameshifts.22 It is bypassed by error-prone Y-family polymerases, including hpol η.23 Once formed, M1dG may oxidize to 3-(2-deoxy-β-D-erythropentofuranosyl)-pyrimido[1,2-f]purine-6,10(3H,5H)-dione (6-oxo-M1dG).19 6-oxo-M1dG represents a strong block to DNA replication,24 but the Y-family polymerase hPol η preferentially inserts dAMP or dGMP across from 6-oxo-M1dG,24 suggesting that like M1dG, 6-oxo-M1dG is mutagenic. Significantly, as compared to the initially formed M1dG lesion, 6-oxo-M1dG is stable in DNA, whereas when placed opposite dC, M1dG rearranges to OPdG.18 Thus, the structural biology of the 6-oxo-M1dG lesion in dsDNA is of interest, as it is anticipated to introduce perturbations into DNA different from those of the OPdG lesion. Crystallographic structures of both insertion and extension complexes with hPol η revealed the role of 3′-neighbor base stacking involving the 6-oxo-M1dG lesion during lesion bypass.24

Base-Displaced Intercalated Structure of 6-oxo-M1dG in dsDNA

Intercalation of 6-oxo-M1dG

The structure of 6-oxo-M1dG in dsDNA emergent from the rMD calculations shows intercalation into the DNA duplex. The complementary base C21 extrudes from the helix toward the major groove. This may be characterized as a “base-displaced” intercalation complex. The intercalated 6-oxo-M1dG lesion creates a localized perturbation of the duplex (Figure 7). This is consistent with the observation that chemical shift perturbations are also localized at the T3:A22, X4:C21, and A5:T20 base pairs (Figure 6), indicating that as one moves beyond the nearest neighbor base pairs, the right-handed helical structure of the oligodeoxynucleotide duplex is reasonably maintained in a B-DNA type of geometry. Likewise, NOEs between the imino and amino regions of the NOESY spectrum (Figure 4) suggest regular Watson–Crick base pairing outside the immediate vicinity of the 6-oxo-M1dG lesion, which is also consistent with a structural perturbation localized at the modified X4:C21 base pair and its immediate 3′- and 5′-neighboring base pairs. Significantly, the 6-oxo-M1dG lesion remains in the anti-conformation about the glycosyl bond (Figure 9), as is evidenced by the weak intranucleotide NOE between the X4 H2 and X4 H1′ protons. The observation of the 6-oxo-M1dG H7 proton resonance at δ 7.81 ppm and X4 H8 proton resonance at δ 5.31 ppm (Table S3 in the Supporting Information) is consistent with the intercalation of the lesion. However, the weak sequential internucleotide NOE between X4 H2 (the imidazole ring proton) and A5 H1′ suggests that it is positioned further toward the major groove than that in the corresponding unmodified duplex. The X4 H2 proton shifts upfield approximately 0.25 ppm, as compared to the unmodified duplex (Figure 5). This is attributed both to the stacking interaction observed between the 6-oxo-M1dG and the 3′-neighboring A5:T20 base pair, as well as the positioning of the X4 H2 proton below the pyrimidine ring of T3 in the 5′-neighboring T3:A22 base pair. Chemical shifts observed in the NOESY spectral region of the pyrimidine H6 and purine H8 protons and their respective deoxyribose H1′ protons are relatively large at X4 and the 5′-neighboring T3 and 3′-neighboring A5 nucleotides in the template strand (Figure 3).

Base-Displaced Extrusion of the Complementary Nucleobase C21

The complementary nucleobase C21 extrudes from the DNA helix and into the major groove. Consistent with this conclusion, in the comparison of the modified vs. unmodified duplexes (Figure 3), nucleotide C21, complementary to the 6-oxo-M1dG nucleotide X4, exhibits the greatest chemical shift perturbation. COSY data show that the C21 H6 resonance (δ 7.75 ppm) shifts downfield by about 0.75 ppm compared with the corresponding C21 H6 resonance of the unmodified DNA duplex, indicative of reduced π stacking interactions. As well, the weak sequential NOE between T20 H1′ and C21 H6, which is observed as a low intensity peak in 300 ms mixing time data (Figure S1 in the Supporting Information) and the weak NOE between C21 H6 and A22 H1′ (Figure 3) are consistent with extrusion of C21 from the helix toward the major groove. The formation of the base-displaced intercalated structure is accompanied by helix unwinding (Figures 7 and 9). Compared with the unmodified duplex (Figure S5 in the Supporting Information), the unwinding of the dsDNA induced by the base-displaced intercalation complex was evidenced by a reduction in the distance between X4 H1′ and A5 H8 as well as the distance between T20 H1′ and C21 H6.

Thermal Destabilization of the Base-Displaced Intercalated Complex

The intercalation of 6-oxo-M1dG disrupts normal Watson–Crick base stacking between the modified X4:C21 base pair and 5′-neighbor base pair T3:A22, as well as the 3′-neighbor base pair A5:T20 (Figure 10). This, combined with the inability of the X4:C21 base pair to support Watson–Crick hydrogen bonding, and the concomitant displacement of C21 into the major groove, likely explains the 15 °C reduction of the thermal stability of the DNA duplex compared with the unmodified duplex (Figure 1). The observation that at the flanking base pairs T3:A22 and A5:T20, both the T3 N3H and T20 N3H imino protons are observed (Figure 4), albeit broadened, indicates that both flanking base pairs remain within Watson–Crick hydrogen bonding distance. This is consistent with the localization of helical disruption to the modified base pair X4:C21 and the two flanking base pairs (Figures 6 and 7). The broadening of the T3 N3H and T20 N3H imino resonances is undoubtedly due to increased exchange with solvent water, consistent with a greater degree of helical disorder. However, structural analyses indicate that 6-oxo-M1dG exhibits superior stacking with 3′-neighboring base pair A5:A20 as compared to 5′-neighboring base pair T3:A22 (Figure 10).

Genotoxicity of 6-oxo-M1dG

It seems plausible that the mutagenic consequences of M1dG formation in DNA are modulated by downstream chemical rearrangements of this lesion in DNA. The respective roles of the initially formed M1dG lesion, and the potential downstream products 6-oxo-M1dG, or OpdG (Scheme 1), in modulating mutagenesis, remain incompletely understood. The 6-oxo-M1dG lesion of interest herein cannot undergo Watson–Crick base pairing and consequently is anticipated to be genotoxic. Consistent with this notion, it blocks DNA replication.24 Christov et al.21 demonstrated that hPol ι incorporated deoxycytidine triphosphate (dCTP) and thymidine triphosphate (dTTP) across from 6-oxo-M1dG with approximately equal efficiencies, but deoxyadenosine triphosphate (dATP) and deoxyguanosine triphosphate (dGTP) were poor substrates. Moreover, after incorporation of a single dNTP opposite the 6-oxo-M1dG, the lesion blocked further replication.21 More recent studies examined mutagenic bypass of 6-oxo-M1dG by additional Y-family polymerases.24 These showed that hPol η inserts dATP or dGTP across from 6-oxo-M1dG,. Crystal structures of DNA containing 6-oxo-M1dG complexed with hpol η in an extension complex with dGTP positioned in the primer opposite 6-oxo-M1dG provided clues as to why the lesion hinders extension.24 In these structures, template 6-oxo-M1dG did not interact with critical residues in the hpol η active site. Moreover, in the active site, the 3′-hydroxyl group of incoming dGTP was not optimally positioned to form a phosphodiester bond with the incoming dCTP nucleotide.24 Instead, 6-oxo-M1dG favored stacking interactions with the 3′-neighbor base pair. Significantly, the present structure of 6-oxo-M1dG in dsDNA shows that absent the polymerase, in the base-displaced intercalated complex, 6-oxo-M1dG also exhibits favorable stacking interactions with the 3′-neighbor base pair (Figure 10), suggesting that they occur independently of glycosylase binding to the damaged DNA.

Previous studies illustrated consequences associated with the reversible structural rearrangement of the initially formed M1dG lesion to OPdG. When present in dsDNA opposite dC, the ring-opened OpdG is favored,18 but in ssDNA, the ring-closed M1dG lesion is favored. At neutral pH, OpdG does not form in dsDNA when thymine is mis-matched opposite M1dG. Thus, it is thought that in dsDNA the rearrangement of M1dG to OPdG is catalyzed by the exocyclic amino group of the complementary cytosine.18 Significantly, OPdG is accommodated in dsDNA with the OPdG 3-oxo-1-propenyl moiety extended into the minor groove, while the complementary cytosine protruded toward the major groove, allowing dsDNA to maintain Watson–Crick hydrogen bonding.14 This is believed to facilitate replication bypass and likely explains why OpdG is less mutagenic than M1dG. Consistent with this notion, the dG adduct derived from acrolein, γ-hydroxyl-1,N2-propano-2′-deoxyguanosine, also exists opposite dC in dsDNA primarily as its ring-opened derivative58 and is not miscoding, in vivo.59,60

Reversion assays in tester strains of Salmonella typhimurium carrying the hisD3052 gene as a reporter revealed the unusual capability of MDA to induce frameshift mutations.61 These were observed in site-specific mutagenesis experiments62 when M1dG was placed into frameshift-prone iterated repeat sequences.22,63 Like M1dG, the exocyclic ring of 6-oxo-M1dG cannot be easily accommodated in the DNA helix, but unlike M1dG, 6-oxo-M1dG cannot reversibly undergo ring-opening when placed opposite the dC in DNA, allowing for Watson–Crick bypass of the damage site. From a structural perspective, 6-oxo-M1dG is anticipated to be similar to 1,N2-propano-dG (PdG), a stable analogue of M1dG (Scheme 1),64 which also cannot undergo ring opening. Site-specific mutagenesis studies employing PdG in iterated CG repeat sequences revealed that in addition to point mutations,65 it induced frameshifts.66,67 Like 6-oxo-M1dG, structural studies of site-specific PdG adducts also revealed base-displaced intercalation. Significantly, PdG undergoes rotation about the glycosyl bond, into the syn conformation, allowing protonated Hoogsteen-like base pairing at the damage site.68−70 Previous studies also examined the structure of both PdG and M1dG incorporated into a duplex containing a two-nucleotide bulge in the modified strand, modeling −2 bp strand slippage deletions associated with the (CpG)3-iterated repeat hotspot for frameshift mutations from the Salmonella typhimurium hisD3052 gene.14 In single-strand DNA, equilibrium between M1dG and OPdG (Scheme 1) favors M1dG, consistent with the observation in studies with M1dG that OPdG was not observed in the bulged duplex. Instead, the bulge was localized, consisting of M1dG and the 3′-neighbor dC. Overall, in the bulged duplex, the structure of M1dG was like that of PdG.71 In contrast, when M1dG was placed into a fully complementary (CpG)3-iterated repeat duplex, as anticipated, ring-opening to OPdG was facilitated.72 In summary, these observations suggest that further studies of 6-oxo-M1dG, probing for Hoogsteen-like protonated base pairing interactions, and in frameshift-prone DNA sequences, such as the reiterated (CpG)3 sequence, are warranted.

Conclusions

The present results reveal a base-displaced intercalated structure of the 6-oxo-M1dG oxidation product of M1dG in dsDNA. The 6-oxo-M1dG lesion cannot undergo ring-opening rearrangement as does M1dG, and it creates a greater structural perturbation to dsDNA than does the OpdG ring-opened rearrangement product of M1dG. The 6-oxo-M1dG intercalates into the helix, while the complementary nucleotide C21 is extruded toward the major groove. The 6-oxo-M1dG lesion remains in the anti-conformation about the glycosyl bond. Normal Watson–Crick base stacking interactions are disrupted between the damaged X4:C21 base pair and its 5′-neighbor base pair T3:A22, and the 3′-neighbor A5:T20 base pair. Superior stacking is observed between 6-oxo-M1dG and the 3′-neighboring A5:T20 base pair. The inability of the damaged X4:C21 base pair to support Watson–Crick hydrogen bonding, and the displacement of C21 into the major groove, combined with the altered base stacking interactions at the lesion site, are each consistent with the reduced thermal stability of the 6-oxo-M1dG modified DNA. This distortion of the DNA duplex is also consistent with the notion that 6-oxo-M1dG is a strong block to DNA replication and that it may be more genotoxic than is the OpdG rearrangement product of M1dG.

Acknowledgments

We thank Dr. Markus Voehler for assistance with NMR. We thank Dr. Michelle L Reyzer and Dr. McDonald Hayes for assistance in MALDI mass spectrometry.

Glossary

Abbreviations

COSY

correlation spectroscopy

NMR

nuclear magnetic resonance

NOESY

nuclear Overhauser effect spectroscopy

RP-HPLC

reverse-phase high-performance liquid chromatography

TOCSY

total correlated spectroscopy

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.chemrestox.3c00226.

  • Table S1, chemical shifts of nonexchangeable DNA protons; Table S2, chemical shifts of exchangeable DNA protons; Table S3, NOE distance restraints used for rMD calculations in AMBER; Table S4, pseudorotation angle restraints used for rMD calculations in AMBER; Table S5, calculated partial charges for 6-oxo-M1dG; Table S6, calculated bond lengths for 6-oxo-M1dG; and Figure S1, hydroxylapatite chromatography data (PDF)

Author Contributions

CRediT: Yizhi (Vera) Fu data curation, formal analysis, investigation, methodology; Robyn M Richie-Jannetta data curation, formal analysis, investigation, methodology; Lawrence J. Marnett funding acquisition, resources, supervision, writing-review & editing; Michael P Stone conceptualization, funding acquisition, project administration, resources, supervision, writing-review & editing.

This work was supported by NIH grants R01 ES-029357 (to M.P.S.) and R01 CA-87819 (to L.J.M.). The Vanderbilt-Ingram Cancer Center was funded by NIH grant P30 CA-068485. Funding for the NMR spectrometers was provided by in part by NIH instrumentation grants S10 RR-05805, S10 RR-025677, and National Science Foundation Instrumentation Grant DBI 0922862, the latter funded by the American Recovery and Reinvestment Act of 2009 (Public Law 111-5). Vanderbilt University assisted with the purchase of NMR instrumentation. Funding for open access charge: National Institutes of Health.

The authors declare no competing financial interest.

Supplementary Material

tx3c00226_si_001.pdf (718.5KB, pdf)

References

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