Three-Dimensional Structure and Reactivity of a Photochemical Cleavage Agent Bound to DNA

Irradiation of the anthraquinone derivative N,N-Bis(3-aminopropyl)-2-anthraquinonesulfonamide dihydrochloride (AQS2) when bound to DNA leads to strand cleavage, primarily at GG steps. The 1.8 Å X-ray structure of AQS2 bound to the DNA duplex [d(CGTACG)]2 shows an intercalated, groove-reversed complex with the out-of-plane “swallow tail” located in the major groove. Two molecules of AQS2 bind with the chromophores intercalated at each CG step. Extensive guanine-AQS2 interactions are observed in the intercalated complex, indicating that the ground-state anthraquinone and DNA are poised for efficient electron transfer after photoexcitation. AQS2 engages in guanine-selective van der Waals contacts and molecular orbital overlap. Molecular orbitals, calculated using coordinates of the X-ray structure, confirm the extensive electronic overlap between the redox partners. The bases adjacent to the intercalation site are well-stacked. The structure supports a model in which electron transfer from base to AQS2 and hole migration within DNA require significant electronic overlap. DNA in its biological environment is oxidatively damaged by the products of normal metabolic processes,1 by ionizing radiation,2,3 and by light.4 One mechanism of these reactions involves the loss of an electron by a DNA base to form a radical cation (electron hole). Interest in such radical cations has expanded as a consequence of the discovery that their formation by electronically excited small molecules such as riboflavin,5 naphthalimide,6 rhodium complexes (Rh),7 and a series of anthraquinone (AQ) derivatives8 provides a model for natural oxidative processes. We showed that the DNA-binding mode of AQ derivatives is controlled by their structure. Most monosubstituted cationic anthraquinones intercalate; others appear to groove-bind.9 The binding mode determines the mechanism of reaction.10 Irradiation of intercalatively bound anthraquinones generates base radical cations. A radical cation can migrate from base to base along the DNA stack. Such hole migration through DNA is well-documented and well-accepted, but the rate of this process is still uncertain. Barton and co-workers have inferred an extremely rapid rate of migration;11 others contend that the rate is slower.12-14 The discrepant results may arise from differing extents of orbital interaction among electron acceptors and DNA bases in various experimental systems. Ultimately a migrating radical cation is stopped at a trap where it reacts irreversibly. The trapped radical cation causes strand cleavage.15 Our long-term goal is to determine how three-dimensional structures of DNA and DNA complexes direct efficiency and mechanism of photocleavage, initiating with photoexcitation, then DNA base to chromophore electron transfer, hole migration, trapping, and covalent oxidative damage. Here we describe the three-dimensional structure of the anthraquinone AQS2 bound to a DNA fragment determined by X-ray crystallography. The X-ray structure can help relate intercalation, extent of baseintercalator interaction, and base-base stacking to efficiency of electron transfer. AQS2 intercalates in DNA, interacting extensively with the DNA bases. Extensive overlap of molecular orbitals in the intercalated AQS2 complex helps explain the very rapid transfer of an electron to the excited quinone from its flanking DNA bases. The DNA bases adjacent to the intercalation site remain stacked, providing an efficient route for base to base hole migration. These results allow us to link the structure of the intercalated anthraquinone to the reaction of the DNA caused by its irradiation. † Current address: Department of Chemistry, Carnegie Mellon University, 4400 Fifth Ave., Pittsburgh, PA 15213. (1) Cheng, K. C.; Cahill, D. S.; Kasai, H.; Nishimura, S.; Loeb, L. A. J. Biol. Chem. 1992, 267, 166-172. (2) Boon, P. J.; Cullis, P. M.; Symons, M. C. R.; Wren, B. W. J. Chem. Soc., Perkin Trans. 2 1984, 1393-1399. (3) Wolf, P. G.; Jones, D. D.; Candeias, L. P.; O’Neill, P. Int. J. Radiat. Biol. 1993, 64, 7-18. (4) Melvin, T.; Plumb, M. A.; Botchway, S. W.; O’Neill, P.; Parker, A. W. Photochem. Photobiol. 1995, 61, 584-591. (5) Kasai, H.; Yamaizumi, Z.; Berger, M.; Cadet, J. J. Am. Chem. Soc. 1992, 114, 9692-9694. (6) Matsugo, S.; Kawanishi, S.; Yamamoto, K.; Sugiyama, H.; Matsuura, T.; Saito, I. Angew Chem., Int. Ed. Engl. 1991, 30, 1351-1353. (7) Arkin, M. R.; Stemp, E. D.; Pulver, S. C.; Barton, J. K. Chem. Biol. 1997, 4, 389-400. 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