Symposium: Applications of Inorganic Photochemistry
Symposium: Applications of Inorganic Photochemistry
Perfect Strangers: Inorganic Photochemistry and Nucleic Acids Pamela J. Carter, Suzanne A. Ciftan, Mark F. Sistare, and H. Holden Thorp* Department of Chemistry, University of North Carolina, Chapel Hill, NC 27599-3290 He was wedded to the unglamorous carbon cycle while younger [scientists] were achieving fame and opulent grants in such fair fields as neurobiology, virology, and the wonderful new wilderness of nucleic acids. John Updike, Couples, 1968.
In his early novel, Couples, John Updike describes Ken Whitman, a midcareer scientist who laments having devoted his scientific efforts to understanding the molecular events of photosynthesis rather than exploring seemingly more exciting fields, including nucleic acid chemistry. What Dr. Whitman (and by association, Mr. Updike) failed to realize was that photosynthesis and nucleic acid chemistry would eventually have an extensive common ground and that nearly thirty years later, the “wilderness of nucleic acids” would be even more new and certainly no less wonderful. The applications of photochemistry, particularly of transition-metal complexes, to nucleic acid chemistry have led to the development of numerous systems with exciting promise for solving important problems in molecular biology and biotechnology (1–4). The challenge now is for inorganic photochemists to wade deeper into the biological waters to utilize these systems in solving specific problems. In this article, we will provide a sampling of the approaches that have been developed (with some historical perspective) and a discussion of the challenges that lie ahead. We hope this latter discussion will highlight how inorganic photochemistry might be particularly suited to some of these challenges. Nucleic Acid Fundamentals There are numerous outstanding textbooks on nucleic acid chemistry (5). A brief summary of important features will be given here. Nucleic acids are polyanions comprised of nucleotide monomers that contain a ribose or deoxyribose sugar, a nitrogenous base, and an anionic phosphate linker (Fig. 1). The nitrogenous bases are guanine and adenine — the bicyclic purines— and cytosine and thymine (or uracil in RNA) — the monocyclic pyrimidines. The phosphodiester bond formed between the 5′-phosphate of the first monomer and the 3′ site of the second monomer creates a directional 5′–3′ linkage. Guanine and adenine pair to cytosine and thymine, respectively. GC base pairs with three hydrogen bonds are more stable than AT base pairs with only two hydrogen bonds. Single-base mismatches, base pairs where A ≠ T or G ≠ C, reduce duplex stability and are detected with enormous fidelity by the biological machinery responsible for DNA replication. Wobble and Hoogsteen base pairs, commonly found in RNA, are mismatches where one or more alternative hydrogen bonds are formed. Deliberate mismatching is found when base stacking is particularly favorable, as in tRNA.
Figure 1. Structures of the DNA nucleotides.
The spiral staircase form of double helices is created by the 5′–3′ strand running parallel to the 3′–5′ strand (the hand rails) with bases toward the inside (the stairs), as shown in Figure 2. Unlike an actual staircase, one can move up the double strand in two paths — either in the minor groove or in the major groove. The anionic phosphate–sugar backbones are much closer together in the minor groove, establishing a high negative charge density that is neutralized by electrostatically bound cations. Unlike its name, the minor groove plays a major role in binding biologically relevant proteins as well as man-made cationic drugs and nucleases. The major groove is wider and deeper with basepairing visible, and the nucleophilic N7 position of guanine is exposed for inner-sphere metal binding. Bases stack upon each other within one strand, analogous to stairs in a spiral staircase, increasing the helical stability. Base stacking is most favorable between the bicyclic purine bases. If the helix is partly unwound, planar drugs and nucleases can sandwich, or intercalate, between the stacked bases.
Figure 2. Structure of the DNA duplex showing the major and minor grooves.
*Corresponding author.
Vol. 74 No. 6 June 1997 • Journal of Chemical Education
641
Symposium: Applications of Inorganic Photochemistry A variety of double helical DNA structures are formed, depending on the solution conditions. The Watson–Crick model of right-handed B-form DNA is most stable at low salt concentrations; in high salt, the helix axis compresses into the A form. Z-DNA favors alternating GC sequences and high concentrations of divalent metal cations, converting the B form into an elongated, left-handed zigzag structure. Unlike DNA, the double helical structures of RNA are restricted to the A form. Probing DNA
Probing with Direct Emission Metal complexes bind to the DNA polyanion by electrostatic interactions, classical intercalation, or a combination of both (1, 3, 4). Intercalation involves the stacking of a ligand between adjacent base pairs of DNA, a strongly favored binding mode, with binding constants on the order of 106 M{1 (6, 7). This stacking interaction requires that the intercalating ligand be a flat, extended aromatic system, often with heterocyclic rings such as pyridine or pyrazine. Hypochromism in the absorption spectrum of the intercalator is observed upon DNA binding owing to the difference in polarity between the aqueous solution and the hydrophobic DNA core. Unwinding of the double helix also must occur to accommodate the intercalator within the stacked DNA bases. Complexes of ruthenium(II) containing the dppz (dipyrido[3,2-a:2′,3′-c]phenazine, ligands shown in Fig. 3) ligand are especially amenable to the study of doublestranded DNA (1, 8–12). When bound to DNA, the Ru → dppz metal-to-ligand charge transfer (MLCT) excited state displays strong emission. In aqueous solution, however, the emission is quenched by proton transfer from water to the phenazine nitrogen rings (13). Intercalation within the DNA double helix prevents water from interacting with the phenazine nitrogens, increasing the lifetime of the excited state. This “molecular light switch” effect observed with dppz complexes allows determination of binding characteristics by emission titration and Scatchard analysis. Because dppz complexes intercalate into DNA, they have been used to study long-range electron transfer mediated by the DNA (1, 14, 15); comparisons have been made with other intercalated and non-intercalated DNA-mediated long-range electron transfer systems (16, 17). Chromophore-quencher systems have also been used to observe DNA binding. The chromophore [fac-(bpy)Re(CO)3py-R]+, undergoes an Re→bpy MLCT transition at 340 nm (18). An anthracene group, covalently attached at R via a flexible linker, effectively quenches the excited state when the complex is free in solution (19). Intercalation of the anthracene places steric constraints on the geometry of the linker group. The required geometry in the intercalated complex is less favorable to the quenching interaction, causing luminescence to return upon intercalation. Inorganic photochemistry can also be applied to characterize complexes containing nucleobase ligands. The photophysical properties of Re(bpy)(CO)3(CH 3CN)+ change dramatically upon displacement of the acetonitrile group with EtG (9-ethylguanine), which binds to the metal via the nucleophilic N7 position (20). The MLCT absorption is redshifted by 2400 cm{1, and the emission maximum is redshifted by 2030 cm{1. The EtG complex has a long-lived excited state, allowing an energy gap law analysis, which relates the lifetime of the excited state to the emission energy. This analysis provides a method to characterize the electronic properties of the coordinated ligand and shows EtG to be a good π-donor, similar to electron-rich pyridines.
642
Figure 3. Structures of ligands.
Also obtained from this analysis is a strategy for designing nucleobase-targeting complexes — electron-poor complexes should bind favorably to guanine in DNA. Electrochemical stimulation can also be used to generate emissive excited states that interact with DNA via a process known as electrogenerated chemiluminescence (ECL). For example, Ru(phen) 32+ is oxidized to the +3 form at approximately 1.0 V (vs. Ag/AgCl), and oxalic acid reacts with the electrochemically generated Ru(phen)3 3+ to form the CO2{· anion (eq 2). The CO2{· radical reduces additional Ru(phen)33+ to generate emissive Ru(phen)32+*. The CO2{· also reduces the ground state Ru(phen)32+ (eq 4); the resultant Ru(phen) 3 + reacts with Ru(phen) 3 3+, also generating Ru(phen) 3 2+ * (eq 6). The reactions that produce the Ru(phen)32+* excited state are severely hindered by DNA, because the polyanion prevents the CO2{· radical from reducing the Ru(phen)33+ (21). This process permits determination of the binding constant of Ru(phen)32+ to DNA using ECL. Ru(II) → Ru(III) + e{
(1)
Ru(III) + C2O42{ → Ru(II) + C2O4{·
(2)
C2O4{· → CO2 + CO2{·
(3)
Ru(II) + CO2 → Ru(I) + CO 2
(4)
Ru(III) + CO2{· → Ru(II)* + CO2
(5)
Ru(III) + Ru(I) → Ru(II)* + Ru(II)
(6)
Ru(II)* → Ru(II) + hν
(7)
{·
The ECL reactions can also be used to detect the presence of double-stranded DNA (22, 23). Although Ru(phen)3 2+ binds strongly to double-stranded DNA, minimal binding is observed in the presence of single-stranded DNA (24). The emissive excited state Ru(phen)3 2+* can also be generated by reacting Ru(phen)33+ with a neutral electron donor, such as tri-n-propylamine, by analogy to eq 5. Using a neutral electron donor eliminates the electrostatic repulsion that retards eq 5 when CO2 {· is the electron donor. A sensor for a given DNA sequence can thus be designed by immobilizing its complementary sequence to an electrode (22). Treatment of the electrode with the target DNA will generate double-
Journal of Chemical Education • Vol. 74 No. 6 June 1997
Symposium: Applications of Inorganic Photochemistry
Figure 4. Mechanisms of strand scission following activation of the 1', 3', 4', and 5' C–H bonds in DNA.
stranded DNA, allowing binding of the ruthenium complex. After rinsing away unbound Ru(phen)3 2+, addition of tri-npropylamine and application of electrochemical potential will yield emission only if the target DNA is present to form double-stranded DNA (23).
Probing with Emission Quenching DNA binding affinities can be measured for metal complexes by quenching the excited state emission of Pt2(pop)44{ (4), which, because of the tetraanionic charge, does not bind to DNA. This technique is based on the regulation of the amount of cationic metal quencher free in solution owing to DNA binding. There are three possible scenarios: (i) only Pt2 (pop)4 4{ is present and the highest emission intensity is observed; (ii) metal quencher is added and the emission intensity decreases dramatically; or (iii) DNA and quencher are present and the emission intensity falls between the first and second scenarios because some quencher is bound to DNA. Therefore, when DNA is titrated into a solution of Pt2 (pop) 44{ and metal quencher, more metal complex is bound to the DNA and less is free in solution for collisional quenching with the tetraanionic complex. The binding constants for low-affinity (100 M{1) and high-affinity (107 M{1) metal complexes of various metals of Ru, Rh, Co, and Os have been determined using this approach (25). The precision of the method is high enough to discern the effects of ionic strength at sufficient resolution to ascribe changes in binding to those predicted by polyelectrolyte theory (26). Modifying DNA
Hydrogen Abstraction The ability of transition metal complexes to oxidize substrates such as alcohols or hydrocarbons via hydrogen atom transfer has been a focus of inorganic chemists for some time (27–29). The sugar moiety of nucleotides also contains several activated C–H bonds, which makes it a good candidate for oxidation by hydrogen-atom abstraction (30, 31). Upon hydrogen atom abstraction from the sugar, the DNA or RNA becomes cleaved to yield diverse products depending upon the site of oxidation; the smaller products such as free or modified bases and modified sugar groups can be detected by HPLC, while the longer nucleic acid frag-
ments can be visualized by high-resolution electrophoresis. The mechanisms of strand scission following breakage of the 1′, 3′, 4′, and 5′ C–H bonds are shown in Figure 4. In the cases discussed here, the transition metal complex must first be activated by photolysis. Numerous other reagents react thermally or act in a catalytic fashion upon addition of a terminal oxidant (30, 31). The advantage of the photochemical approach is that the excited-state lifetime of the oxidant provides a lower limit for the rate of the hydrogen abstraction, which is generally much faster than binding and recognition processes. Therefore, the cleavage pattern for photochemical nucleases usually reflects faithfully the binding pattern (or lack thereof) of the oxidant. A range of complexes containing d 6 metals such as Rh(III) or Co(III) have been explored as DNA photocleavage agents (3, 32). Among these compounds are the phenanthrenequinone diimine (phi) complexes of Rh(III), which undergo a ligand-centered π → π* transition in the near-UV region. These Rh(III) complexes bind tightly to DNA via intercalation of the hydrophobic phi ligand between adjacent base pairs (33) and produce strand scission at the site of binding upon photolysis. The cleavage mechanism involves oxidation of the deoxyribose at the 3′ sugar position to yield oligomers containing 3′- and 5′-phosphate termini, a free base, and a sugar fragment (32). The mechanism on the metal complex side of the reaction probably involves hydrogen abstraction by the coordinated phi ligand. Complexes of Co(III) also undergo light-induced reactions that result in DNA cleavage. Both Co(phen)33+ and Co(DiP)33+ (DiP = diphenylphenanthroline) produce singlestranded nicks in plasmid DNA, as monitored by agarose gel electrophoresis (34). The Co(III) form of bleomycin (BLM), a glycopeptide-derived antibiotic, is another example of a metal-containing photocleavage reagent (31, 35). Although this compound is not as widely understood as its Fe(II)–BLM counterpart, studies done on model complexes have suggested that irradiation of Co(III)-BLM in aqueous solution generates ·OH, which might be responsible for DNA damage (36). Nucleic acids, particularly RNA, assume diverse conformations and present regions of structural distortions that are important for protein recognition (5). Understanding protein–nucleic acid interactions is essential for understanding gene expression and could aid in the design of novel chemotherapeutic agents. Transition metal complexes that cleave DNA or RNA can provide valuable information about protein–nucleic acid interactions in two ways. First, the complex may be designed to recognize DNA structural elements via hydrogen bonding or van der Waals interactions in a manner that mimics recognition by proteins (37). This approach has been pioneered by the Barton group; it is described in numerous review articles (2, 3), and therefore not treated in detail here. At the heart of this approach is the ability to determine the sites of DNA or RNA cleavage by high-resolution gel electrophoresis. A simple explanation of this technique is provided in recent reviews (3, 4). The second approach to using inorganic photochemistry to understand protein–nucleic acid interactions involves complexes that cleave DNA or RNA in a sequence-independent manner. A technique referred to as protein footprinting involves cleaving sequences independently with and without a protein that binds to a particular nucleic acid (38). In this experiment, some nucleotides are protected from cleavage by binding of the protein; this protected region is referred to as the “footprint” (Fig. 5). A thermal approach to footprinting has been developed by generating ·OH with Fe(EDTA)2{ (39, 40) (EDTA = ethylenediamine tetraacetic acid); a photochemical approach has been developed with
Vol. 74 No. 6 June 1997 • Journal of Chemical Education
643
Symposium: Applications of Inorganic Photochemistry product with decreased mobility during gel electrophoresis, indicating the formation of a DNA–metal adduct in high yield. Adduct formation appears to be specific to guanine sites, implicating strongly a mechanism in which the initial step is an electron transfer process (48). Recent studies show that long-wavelength irradiation of Rh(III) complexes leads to base-labile scission due to electron abstraction from guanine (49). Perspectives
Figure 5. Summary of footprinting technique, adapted from ref 38 .
one of the most widely studied photoreactive complexes: Pt2 (pop)4 4{ (pop = P2O 5H 22{) (29). The complex is a d 8–d 8 dimer that consists of two square-planar platinum centers bridged by four pyrophosphito ligands. Upon irradiation with 367-nm light, this complex undergoes a dσ* → pσ transition to generate a long-lived (τ = 9.5 µs) excited state that is a hydrogen abstractor for a variety of substrates including DNA (29, 41, 42). In duplex DNA, Pt2(pop)4 4{ cleaves DNA by abstracting a hydrogen from both the 4′ and 5′ ribose carbons, as illustrated in Figure 4 (43). Cleavage at these sites results in frank scission that is immediately apparent on a sequencing gel; some additional cleavage is observed after treatment with piperidine. Because Pt2(pop)44{ is anionic, it does not bind to DNA and therefore exhibits a sequenceneutral cleavage pattern. A high resolution footprint of the λ repressor protein bound to its OR 1 operator sequence has been obtained using Pt2(pop)44{ (43). The footprint is similar to that obtained with ?OH (39, 40), and both reagents provide footprints that are high enough in resolution to distinguish binding of the protein to a single side of the DNA duplex. This discrimination is possible when two separate half-sites of protection are observed because of the coiling of the DNA strands. An advantage of the diplatinum complex is that it does not produce a diffusible intermediate such as ?OH, which is easily quenched by reagents commonly used to maintain the native form of the protein, such as DTT or glycerol. The complex Rh(phi)2(bpy)3+ also exhibits a sequence-neutral photocleavage pattern and has been used to footprint the binding of both EcoRI and distamycin on a restriction fragment (44).
Electron Transfer The MLCT excited state of ruthenium polypyridyl complexes has greater oxidizing and reducing power than the corresponding ground state (45). Two photoreactions are known between ruthenium polypyridyls and DNA: base modification and photoadduct formation. Excited Ru(bpy)3 2+ can excite ground-state 3 O2 to 1O2 , which attacks the guanine base to form 8-oxoguanine. Subsequent treatment of the oxidized DNA with piperidine causes loss of the modified base and strand cleavage (46). The complex Ru(TAP)3 2+ directly oxidizes GMP and AMP by one electron upon photolysis (47). The photoreaction between Ru(TAP) 32+ or Ru(TAP)2(bpy) 2+ and labeled oligonucleotides generates a
644
Using inorganic photochemistry to understand the structures of nucleic acids and nucleoprotein complexes is more important than ever. This point is illustrated by the proliferation of new sequences afforded by the SELEX technique (50). In SELEX, a random sequence of DNA is synthesized across a given number of nucleotides. For example, if a random 100-mer is prepared, then there are 4100 possible sequences, of which as many as 1015 might be obtained following the preparation on a conventional synthesizer. Using a combination of PCR and affinity chromatography, the best of these sequences are selected based on binding to a given target. The individual sequences are then determined by cloning into bacteria. Related strategies are possible for RNA by introducing reverse transcription steps. These sequences provide lead compounds for inhibition of the target of interest. Therefore, pharmaceutical chemists have the ability to generate leads for any target that can be immobilized and screened against the random library. The SELEX process therefore generates mind-boggling numbers of novel nucleic acids and nucleoprotein complexes at a high rate. Once the lead sequence is obtained, many challenges for preclinical drug development are immediately presented. If the goal is to use an actual nucleic acid as a drug, the first goal is to obtain the minimum necessary sequence for binding to the target, and footprinting provides this information rapidly. If the goal is to use the nucleic acid as a scaffold from which to design small organic molecules that produce similar binding (51), then even more information is required. In this latter case, both the threedimensional structure of the nucleic acid and the nature of the nucleic acid–protein contacts are important. Here, information is needed on the secondary and tertiary structure of the nucleic acid in the absence of the protein and on the precise nucleotides that contact the protein. This information can serve as an indicator of whether more detailed structural studies are likely to be productive. It seems likely that inorganic photochemistry will be part of this nucleic acid-based drug discovery. Acknowledgment We thank the National Science Foundation and the David and Lucile Packard Foundation for supporting our work in this area. Literature Cited 1. Arkin, M. R.; Jenkins, Y.; Murphy, C. J.; Turro, N. J.; Barton, J. K. In Mechanistic Bioinorganic Chemistry; Thorp, H. H.; Pecoraro, V. L., Eds.; Advances in Chemistry 246; American Chemical Society: Washington, DC, 1995; pp 449–469. 2. Chow, C. S.; Barton, J. K. Meth. Enzymol. 1992, 212, 219–241. 3. Pyle, A. M.; Barton, J. K. Prog. Inorg. Chem. 1990, 38, 413. 4. Thorp, H. H. Adv. Inorg. Chem. 1995, 43, 127–177. 5. Nucleic Acids in Chemistry and Biology; Blackburn, G. M.; Gait, M. J., Eds.; IRL: Oxford, 1990. 6. Waring, M. J. J. Mol. Biol. 1965, 13, 269.
Journal of Chemical Education • Vol. 74 No. 6 June 1997
Symposium: Applications of Inorganic Photochemistry 7. Lippard, S. J. Acc. Chem. Res. 1978, 11, 211–217. 8. Friedman, A. E.; Chambron, J. C.; Sauvage, J. P.; Turro, N. J.; Barton, J. K. J. Am. Chem. Soc. 1990, 112, 4960. 9. Jenkins, Y.; Barton, J. K. J. Am. Chem. Soc. 1992, 114, 8736– 8738. 10. Gupta, N.; Grover, N.; Neyhart, G. A.; Liang, W.; Singh, P.; Thorp, H. H. Angew. Chem. Int. Ed. Engl. 1992, 31, 1048. 11. Welch, T. W.; Corbett, A. H.; Thorp, H. H. J. Phys. Chem. 1995, 99, 11757–11763. 12. Schoch, T. K.; Hubbard, J. L.; Zoch, C. R.; Yi, G.-B.; Sørlie, M. Inorg. Chem. 1996, 35, 4383–4390. 13. Turro, C.; Bossmann, S. H.; Jenkins, Y.; Barton, J. K.; Turro, N. J. J. Am. Chem. Soc. 1995, 117, 9026–9032. 14. Murphy, C. J.; Arkin, M. R.; Jenkins, Y.; Ghatlia, N. D.; Bossmann, S. H.; Turro, N. J.; Barton, J. K. Science 1993, 262, 1025–1029. 15. Murphy, C. J.; Arkin, M. R.; Ghatalia, N. D.; Bossman, S.; Turro, N. J.; Barton, J. K. Proc. Natl. Acad. Sci. USA 1994, 91, 5315– 5319. 16. Meade, T. J.; Kayyem, J. F. Angew. Chem. Int Ed. Engl. 1995, 34, 352–354. 17. Brun, A. M.; Harriman, A. J. Am. Chem. Soc. 1994, 116, 10383– 10393. 18. Schanze, K. S.; MacQueen, D. B.; Perkins, T. A.; Cabana, L. A. Coord. Chem. Rev. 1993, 32, 63. 19. Thornton, N. B.; Schanze, K. S. Inorg. Chem. 1993, 32, 4994– 4995. 20. Oriskovich, T. A.; White, P. S.; Thorp, H. H. Inorg. Chem. 1995, 34, 1629–1631. 21. Carter, M. T.; Bard, A. J. Bioconjugate Chem. 1990, 1, 257. 22. Xu, X.-H.; Yang, H. C.; Mallouk, T. E.; Bard, A. J. J. Am. Chem. Soc. 1994, 116, 8386–8387. 23. Xu, X.-H.; Bard, A. J. J. Am. Chem. Soc. 1995, 117, 2627–2631. 24. Welch, T. W.; Thorp, H. H. J. Phys. Chem. 1996, 100, 13829– 13836. 25. Kalsbeck, W. A.; Thorp, H. H. J. Am. Chem. Soc. 1993, 115, 7146– 7151. 26. Kalsbeck, W. A.; Thorp, H. H. Inorg. Chem. 1994, 33, 3427– 3429. 27. Meyer, T. J. J. Electrochem. Soc. 1984, 131, 221C.
28. Meunier, B. Chem. Rev. 1992, 92, 1411. 29. Roundhill, D. M.; Gray, H. B.; Che, C.-M. Acc. Chem. Res. 1989, 22, 55. 30. Pratviel, G.; Bernadou, J.; Meunier, B. Angew. Chem. Int. Ed. Engl. 1995, 34, 746–769. 31. Stubbe, J.; Kozarich, J. W. Chem. Rev. 1987, 87, 1107. 32. Sitlani, A.; Long, E. C.; Pyle, A. M.; Barton, J. K. J. Am. Chem. Soc. 1992, 114, 2303. 33. David, S. S.; Barton, J. K. J. Am. Chem. Soc. 1993, 115, 2984. 34. Barton, J. K.; Raphael, A. L. J. Am. Chem. Soc. 1984, 106, 2466– 2468. 35. Chang, C. H.; Meares, C. F. Biochemistry 1982, 21, 6332. 36. Tan, J. D.; Hudson, S. E.; Brown, S. J.; Olmstead, M. M.; Mascharak, P. K. J. Am. Chem. Soc. 1992, 114, 3841–3853. 37. Krotz, A. H.; Hudson, B. P.; Barton, J. K. J. Am. Chem. Soc. 1993, 115, 12577–12578. 38. Papavassiliou, A. G. Biochemistry 1995, 305, 345–357. 39. Tullius, T. D.; Dombroski, B. A.; Churchill, M. E. A.; Kam, L. Meth. Enzymol. 1987, 155, 537–558. 40. Tullius, T. D.; Dombroski, B. A. Proc. Natl. Acad. Sci. USA 1986, 83, 5469–5473. 41. Kalsbeck, W. A.; Grover, N.; Thorp, H. H. Angew. Chem. Int. Ed. Engl. 1991, 30, 1517. 42. Kalsbeck, W. A.; Gingell, D. M.; Malinsky, J. E.; Thorp, H. H. Inorg. Chem. 1994, 33, 3313–3316. 43. Breiner, K. M.; Daugherty, M. A.; Oas, T. G.; Thorp, H. H. J. Am. Chem. Soc. 1995, 117, 11673–11679. 44. Uchida, K.; Pyle, A. M.; Morii, T.; Barton, J. K. Nucleic Acids Res. 1989, 17, 10259–10279. 45. Kalyanasundaram, K. Coord. Chem. Rev. 1982, 46, 159. 46. Mei, H.-Y.; Barton, J. K. Proc. Natl. Acad. Sci. USA 1988, 85, 1339–1343. 47. Lecomte, J.-P.; Kirsch-De Mesmaeker, A.; Feeney, M. M.; Kelly, J. M. Inorg. Chem. 1995, 34, 6481–6491. 48. Jacquet, L.; Kelly, J. M.; Kirsch-De Mesmaeker, A. J. Chem. Soc., Chem. Commun. 1995, 913–914. 49. Hall, D. B.; Holmlin, R. E.; Barton, J. K. Nature 1996, in press. 50. Gold, L. J. Biol. Chem. 1995, 270, 13581–13584. 51. Kenan, D. J.; Tsai, D. E.; Keene, J. D. Trends Biochem. Sci. 1994, 19, 57–64.
Vol. 74 No. 6 June 1997 • Journal of Chemical Education
645
