Structure of an oligonucleotide containing a N-(2-deoxy-β-D-erythro-pentofuranosyl)formamide residue facing a guanine

Biochimie 82 (2000) 65−69 © 2000 Société française de biochimie et biologie moléculaire/Éditions scientifiques et médicales Elsevier SAS. All rights reserved. S0300908400003424/FLA

Structure of an oligonucleotide containing a N-(2-deoxy-β-D-erythro-pentofuranosyl)formamide residue facing a guanine Corinne Maufraisa, George Victor Fazakerleya, André Guyb, Jean Cadetb, Yves Boularda* a

CEA, Service de Biochimie et Génétique Moléculaire, Bât. 142, CEA Saclay, 91191 Gif-Sur-Yvette cedex, France b CEA, Service de Chimie Inorganique et Biologique, Département de Recherche Fondamentale sur la Matière Condensée, 38054 Grenoble cedex, France (Received 20 July 1999; accepted 1 October 1999)

Abstract — Formamide residue is a major oxidative DNA damage product from ionizing radiation on thymine residues in DNA. We report NMR and molecular modeling studies on a DNA duplex structure which contains guanine opposite formamide residue. Formamide residue exists as either the cis and trans isomer. For the trans and the cis isomers, we find that guanine and formamide are stacked inside the helix and are hydrogen bonded. The oligonucleotide adopts globally a B form structure for the two isomers. Conformational changes are observed between the two isomers. © 2000 Société française de biochimie et biologie moléculaire/ Éditions scientifiques et médicales Elsevier SAS DNA lesions / NMR / molecular modeling

1. Introduction Ionizing radiation produces free radicals in aqueous solution and can give rise to base modifications, single and double strand breaks [1, 2]. These lesions have been implicated in biological processes such as mutagenesis, carcinogenesis and lethality. When thymine is γ-irradiated in aqueous solution under aerobic conditions the predominant degradation product is N-(2-deoxy-β-D-erythropentofuranosyl)formamide. Only a few results have been reported on the assessment of the biological role of the formamide. It was shown that the base fragmentation products such as formamide can be bypassed by DNA polymerase or is able to either promote nucleotide incorporation or induce a deletion [3]. Formamide residues promote, preferentially, misincorporation of guanine. The formamide lesion prevents hydrolysis of the 3’ phosphoester bond to the modified nucleoside by nuclease P1 [4], but no enzyme repair study of formamide by either Fpg or endo III has been performed up to now. Results obtained with the formamide residue can be compared with those reported for the apurinic sites in term of mutagenicity [57]. The formamide lesion has lost part but not all of the coding information and it is still able to form hydrogen bonds. Formamide is characterized by the presence of one

Figure 1. Structure of deoxyriboformamide (F). Left, trans isomer about the N1-C2 amide bond; right, cis isomer about the N1-C2 amide bond.

hydrogen bond acceptor and one donor. The rotation of the amide bond of the formamide product leads to cis and trans isomers (figure 1), which have been previously defined and observed [8, 9]. In an attempt to understand why the formamide residue can act as a strong block to DNA polymerase and to determine the reason for the preferential insertion of guanine, we report NMR and molecular modeling studies on an oligonucleotide containing a guanine opposite the formamide lesion in DNA. 2. Materials and methods 2.1. NMR spectroscopy

* Correspondence and reprints Abbreviations: NMR, nuclear magnetic resonance; NOESY, nuclear Overhauser spectroscopy; DQF-COSY, doublequantum-filtered-correlation spectroscopy.

The decamer containing the N-(2-deoxy-β-D-erythropentofuranosyl) formamide residue was synthesized as previously reported [3]. We mixed the two single strands

66 at 80 °C and followed the ratio between the two strands by integrating 1D NMR spectra. The sequence is: 5’-A1-G2-G3-A4-G5-C6-C7-A8-C9G10-3’; 3’-T20-C19-C18-T17-F16-G15-G14-T13-G12C11-5’. The duplex was 2 mM in single strand concentration dissolved in 10 mM phosphate buffer, 150 mM NaCl and 0.2 mM EDTA. Chemical shifts were measured relative to the internal reference 3-(trimethylsilyl)propionate (TSP) at 0 ppm. NMR spectra were recorded on either DRX500 or DRX600 Bruker spectrometers in 99.99% D2O or 90%H2O/10%D2O. NOESY spectra were recorded with 40 to 400 ms mixing times, in the phase sensitive mode [10]. In H2O, the water signal was suppressed with the Watergate sequence [11]. DQF-COSY were recorded with time-proportional phase incrementation [12]. 2.2. Molecular modeling All initial structures were generated from canonical B-DNA [13] and all energy refinements were carried out with the program Amber [14]. The formamide position, inside the helix, was optimized by exploring the conformational space. A set of 24 conformations was generated by varying the χ angle of the formamide. Three types of constraints were applied during the first minimization: torsion angles, NMR distances and weak reinforcement of the hydrogen bonds of the terminal base pairs to avoid end fraying. The second minimization was performed without any constraints in order to relax the obtained conformation [15, 16]. All minimizations were stopped when the root mean square (rms) energy gradient was less than 0.1 kcal/mol–1 Å–1. 3. Results and discussion 3.1. NMR spectroscopy A NOESY spectrum of the non-exchangeable protons recorded with a 400 ms mixing time gave the proton assignment. We were able to follow the classical intra- and internucleotide connectivities for the two strands. Figure 2 shows part of the NOESY spectrum for the duplex. The interactions for the central part of the strand are normal. In particular, internucleotide interactions with G5H8 suggest that no major disruption of the helix takes place from A4 to C6, the residues which could be influenced by the formamide on the opposite strand. For the strand containing the formamide residue, the chain of connectivities can be followed from C11 to T20 without interruption. For the central part of the duplex, we observe a series of interactions between the residues G15-F16-T17 which show that these bases are sequentially stacked over each other. For this strand, all the interactions observed are typical of a B-DNA conformation. One or both of the interbase NOEs with G5 and F16 are observed. This means that forma-

Maufrais et al. mide, as well as G5, are well stacked inside the helix. We note here and in other regions of the NOESY spectrum the presence of a minor species. In a NOESY spectrum recorded with a short mixing time (60 ms), we observed that all the bases were anti and all the sugars of the Watson-Crick residues were predominantly C2’-endo [17]. In a DQF-COSY spectrum, we can measure the necessary coupling constants to determine the sugar conformation of formamide. The sugar pucker of formamide is predominantly C2’-endo. NOE’s build-up curves have been measured in order to generate a low resolution structure. Part of an NMR spectrum in H2O (figure 3) shows the intra- and internucleotide connectivities corresponding to the formation of the Watson-Crick base pairs of the duplex. The formation of the base pairs C6-G15 and A4-T17 is shown by the interactions of the imino proton of G15 with the amino and H5 protons of C6 respectively and by the interactions of the imino proton of T17 with the amino and H2 protons of A4. The chemical shift of the imino proton of G5 is characteristic of an imino proton hydrogen bonded to an oxygen atom. We observed an interaction between the G5 imino proton and a resonance which corresponds to its amino protons not involved in interstrand hydrogen bonding [16]. For normal hydrogen bonding the amino protons would give rise to a broad and generally not observable resonance. We have identified the amide proton of F16 from its interactions with G5 and the neighboring base pairs. This proton observed at 8.24 ppm indicates that it is not hydrogen bonded to a nitrogen. There remain, however, unassigned peaks in the spectra in D2O and H2O. In the NOESY spectra, we were able to find certain connectivities between these peaks. They belong to a minor species which correspond to ca. 40% of the total sample. In D2O, we observe that the chemical shifts of the residues are different only at the center of the duplex between the two species. For the G strand, we observe a splitting of the resonances. For the formamide containing strand, we can follow the connectivities expected for a right-handed B-DNA helix. Only the assignment of the center of the duplex is modified compared to that of the major species. The chemical shift of the F16*H2 proton of the minor species moves from 7.16 ppm to 6.71 ppm. However, the NOE cross-peaks observed between the formyl proton of formamide and the protons of the bases on each side show that formamide of the minor species is stacked inside the helix, too. In a NOESY spectrum recorded with a short mixing time (60 ms), we observed that all the bases of the minor species were anti and all the sugars of the Watson-Crick residues were predominantly C2’-endo. In a DQF-COSY spectrum, the formamide sugar pucker could not be determined due to resonance overlap. In H2O, as for the non-exchangeable protons, the structural differences between the two species only lead to differences in the chemical shifts at the center of the

Conformational changes in formamide residue

67

Figure 2. Expanded contour plot of the aromatic region of the NOESY spectrum (400 ms mixing time) of the duplex containing the formamide in D2O, 13 °C and pH 7.0. The interactions between the F16 formyl proton and the neighboring bases are shown in the boxes.

duplex. The chemical shifts of the imino protons of the minor species and the interactions of these imino protons with the amino, H2 and H5 protons showed that all the guanines and thymines of the minor species formed Watson-Crick base pairs. We note that for the minor species the F16*NH proton is involved in a hydrogen bonding with a nitrogen. Finally, as for the major species, the G5*NH proton forms an hydrogen bond with an oxygen atom while its amino group does not. 3.2. Molecular modeling The two species observed in our NMR data correspond to the trans and cis isomers of formamide as shown according to the synthesis of the monomer. In a previous NMR study, it has been shown that the major species is trans and the minor species is cis [8]. The two species are intrahelical from NOEs observed between formamide and the adjacent bases. Thus, the initial structures for the duplex were based upon the NMR observations that the duplexes adopt globally a B-DNA conformation with no major distortion. We have taken into account that the sugar

pucker of the major species is C2’-endo, but cannot define that of the minor species. To define the sugar/formamide orientation of the cis and trans isomers, we have generated a series of structures by varying the χ angle from the anti to the syn conformation. For the major species, a series with a C2’-endo conformation for formamide was generated, while for the minor species two series were built: with a C2’-endo and with a C3’-endo conformation. The structures, fitting best the NMR constraints and optimizing the hydrogen bond between the imino proton of G5 and the oxygen atom of F16 as observed in H2O show a anti conformation for the major (trans) species of the formamide and a syn conformation for the minor (cis) species. For the minor species, the structure fitting best the NMR data has formamide with a C2’-endo conformation. The minimizations were carried out with the constraints as described above. The best final structures were chosen from those found to be in good agreement with the NMR data and low energy value. For the major and minor species, the best final structures are shown in figure 4A, B, respectively. These results

68

Maufrais et al.

Figure 3. Expanded contour plot of the imino and the amino/H2 region of a NOESY spectrum (150 ms mixing time) recorded at 2 °C in H2O. The pair of protons of the amino groups of cytosine (C) and adenine (A) are connected by a continuous line. Only the cross-peaks of the central part are labeled. The major species is labeled with a solid line and the minor species with a star and dotted line.

clearly show the possible formation of a hydrogen bond between the formamide and G5 on the opposite strand. For the cis isomer, it is possible to form a hydrogen bond between F16NH...G15N3 without deforming the structure from that of a B-DNA helix. 4. Conclusion With the oligonucleotide containing guanine opposite formamide lesion, our results show that the cis and trans isomers are integrated in a right-handed B-DNA WatsonCrick helix. The insertion of the formamide in the helix does not lead to a local perturbation of the duplex geometry and does not disturb the adjacent base paring. From our NMR data and from model building studies, the

cis and trans isomers of formamide can pair with guanine on the opposite strand with one hydrogen bond. The cis isomer has a syn orientation to favor the formation of this hydrogen bond and formamide is stabilized with an additional hydrogen bond with guanine on the 5’ side. We note that for both isomers, the amino group of guanine facing formamide is not involved in hydrogen bonding. When guanine is incorporated in a duplex structure opposite an apurinic site, an intrahelical form has been observed for both guanine and deoxyribose. Our result show that formamide facing guanine has a similar structure to that of the apurinic site [17]. More detailed NMR and molecular dynamics studies (in particular with explicit water molecules) on the systems described here are currently in progress. Duplexes with either an adenine or no base (a frameshift) facing

Conformational changes in formamide residue

69 [2] Wallace S.S., Ide H., Structure/function relationships involved in the biological consequences of pyrimidine ring saturation and fragmentation products, Ionizing radiation damage to DNA: molecular aspects, Wiley-Liss, Inc, 1990, pp. 1–15. [3] Guy A., Duplaa A.M., Ulrich J., Téoule R., Incorporation by chemical synthesis and characterization of deoxyribosylformylamine into DNA, Nucleic Acids Res. 19 (1991) 5815–5820. [4] Falcone J.S., Box H.C., Selective hydrolysis of damaged DNA by nuclease P1, Biochem. Biophys. Acta 1337 (1997) 267–275. [5] Hevroni D., Livneh Z., Bypass and termination at apurinic sites during replication of single-stranded DNA in vitro: a model for apurinic site mutagenesis, Proc. Natl. Acad. Sci. USA 85 (1988) 5046–5050. [6] Boiteux S., Laval J., Coding properties of poly (deoxycytidylic acid) templates containing uracil or apyrimidinic sites: in vitro modulation of mutageness by deoxyribonucleic acid repair enzymes, Biochemistry 21 (1982) 6746–6751. [7] Loeb L.A., Preston B.D., Mutagenesis by apurinic/apyrimidic sites, Annu. Rev. Genet. 20 (1986) 201–230. [8] Cadet J., Nardin R., Voituriez L., Remin M., Hruska F.E., A 1H and 13 C nmr study of radiation-induced degradation products of 2’-deoxythymidine derivatives: N-(2’-deoxy-β-D-erythropentofuranosyl) formamide, Can. J. Chem. 59 (1981) 3313–3318. [9] Aurine J., Laplanche L.A., Rogers M.T., cis and trans Configurations of the peptide bond in N-monosubstitued amides by nuclear magnetic resonance, J. Am. Chem. Soc. 86 (1964) 337–341. [10] Bodenhausen G., Kogler H., Ernst R.R., Selection of coherencetransfer pathways in NMR pulse experiments, J. Magn. Res. 58 (1984) 370–388. [11] Piotto M., Saudek V., Sklénar V., Gradient-tailored excitation for single-quantum NMR spectroscopy of aqueous solutions, J. Biomol. NMR 2 (1992) 661–665.

Figure 4. Stereoviews of the models of the two duplexes. A. The duplex containing the trans isomer. The arrow indicates the formamide. B. The duplex containing the cis isomer.

formamide are under investigation to compare the structures with those obtained for an apurinic site and to discuss the biological implications. Acknowledgments We thank Marc Le Bret for the kind gift of the programs OCL and MORCAD. This work was, in part, supported by funds from the Ministère de l’Éducation Nationale et de l’Enseignement Supérieur, ACC-SV8.

References [1] Fuciarelli A.F., Zimbrick J.D., Radiation Damage in DNA, Battelle Press, Columbus, OH, 1995.

[12] Schmitz U., Zon G., James T.L., Deoxyribose conformation in (d(GTATATAC) 2: Evolution of sugar pucker by simulation of double-quantum-filtered COSY cross-peaks, Biochemistry 29 (1990) 2357–2368. [13] Arnott S., Smith P.J.C., Chandrasekaran R., Atomic coordinates and molecular conformations for DNA-DNA, RNA-DNA and DNA-RNA helices., Vol. 2, 3 edn., CRC Press, Cleveland, OH, 1976. [14] Cornell W.D., Cieplak P., Bayly C.I., Gould I.R., Merz K.M., Ferguson D.M., Spellmeyer D.C., Fox T., Caldwell G.W., Kollman P.A., A second generation force field for the simulation of proteins, nucleic acids, and organic molecules, J. Am. Chem. Soc. 117 (1995) 5179–5197. [15] Roll C., Ketterlé C., Faibis V., Fazakerley G.V., Boulard Y., Conformation of nicked and gapped DNA structure by NMR and molecular dynamic simulations in water, Biochemistry 37 (1998) 4059–4070. [16] Faibis V., Cognet J.A.H., Boulard Y., Sowers L.C., Fazakerley G.V., Solution structure of two mismatches G.G and I.I in the K-ras gene context by nuclear magnetic resonance and molecular dynamics, Biochemistry 35 (1996) 14452–14464. [17] Cuniasse P., Fazakerley G.V., Guschlbauer W., Kaplan B., Sowers L.C., The abasic site as a challenge to DNA polymerase, J. Mol. Biol. 213 (1990) 303–314.