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Bioconjugate Chem. 1999, 10, 1122−1130
Synthesis of Metallointercalator-DNA Conjugates on a Solid Support R. Erik Holmlin, Peter J. Dandliker,† and Jacqueline K. Barton* Division of Chemistry and Chemical Engineering, California Institiute of Technology, Pasadena, California 91125. Received June 21, 1999; Revised Manuscript Received August 26, 1999
Metallointercalator-DNA conjugates were prepared by amide bond formation between active esters on the nonintercalating ligands of transition metal complexes and primary amines presented at the 5′ or the 3′ termini of oligonucleotides attached to solid supports. The conjugates were liberated from the support by aminolysis and purified by HPLC on C18 or C4 stationary phases, which separates the two diastereomeric forms of the conjugates containing either the Λ or the ∆ enantiomer of the octahedral metal complex. The coupling reaction proceeds with ∼75% conversion of the aminoterminated oligonucleotide into the conjugate; the isolated yield is ∼200 nmol for syntheses initiated on DNA-synthesis columns with a loading of 2 µmol. The conjugates were characterized by ultravioletvisible and circular dichorism absorption spectroscopy, electrospray ionization mass spectrometry, enzymatic digestion, and polyacrylamide gel electrophoresis (PAGE). Oligonucleotides bearing [Rh(phi)2(bpy′)]3+ (phi ) 9,10-phenanthrene quinone diimine; bpy′ ) 4-butyric acid-4′-methyl bipyridyl) form 1:1 duplexes with the complementary strand, and the electrophoretic mobility under nondenaturating PAGE of duplexes containing ∆-Rh is notably different from duplexes containing Λ-Rh. Highresolution PAGE of DNA photocleavage reactions initiated by irradiation of the tethered Rh complexes reveal intercalation of the complex only near the tethered end of the duplex. Analogous DNA-binding properties were observed with [Rh(phi)2(bpy′)]3+ tethered to the 3′ terminus. By combining the 3′ and 5′ modification strategies, a mixed-metal DNA conjugate containing both [Os(phen)(bpy′)(Me2-dppz)]2+ (Me2-dppz ) 7,8-dimethyldipyridophenazine) on the 3′ terminus and [Rh(phi)2(bpy′)]3+ on the 5′ terminus was prepared and isolated. Taken together, these strategies for preparing metallointercalator-DNA conjugates offer a useful approach to generate chemical assemblies to probe long-range DNA-mediated charge transfer where the redox initiator is confined to and intercalated in a welldefined binding site.
INTRODUCTION
Investigations of long-range electron transfer (ET) reactions mediated by the stacked, heterocyclic bases of DNA are important to fundamental studies of ET, as well as the complete biochemical characterization of the genetic material. In our laboratories we employ transition metal complexes that bind to DNA by intercalation, metallointercalators, with well-defined redox properties to probe DNA-mediated electron transfer (1). Here we describe synthetic methods to prepare oligonucleotide conjugates in which a metallointercalator, [Rh(phi)2(bpy′)]3+ (Figure 1; phi ) 9,10-phenanthrene quinone diimine; bpy′ ) 4-butyric acid-4′-methyl bipyridyl), is attached covalently to DNA at either the 5′ or 3′ terminus while the assembly is immobilized on a solid support. Photoinduced ET between metallointercalators bound to DNA is remarkably fast and efficient (2-5). In titrations of electron acceptor (∆-[Rh(phi)2(bpy)]3+) into solutions of electron donor (∆-[M(phen)2(dppz)]2+) bound to DNA, 50% of the donors were quenched by just 1 equiv of ∆-[Rh(phi)2(bpy)]3+ on a time scale of 85%
Synthesis of Metallointercalator−DNA Conjugates
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Figure 3. Circular dichroism of the two rhodium-containing products (∆-Rh-151, Λ-Rh-151) isolated from the reaction mixture shown in Figure 2.
Figure 2. HPLC chromatogram of an Rh-DNA conjugate reaction mixture before purification (upper panel). UV-vis absorption spectra of the two rhodium-containing fractions at elution times of 28 min (left) and 34 min (right) (lower panel). The sequence of the oligonucleotide was as follows: 5′-Rh-ACG ATA TAC TAA CGT-3′.
yield and can be followed by reversed-phase HPLC (Supporting Information). At this stage, the oligonucleotide is poised for reaction with an electrophile. We exploited simple, high-yield amide bond formation to tether the metal complex to the functionalized oligonucleotide on the solid support. The carboxylate of [Rh(phi)2(bpy′)]3+ was converted to the NHS ester by treatment with TSTU. TSTU was used to synthesize an active ester with a derivative of [Ru(phen)3]2+ for conjugation to amino-modified DNA in solution (28-30), and in our work we favored TSTU because we could follow the activation of the carboxylate by TLC, and isolate the NHS ester for characterization. A matrix-assisted laser-desorption mass spectrum of the NHS ester of [Rh(phi)2(bpy′)]3+ is provided in the Supporting Information. We executed the coupling reaction by combining the solution of the NHS ester of [Rh(phi)2(bpy′)]3+ (without isolation) and the CPG beads coated with the aminomodified DNA and stirring the slurry overnight. Owing to the tendency of the conjugate attached to the bead to be well solvated by organic solvents, we observed significant solvatochromism in the coloration of the CPG beads after the coupling reaction, which is consistent with incorporation of the transition metal chomophore. The final step in this procedure is to liberate simultaneously the conjugate from the bead and remove the protecting groups by treatment with aqueous ammonia. The conjugates are purified by reversed-phase HPLC under the same conditions employed for purification of unmodified oligonucleotides. We estimate the yield of the coupling step to be ∼75% by comparing the amount of amino-modified DNA present (relative to the unmodified strand as an internal standard) before and after coupling. A similar procedure was employed to attach a metallointercalator containing ruthenium to the 5′ terminus of oligonucleotides in a study of long-range oxidative damage to guanine in DNA duplexes (9). (B) Isolation of of 5′-Rh-Modified Oligonucleotides by HPLC. Figure 2 shows a typical chromatogram of an unpurified reaction mixture for the synthesis
of 5′-Rh-modified DNA monitored at two wavelengths: 260 nm where everything in the mixture absorbs and 390 nm where only the rhodium complex absorbs. This reaction yields two products that contain rhodium. Owing to the hydrophobicity added by the metallointercalator, the conjugates elute with later retention times compared to the unmodified DNA, which simplifies their purification. The absorption spectra of the two rhodium-containing products (Figure 2) are identical and consistent with expectation for Rh-modified DNA. The intense absorption centered at 260 nm reflects the contribution of the DNA bases, while the absorption centered at 390 nm corresponds to absorption of the metal complex. The transition centered at 390 nm is hypochromic compared to that for the metal complex free in solution. Such hypochromicity for the conjugate indicates that the metal complex interacts preferentially with the bases of the singlestranded DNA (20, 24). The HPLC chromatogram in this figure reveals several side products that do not contain a rhodium complex. Since we find that the amounts of these species vary considerably from one synthesis to the next, we suggest that they likely reflect side products and failed sequences in the DNA synthesis. It is important to note that the relative amount of these products is somewhat exaggerated by the fact that the Rh-modified DNA elutes as two peaks, so the relative height of the product peak is reduced by a factor of 2 while the height of the peaks due to the side products are constant. (C) Circular Dichroism Spectroscopy of RhModified DNA. To answer the question of why two products with identical absorption spectra are formed, we measured the circular dichroism (CD) spectra of each species (Figure 3). On the basis of the equal and opposite relationship of the spectra in this region where the metal complex absorbs and by comparison to CD spectra for the enantiomers of the parent [Rh(phi)2(bpy)]3+, we concluded that into one product we had incorporated the ∆ enantiomer of [Rh(phi)2(bpy′)]3+, while we introduced Λ-[Rh(phi)2(bpy′)]3+ into the other. Remarkably, achiral HPLC was the only purification method required to separate these two diastereomeric Rh-DNA conjugates. Moreover, neither diastereomer was formed preferentially. (D) Electrospray Ionization Mass Spectrometry of Rh-DNA Conjugates. We used electrospray ionization (ESI) MS to measure the mass of Rh-modified oligonucleotides. ESI MS can be used to confirm not only that the conjugate is intact, but also it confirms that all of the protecting groups have been eliminated (31). Figure 4 shows the ESI MS of [Rh(phi)2(bpy′)]3+ attached by a diaminononane linker to the 5′-hydroxyl of a 20 nt oligonucleotide. The data shown is a deconvoluted spectrum constructed from mass:charge (m/z) data for several charge states (31). The calculated mass for the
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Figure 5. HPLC chromatograms monitored at 260 nm of the enzymatic digestion of an unmodified 14-mer (top) and a Rhmodified 14-mer (bottom). The area under the peak for dA is clearly reduced in the data for the conjugate. This loss of dA is consistent with the coupling of the metallointercalator to the 5′ terminus of this oligonucleotide.
Figure 4. Electrospray ionization (ESI) mass spectrum of [Rh(phi)2(bpy′)]3+ tethered to a 20-mer oligonucleotide. The calculated mass for this conjugate is 6964.8 amu and the observed mass is 6965.0. Scheme 2. Schematic Illustration of the Enzymatic Digestion of Oligonucleotides by Snake Venom Phosphodiesterase To Give Nucleotide Monophosphates (NMPs)a
a Hydrolysis of NMPs by alkaline phosphatase gives the nucleotides. The 5′-terminal base remains attached to the metallointercalator because the enzymes do not hydrolyze the carbamate linkage. The products are separated by HPLC and the numbers of each base are counted.
conjugate is 6964.8, and the observed mass is 6965.0. From this spectrum we can say (i) the product is the desired conjugate and (ii) all of the protecting groups are remove upon treatment with aqueous ammonia without degredation of the metal complex or the DNA. (E) Enzymatic Digestion of Rh-Modified DNA to Establish Base Content. To establish that covalent modification occurred at the targeted terminus as designed, we determined the base content of Rh-modified DNA and compared the results to data for unmodified DNA. The premise of this assay, outlined in Scheme 2, is to detect the modification of the 5′-terminal base. The oligonucleotide is treated with a cocktail of snake venom phosphodiesterase (SVP) and alkaline phosphatase (AP) (32). SVP cleaves the phosphodiester bonds of the oligonucleotide to release the nucleotide monophosphates (NMPs). AP then hydrolyzes the NMPs to give the neutral nucleotides, which are significantly more amenable to chromatographic separation. The reaction mix-
ture is then analyzed by reversed-phase HPLC. Since the enzymes do not recognize the carbamate linkage that joins the nucleotide and the metal complex through an aliphatic linker, the modified DNA will appear to have lost one base in the HPLC analysis. Figure 5 shows the results of the digestion of two oligonucleotides: an unmodified 14-mer and the corresponding Rh-modified 14-mer. For this conjugate, the rhodium complex is attached to a deoxyadenosine (dA). The quantity of each base, determined by integration of the chromatograms, is consistent with expectation based on the known sequence. The area under the peak corresponding to dA is clearly diminished for the conjugate compared to the unmodified strand. These results establish that the conjugation reaction is selective for the 5′ terminus and that no other bases are modified. Following the digestion, the metal complex attached to the 5′ terminal nucleoside adsorbs to the filter before analysis and therefore does not appear in the chromatogram in Figure 5. Synthesis of Rh-Modified Oligonucleotides with the Metallointercaltor Attached to the 3′ Terminus. To generalize our method of generating metallointercalator-DNA conjugates, we developed a procedure to modify the 3′ terminus of oligonucleotides (Scheme 3). As with conjugation to the 5′-hydroxyl, this approach relies on derivatization of the strand on the CPG resin. A commercially available CPG bead functionalized with a hydroxy aminoalkane masked with Fmoc and dimethoxy trityl (DMT) protecting groups was used as the support for automated DNA synthesis. Following selective removal of the DMT group, the DNA was synthesized by the standard phosphoramidite approach to give an oligonucleotide functionalized at the 3′ terminus with an Fmoc protected, alkylamine while the assembly remains attached to the solid support. The amine is selectively unmasked by treatment with piperidine in DMF without affecting the protecting groups on the DNA bases or backbone (23). The oligonucleotide is then coupled to the NHS ester of [Rh(phi)2(bpy′)]3+ as described for the 5′-modification strategy. Before removing the product from the bead, the DMT protecting group is removed from the 5′-hydroxyl by treatment with trichloroacetic acid in acetonitrile. Oligonucleotide conjugates prepared by this procedure are also purified by HPLC and can be resolved into the ∆and Λ-rhodium-containing diastereomers (33).
Synthesis of Metallointercalator−DNA Conjugates Scheme 3. Coupling of [Rh(phi)2(bpy′)]3+ to the 3′ Terminus of Oligonucleotides Attached to a Solid Supporta,b
a Reagents: (a) Automated DNA synthesis (ABI 391 DNA synthesizer); (b) piperidine (20% v/v), DMF, rt, 30 min; (c) TSTU, DIEA, CH3OH, CH3CN, CH2Cl2 (1:1:1), rt, 1 h; (d) heterogeneous coupling, DIEA, rt, 12 h; (e) TCAA, CH3CN, 18 s, rt. b Abbreviations: CPG ) controlled pore glass; DMT ) dimethoxy trityl; Rh ) [Rh(phi)2(bpy′)]3+.
Figure 6. Nondenaturing polyacrylamide (20%) gel for following the hybridization of 32P-end-labeled complement to Rhmodified DNA. Contents of lanes: (1) labeled complement (single stranded); (2) unmodified duplex (complement plus rhodium strand; 1:1); (3-7) labeled complement (8 µM) plus increasing concentrations of ∆-Rh-DNA (6, 7, 8, 9, 10 µM). Lanes 8-14 are the same as 1-7, with Λ-Rh substituted for the ∆ isomer.
Characterization of Rh-Modified DNA Duplexes. (A) Monitoring the Hybridization of Rh-Modified Oligonucleotides to Their Complements by PAGE. We followed the hybridization of 32P-end-labeled DNA to Rh-modified DNA on nondenaturing polyacrylamide gels (free of urea). Owing to the stiffness of duplex DNA relative to single strands, the duplexes are retarded relative to single stranded DNAs under electrophoresis on polyacrylamide gels. Figure 6 illustrates a typical hybridization experiment for Λ-Rh-DNA and ∆-Rh-DNA (25). The first lane of each titration contains only the 32P-end-labeled complement of the Rh-modified strand ([32P]Rh-DNA-C). The second lane contains [32P]Rh-DNA-C hybridized to 1 equiv of the unmodified rhodium strand. As expected for the formation of a 1:1 complex, a new band appears in the gel retarded relative to the single strand and there is no [32P]Rh-DNA-C remaining. The next five lanes contain [32P]Rh-DNA-C (8 µM) and increasing concentrations of RhDNA (6, 7, 8, 9, and 10 µM). As in the case of the unmodified duplex, a new band appears near the top of the gel. The intensity of this band increases with RhDNA until it reaches saturation at 8 µM. Similarly, the band for [32P]Rh-DNA-C decreases in intensity until disappearing from the gel when the sample is fully duplexed. There is an exogenous source of radioactivity
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present at equal levels in the sample that is present in all lanes and migrates just above the single strand. It is noteworthy that the Rh-modified duplexes migrate above the unmodified duplex. Such an observation is consistent with the positively charged metal complex reducing the total negative charge of the assembly and diminishing its attraction to the positive pole of the gel (25). Upon comparison of migration profiles of the ∆ and Λ-Rh-modified duplexes, the diastereomer containing Λ-Rh moves slower on the gel than the diastereomer containing ∆-Rh. Owing to the symmetry matching of the right-handed DNA duplex with the right-handed ∆-Rh, we expect the complex to be intercalated more deeply into the duplex than the Λ-isomer (34). The migration behavior observed in the gels is consistent with this notion, since a more compact structure should be less hindered by the gel, as is the duplex bearing ∆-[Rh(phi)2(bpy′)]3+. In the lanes for duplexes with ∆-Rh, there is a faint band that migrates just ahead of the major band for the doublestranded conjugates; this species may reflect a small population of assemblies where the complex is intercalated at a site other than the predominant binding site. These experiments demonstrate that the presence of the rhodium complex does not preclude the stoichiometric formation of duplexes, and that the association of the Λ enantiomer of the metallointercalator with the duplex is different from the interactions of the ∆ enantiomer. (B) Establishing the Intercalation Site by Photocleavage in Rh-Modified DNA Duplexes. To apply Rh-modified oligonucleotides in studies of long-range charge transfer in DNA, it is critical that we know the site at which the reactants are bound to the DNA. Upon photoactivation with UV light (λ ≈ 313 nm) in the presence of DNA, phi complexes of Rh(III) may abstract a hydrogen atom from the sugar residue at the base step of intercalation (20), which leads to direct strand scission and marks the site of intercalation. The products of the photochemical reaction with 32P-labeled DNA are separated by PAGE and visualized by phosphorimagery (25). This experiment provides single-base resolution of the site of cleavage and permits the determination of the molecular distances that separate electron donors and acceptors during charge-transfer experiments. Figure 7 shows the polyacrylamide gel used to reveal the site of intercalation in rhodium-modified DNA duplexes as a function of the length of the linker and the binding site. To clarify these results, the data are summarized schematically in Figure 8. We examined photocleavage by the ∆ and Λ enantiomers of [Rh(phi)2(bpy′)]3+ tethered by diaminononane (C9) or diaminohexane (C6) linkers to a 15 base pair (bp) duplex (lanes 4-7; Figure 7). In lanes 8-11, the Rhmodified strand contains 16 nt, but it is hybridized to the 15 nt complement used for duplexes in lanes 4-7. This hybrid duplex has a thymine that extends the effective length of the linker joining the duplex and the C9 or C6 tether. These reactions were also compared to photocleavage by the parent complex [Rh(phi)2(dmb)]3+ (dmb ) 4,4′-dimethyl bipyridine) bound noncovalently to the unmodified duplexes (lane 14). For the samples where the complex was bound noncovalently, the cleavage is distributed fairly evenly throughout the duplex, with some specificity for binding between the 5′-G(3) and 5′A(4), as designed. In fact, the 5′-ACGA-3′ sequence was chosen to accommodate the tethered complex since this is a preferred binding sight of [Rh(phi)2(dmb)]3+. Cleavage by the tethered complex is only observed near the end of
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Figure 7. Phosphorimagery results depicting the products of the direct photocleavage of Rh-modified duplexes upon irradiation at 313 nm for 8 min. A schematic representation of these results is shown in Figure 8. The experiment measures the site of intercalation as a function of configuration about the metal center, linker length, and base sequence for a series of rhodiumDNA conjugates. All samples were 8 µM in duplex in 5 mM Na3PO4 and 50 mM NaCl, pH 7.0. The contents of each lane are as follows: 1-3, 15 bp duplex, 16/15-mer hybrid duplex, 16 bp, duplex, respectively, no irradiation (dark control); 4, 5, Λ-Rh15 bp duplex, and ∆-Rh-15 bp duplex with C9 linker; 6, 7, Λ-Rh15 bp duplex, and ∆-Rh-15 bp duplex with C6 linker; 8, 9, Λ-Rh16/15 hybrid duplex, and ∆-Rh-16/15 hybrid duplex with C9 linker; 10, 11, Λ-Rh-16/15 hybrid duplex, and ∆-Rh-16/15 hybrid duplex with C6 linker; 12, 13, Maxam-Gilbert sequencing lanes for 15-mer (A + G, 12; C + T, 13); 14, ∆-[Rh(phi)2(dmb)]3+ bound noncovalently to 15 bp duplex.
the duplex to which the metallointercalator is confined by the linker. For modified duplexes bearing the C9 or C6 linkers, the Λ isomer binds primarily two bases in from the 5′terminus and the ∆ isomer binds three bases from the terminus. This observation has been made repeatedly for rhodium complexes tethered to the sequence 5′-ACGA(T). In quantitative comparisons of cleavage intensities, it is apparent that there is a diminution in the amount of cleavage with the C6 linker, suggesting that this linker might not be long enough to permit efficient access of the preferred binding site. The presence of the overhanging thymine in the hybrid duplexes (Rh-16-mer hybridized to a 15 nt complement) does not seem to affect the site of intercalation, but the cleavage intensity does appear to decrease. Synthesis of 3′,5′-bis-Hetereometalated Oligonucleotides. Scheme 3 outlines the preparation of oligonucleotides with a metallointercalator attached to the 3′hydroxyl group. Over the course of these reactions, the 5′-hydroxyl is protected as the DMT ether (19). The last steps in that procedure involve deprotection of the 5′hydroxyl group and liberation of the conjugate from the resin. We recognized that if we unmasked the 5′-hydroxyl of the 3′-modified oligonucleotide, left on the resin, we would be free to functionalize the 5′-hydroxyl as well (26, 27). We could synthesize oligonucleotides that contain an electron donor, for example, at one terminus and an electron acceptor at the other. We developed the strategy in Scheme 4, which permits the synthesis of so-called 3′,5′-bis-heterometalated oligo-
Holmlin et al.
Figure 8. Summary of photocleavage of a 15 bp duplex by [Rh(phi)2(bpy′)]3+ tethered to the 5′-ACGA terminus as indicated. The columns correspond to lane numbers in Figure 7 as follows: (left column, top to bottom) lane 4, lane 6, lane 8, lane 10; (right column, top to bottom) lane 5, lane 7, lane 9, lane 11. Arrows represent sites of cleavage and their height represents relative cleavage intensity.
Figure 9. ESI mass spectrum of a 3′,5′-bis-heterometalated oligonucleotide. The calculated mass for the fully protonated species is Mcalc ) 6643.2. According to the distributions of ions present in the mass spectrum, the mass of the ion plus the appropriate number of hydrogens to achieve the targeted charge, leads to a mass of Mobs ) 6643.5. This remarkable accuracy confirms that the material is the desired compound, and that all protecting groups have been removed.
nucleotides. The first phase of the synthesis consists of incorporating a metallointercalator onto the hydroxy aminoalkyl linker tethered to the 3′ terminus of the oligonucleotide according to the procedure developed for preparing 3′-modified DNA. Next, the DMT protecting group is removed from the 5′ hydroxyl by treatment with trichloroacetic acid in CH2Cl2. After coupling the appropriate diaminoalkyl linker to the 5′ hydroxyl, the second metallointercalator is coupled to the free amine
Synthesis of Metallointercalator−DNA Conjugates Scheme 4. Synthesis of 3′,5′-bis-Heterometallated Oligonucleotides on a Solid Supporta,b
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DNA (33), and to compare long-range charge-transfer reactions in duplexes with 5′-GG-3′ and thymine dimer sites to those with either 5′-GG-3′ or thymine dimers (36). This work established for the first time that DNA damage and repair chemistry can be initiated from a remote site. Rh-DNA conjugates have also been useful, along with ethidium-modified DNA to characterize DNA-mediated electron-transfer chemistry using photophysical techniques (37). For future applications, these conjugates, including the 3′,5′-bis-heterometalated oligonucleotides, may provide the basis for developing novel DNA-based sensor technologies based on charge-transfer reactions involving the DNA base stack. ACKNOWLEDGMENT
a
Reagents: (a) CDI, dioxane, 30 min, rt; (b) H2N(CH2)9NH2, dioxane/H2O (9:1), 25 min, rt; (c) TSTU, DIEA, CH3OH, CH3CN, CH2Cl2 (1:1:1), rt, 1 h; (d) heterogeneous coupling, DIEA, rt, 12 h; (e) NH3(aq), 60 °C, 6 h. b Abbreviations: CPG ) controlled pore glass; Os ) [Os(phen)(bpy′)(Me2-dppz)]2+.
according to the procedure established for modification of the 5′ terminus. Finally, the whole assembly is liberated from the conjugate and deprotected by treatment with NH3(aq), and isolated by HPLC. We executed this procedure to validate our method and isolated one major product. It is important to note that using racemic mixtures of each metallointercalator gives rise to eight possible isomers of the 3′,5′-bis-heterometalated oligonucleotide. The ESI MS of the product isolated in this reaction is shown in Figure 9. This methodology will be important in the application of DNA-mediated electron transfer for metallointercalator-DNA conjugates in sensing technology. SUMMARY AND CONCLUSIONS
We have established methods to prepare oligonucleotides bearing metallointercalators on either the 5′, 3′, or both 5′ and 3′ termini. The procedures are based on a solid-phase approach in that the conjugates are prepared while the oligonucleotide is immobilized on the CPG bead used in automated DNA synthesis. This approach is attractive for three reasons: (i) excess reagents that drive reactions to completion can be easily washed away from the solid support following reactions; (ii) the phosphate groups are protected so the oligonucleotide is uncharged and well solvated; and (iii) all exocyclic amines on the DNA bases are protected so the only reactive functionality present is the desired point of attachment. These protocols all provide a means to synthesize, in high, reproducible yields, a variety of conjugates that vary in the DNA sequence of the oligonucleotide as well as the length of the linker used to join the two species. We employed several methods to characterize the materials and establish the chemical composition of the conjugates as well as their ability to form duplexes with their complementary strands and the site of intercalation for the metallointercalator. The methodology described in this paper has now been employed to prepare metallointercalator-DNA conjugates for investigations of the fundamental properties of DNA as a medium for charge transfer. Rh-modified oligonucleotides have been employed to demonstrate long-range oxidative DNA damage to 5′-GG-3′ sites (8), as well as to examine the effect of intervening bulges in the base stack (35). We have used Rh-modified DNA to effect the long-range repair of thymine-dimer lesions in
We are grateful to the NIH for their financial support. We also thank the Parsons Foundation (R.E.H.) and the Damon-Runyon Walter Winchell Cancer Fund (P.J.D.) for fellowship support. Supporting Information Available: Illustration of the NHS ester of [Rh(phi)2(bpy′)]3+, MALDI-TOF mass spectrum of [Rh(phi)2(bpy′-NHS)]3+, and HPLC chromatograms. This material is available free of charge via the Internet at http://pubs.acs.org. LITERATURE CITED (1) Holmlin, R. E., Dandliker, P. J., and Barton, J. K. (1997) Angew. Chem., Int. Ed. Engl. 36, 2714-2730. (2) Arkin, M. R., Stemp, E. D. A., Holmlin, R. E., Barton, J. K., Ho¨rmann, A., Olson, E. J. C., and Barbara, P. F. (1996) Science 273, 475-480. (3) Holmlin, R. E., Stemp, D. A., and Barton, J. K. (1996) J. Am. Chem. Soc. 118, 5236-5244. (4) Murphy, C. J., Arkin, M. R., Ghatlia, N. D., Bossmann, S., Turro, N. J., and Barton, J. K. (1994) Proc. Natl. Acad. Sci. U.S.A. 91, 5315-5319. (5) Arkin, M. R., Stemp, E. D. A., Turro, C., Turro, N. J., and Barton, J. K. (1996) J. Am. Chem. Soc. 118, 2267-2274. (6) Lincoln, P., Tuite, E., and Norde´n, B. (1997) J. Am. Chem. Soc. 119, 1454-1455. (7) Olson, E. J. C., Hu, D., Hormann, A., and Barbara, P. F. (1997) J. Phys. Chem. B. 101, 299-303. (8) Hall, D. B., Holmlin, R. E., and Barton, J. K. (1996) Nature 382, 731-735. (9) Arkin, M. R., Stemp, E. D. A., Pulver, S. C., and Barton, J. K. (1997) Chem. Biol. 4, 389-400. (10) Murphy, C. J., Arkin, M. R., Jenkins, Y., Ghatlia, N. D., Bossmann, S. H., Turro, N. J., and Barton, J. K. (1993) Science 262, 1025-1029. (11) Hurley, D. J., and Tor, Y. (1998) J. Am. Chem. Soc. 120, 2194-2195. (12) Khan, S. I., and Grinstaff, M. W. (1999) J. Am. Chem. Soc. 121, 4704-4705. (13) Meggers, E., Kusch, D., and Giese, B. (1997) Helv. Chim. Acta 80, 640-652. (14) Bannwarth, W., and Schmidt, D. (1989) Tetrahedron Lett. 30, 1513-1516. (15) Manchanda, R., Dunham, S. U., and Lippard, S. J. (1996) J. Am Chem. Soc. 118, 5144-5145. (16) Schliepe, J., Berghoff, U., Lippert, B., and Cech, D. (1996) Angew. Chem., Int. Ed. Engl. 35, 646-648. (17) Mucic, R. C., Herrlien, M. K., Mirkin, C. A., and Letsinger, R. L. (1996) Chem. Commun. 555-557. (18) Magda, D., Crofts, S., Lin, A., Miles, D., Wright, M., and Sessler, J. L. (1997) J. Am. Chem. Soc. 119, 2293-2294. (19) Caruthers, M. H. (1985) Science 230, 281-285. (20) Sitlani, A., Long, E. C., Pyle, A. M., and Barton, J. K. (1992) J. Am. Chem. Soc. 114, 2303-2311.
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