Naphthalenedicarboxamides as Fluorescent Probes of Inter- and

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J. Phys. Chem. B 1999, 103, 2570-2578

Naphthalenedicarboxamides as Fluorescent Probes of Inter- and Intramolecular Electron Transfer in Single Strand, Hairpin, and Duplex DNA Frederick D. Lewis,* Yifan Zhang, Xiaoyang Liu, Ning Xu, and Robert L. Letsinger Department of Chemistry, Northwestern UniVersity, EVanston, Illinois 60208 ReceiVed: NoVember 30, 1998; In Final Form: January 28, 1999

The 2,6-naphthalenedicarboxamide chromophore has been investigated as a fluorescent probe for DNA hairpin and duplex formation and DNA electron transfer. The high fluorescence quantum yield and long singlet lifetime of this chromophore make it an attractive candidate for these studies. The kinetics of intermolecular quenching of a naphthalenedicarboxamide by nucleosides is dependent upon the nucleoside oxidation potential and solvent. Bis(oligonucleotide) conjugates containing naphthalene linkers have been prepared by means of conventional phosphoramidite chemistry. The base-sequence dependence of the naphthalene fluorescence intensity and decay times in both single-strand and hairpin conjugates indicates that singlet naphthalene is quenched by neighboring dA more efficiently than by dT, in accord with an electron-transfer quenching mechanism. These data are analyzed by means of a three-state model which includes a nonemissive “dark” state. Duplexes formed between complementary naphthalene-linked oligonucleotides display naphthalene excimer emission. The base-sequence dependence of the excimer emission quantum yields indicates that the excimer is not quenched by neighboring dA but that distance-dependent electron-transfer quenching by dG may occur. Quenching serves to protect the naphthalene chromophore from photobleaching in both single strand and duplex structures.

Introduction Oligonucleotides containing non-nucleotide linkers can form duplex and hairpin structures in which the linker enhances the stability of the base-paired structure.1-13 We have made use of stilbenedicarboxamide linkers as fluorescence probes for the formation and properties of duplex and hairpin DNA.8-13 Hybridization of complementary pairs of mixed-base sequence bis(oligonucleotide) conjugates which contain appropriately positioned stilbenedicarboxamide linkers results in formation of a duplex in which the stilbenes associate to form a groundstate dimer.12,13 Electronic excitation of this dimer results in both excimer fluorescence and stilbene photodimerization. Conjugates in which a stilbenedicarboxamide serves as a linker between short complementary base pair sequences can form stable hairpin structures in which the stilbene is parallel to the adjacent base pair.9-11 Stilbene fluorescence is selectively quenched by guanine via an electron transfer mechanism in which stilbene acts as an electron acceptor and guanine as an electron donor. Investigation of the kinetics of electron transfer for a family of stilbene-linked hairpins containing a single dGdC base pair separated from the stilbene by a variable number of dT-dA base pairs provided the first direct measurement of the distance dependence of electron transfer in DNA.10 The stilbenedicarboxamides provided a useful entry point for our investigations. However, their use as fluorescence probes is compromised by their lack of photostability and relatively short singlet lifetime.14 Photoisomerization of the fluorescent trans-stilbene to the nonfluorescent cis isomer in single strand and hairpin DNA9 and stilbene photodimerization in duplex DNA13 results in bleaching of the monomer and excimer fluorescence, respectively. The short stilbene singlet lifetime * E-mail: [email protected]. Telephone: 847-491-3441. Fax: 847467-2184.

precludes investigation of either intermolecular electron transfer or slow intramolecular ( 8 > 7, parallel the relative fluorescence intensities. Thus, quenching of singlet naphthalene by neighboring nucleobases serves to protect it from photodegradation. Hairpin Conjugates. The bis(oligonucleotide) conjugates 12 and 13 can adopt hairpin structures in which the naphthalene dicarboxamide serves as a linker between the complementary poly(dT) and poly(dA) segments. We have reported the formation of related synthetic DNA hairpins with stilbenedicarboxamide and terephthalamide bridges.8-10 The TM for 12 is intermediate between the values reported for stilbenedicarboxamide-bridged hairpins with four or six dT-dA base pairs for which TM ) 49 and 59 °C, respectively.9 Introduction of a dTdT mismatch at the top of the hairpin results in only a small decease in TM for 13 vs 12 (Table 1). The TM values for the arenedicarboxamide linked hairpins are independent of concentration and are higher than those expected for the duplexes

formed between these self-complementary sequences. For example, the TM calculated for the dimer of 12 in 0.1 M NaCl using the method of SantaLucia34 is 25 °C, substantially below the observed value (Table 1). Molecular modeling of 12 (Figure 7a) indicates that it can adopt a B-form double helical structure with the naphthalenedicarboxamide roughly parallel to the adjacent dT-dA base pair. However, the minimum naphthalenedA plane-to-plane distance is 5.0 Å, too long for strong electronic interaction. Additional experimental evidence is necessary to define the geometry in the region of the naphthalene linker. The hairpins 12 and 13, like the single strand conjugates 5-11, have lower fluorescence quantum yields than the diol 2 and display dual exponential fluorescence decay. In the case of 12 both the fluorescence quantum yield and the lifetime of the major 0.31-ns decay component are lower than those of 2 by a factor of 40. Thus, there is no need to postulate a role for a dark state. Assuming that quenching is the result of electron transfer from dA or dT to the singlet naphthalene, the reduction in lifetime can be used to estimate the rate constant for quenching by neighboring dA or dT bases (kq ) τ12-1 - τ2-1), kq ) 3 × 109 s-1. On the basis of the lower oxidation potential of dA vs dT and the faster rate constant for intermolecular quenching by dA vs dT (Table 3), intramolecular quenching by dA is assumed to be more rapid than quenching by dT. In accord with this assumption, placement of a dT-dT mismatch between the naphthalene and first dA-dT base pair in 13 results in an increase in both φf and the two decay times (Table 2). However, the value of φf for 13 is significantly smaller than that for 2 or 11, indicating that quenching by neighboring dA remains relatively efficient. Broadening in the long-wavelength absorption band of 13 vs 12 or 2 provides evidence for groundstate interaction between naphthalene and dA which could account for substantial dark-state formation in 13 but not in 12. The calculated rate constant of 3 × 109 s-1 for quenching of naphthalenedicarboxamide by dA is much slower than the measured rate constant of 1012 s-1 for electron transfer quenching of stilbenedicarboxamide by neighboring dG in a stilbenebridged hairpin.10 Relatively slow quenching seems inconsistent with the value of ∆Get ) - 0.33 eV calculated from Weller’s equation (eq 3) and the value of Eox for dA measured in aqueous solution (Table 3). It is more consistent with a value of ∆Get ) + 0.19 calculated using the value of Eox for nonaqueous solution. Our previous observation that the singlet stilbenedi-

Fluorescent Probes of Electron Transfer in DNA caboxamide linker is not efficiently quenched by dA-dT base pairs in stilbene-bridged hairpins 10 is also more consistent with the value of ∆Get obtained using nonaqueous vs aqueous oxidation potentials. Since the π-stacked nucelobases in duplex DNA are in a hydrophobic environment, it may be more appropriate to use nonaqueous oxidation potentials to calculate the free energy of electron transfer, particularly in case where both the forward and return electron-transfer process are rapid and thus not coupled with proton transfer. However, the use of aqueous oxidation potentials may be more appropriate for single strand or disordered regions of duplex DNA. Clearly, values of ∆Get for electron-transfer processes in duplex DNA obtained using either aqueous or nonaqueous redox potentials measured for single nucleosides should be viewed as being highly approximate. Duplex Conjugates. Pairing of the complementary naphthalene-containing conjugates 5-10 results in hypochromicity of the 340-nm absorption band assigned to the naphthalenedicarboxamide as well as the 260-nm absorption band assigned predominantly to the nucleobases (Figure 2). The temperature dependence of the normalized absorbance (Figure 3) provides values of TM for 7:8 reported in Table 1. The higher conjugate concentrations employed for TM measurements at 340 vs 260 nm can account for the higher TM values obtained at 340 nm. Also reported in Table 1 is the TM for a model duplex having the same base sequence as 5:6 but lacking the naphthalene linkers.13 The TM of 5:6 is slightly lower than that of the model, as previously observed for bis(oligonucleotide) conjugates having stilbenedicarboxamides9,13 or hydrophobic, non-hydrogenbonded isosteres of pyrimidines or purines inserted in the middle of a duplex.35 Evidently hydrophobic association of these unnatural conjugates does not fully compensate for disruption of duplex π-stacking. Also reported in Table 1 are values of TM for the model and three duplexes calculated using the method of SantaLucia,34 assuming that no correction is necessary for the presence of the linker. The calculated values for both model duplex and the duplex conjugates are several degrees higher than the observed values but follow the same order, increasing with the number of dG-dC base pairs. Duplex formation results in a decrease in monomer fluorescence intensity and the appearance of structureless emission at longer wavelength attributed to an excimer (Figures 5 and 6). Excimer fluorescence is not observed from 9:10 at 30 °C, which is above its Tm, but is readily detected at lower temperatures (Figure 6). The excimer fluorescence quantum yield for 7:8 also increases with decreasing temperature (Table 3), however the excimer fluorescence of 5:6 remains weak even at low temperatures. The excimer fluorescence decay of duplexes 7:8 and 9:10 is dual exponential (Table 3). Both the short and long decay times increase with decreasing temperature, however the change is more pronounced for the short decay. Formation of the naphthalene excimer in duplex DNA might occur either via excitation of a ground state naphthalene dimer by quenching of singlet naphthalene by a neighboring naphthalene. Several lines of evidence favor the excitation of a ground-state dimer. The observation of 340-nm hypochromism indicates that the naphthalenes are π-stacked in the ground state. The absence of a rising component in the excimer fluorescence decay profile indicates that the excimer is completely formed within the ca. 0.2-ns time resolution of our fluorescence lifetime apparatus. Molecular modeling confirms that the duplexes can adopt geometries in which the two naphthalenes are associated in the ground states. As shown in Figure 7, two energy minima were located for a truncated duplex with two dT-dA base pairs

J. Phys. Chem. B, Vol. 103, No. 13, 1999 2577 on either side of the naphthalenes, which can adopt either a parallel sandwich structure (Figure 7b) or a partially overlapping crossed structure (Figure 7c). The two dimer geometries may be responsible for the observation of dual exponential excimer fluorescence decay (Table 3), as observed for intramolecular excimers which have two ground-state conformations of comparable energy.19,36 The observation of naphthalene excimer fluorescence from the duplexes further illustrates the utility of DNA base pairing to align pairs of conjugated molecules without the necessity of covalent attachment.12,13 Singlet naphthalene in homogeneous solution fails to undergo self-quenching or excimer formation. However, intramolecular excimer fluorescence is observed for 1,3-di-2-naphthylpropane 1417,18 and naphthalenophanes such as 15.19 The red-shifts for excimer vs monomer fluorescence in 14 (420 vs 340 nm) and 15 (404 vs 335 nm both in alkane solvent) are somewhat larger than for the duplex 7:8 (410 vs 360 nm in aqueous solution). Since excimer fluorescence maxima display small but finite solvatochromic shifts,37 the redshift for duplex 7:8 might be even smaller in comparison to those of 13 and 14 in the same solvent. The smaller red-shift may reflect an excimer geometry which does not have optimal orbital overlap, as observed for partially overlapping diarylalkanes and cyclophanes.19,36

Of the three duplexes investigated 5:6 forms the most stable duplex (Table 1) but displays the weakest excimer fluorescence (Figure 5a). The observation of hypochromism for the 340-nm absorption band of 5:6 (Table 1) and of very weak naphthalene monomer fluorescence (Figure 5a) indicates that a ground-state dimer is formed, but that it is very weakly fluorescent. The most obvious structural difference between the three duplexes is shorter distance between the closest dG-dC base pair and the naphthalene dimer in 5:6. Excimer fluorescence quenching by the closest dG could account for weaker excimer fluorescence from 5:6 than for 7:8 and 9:10, which have two additional dTdA base pairs between the naphthalenes and nearest dG. Quenching of the naphthalene excimer should be less favorable than quenching of the naphthalene monomer due to the lower energy of the excimer.38 Furthermore, unlike arene cation radicals which form stable dimer cation radicals, dimer anion radicals are dissociative.39 Comparison of the monomer fluorescence quantum yield and lifetime for the hairpin 12 with those of the duplexes 7:8 or 9:10 indicates that quenching of the naphthalene excimer by dA is much less efficient than quenching of the monomer by dA. However, dG is a better donor than dA and thus might be able to quench naphthalene excimer even when separated from the excimer by one dT-dA base pair in the duplex 5:6. Assuming a normal distance dependence for electron transfer in DNA,10 separation of the excimer from dG by two additional base pairs in duplexes 7:8 or 9:10 should reduce the rate constant for electron-transfer quenching by at least 10-fold. It is interesting to note that quenching of stilbene excimer by dG was not observed in our investigation of duplex formation from the stilbene-containing analogues of conjugates 5 and 6.13 The quenching of naphthalene excimer, but not that of stilbene excimer, may reflect the higher singlet energy of the former excimer. Comparison of the excimer fluorescence maxima of the naphthalene and stilbene-containing duplexes (λmax ) 410

2578 J. Phys. Chem. B, Vol. 103, No. 13, 1999 and 455 nm, respectively) indicates that electron-transfer quenching of the naphthalene excimer should be more exergonic by ca. 0.3 eV. Concluding Remarks. Oligonucleotide conjugates containing 2,6-naphthalenedicarboxamide linkers can be readily synthesized from the commercially available dicarboxylic acid. Naphthalenelinked hairpin and duplex structures are comparable in thermodynamic stability to the structures of the previously investigated stilbene-linked analogues.9-13 The long-fluorescence lifetime and high fluorescence quantum yield for the naphthalenedicarboxamide chromophore make it well-suited for use of as a fluorescence probe. However, a weak long-wavelength absorption band necessitates use at concentrations higher than those for probes with fully allowed absorption bands. Intermolecular singlet quenching of the diol 2 by individual nucleosides is strongly solvent-dependent. Rate constants for quenching by pyrimidine bases are substantially smaller than for quenching by purine bases in DMF solution but are more nearly comparable in aqueous solution, in accord with differences in nucleoside oxidation potentials in aqueous vs nonaqueous solution. Thus, the energetics of electron-transfer quenching by nucleotides in single strand or duplex DNA may depend on the location of the nucleobase in either a hydrophobic or hydrophilic environment. The fluorescence quantum yield and lifetime of naphthalenelinked nucleotides is dependent upon the identity of the neighboring nucelobases. From the base-sequence dependence of the fluorescence quantum yields for single strand conjugates 5-11 it is evident that dA is a more effective quencher than dT, in accord with the lower oxidation potential of dA. The observation of more extensive quenching of fluorescence intensity vs decay times can be accounted for by use of a threestate model in which ground-state interaction of naphthalene and dA results in the formation of a nonemissive dark state. Comparison of the decay times of the diol 2 and the hairpin 12 provides a rate constant for quenching of naphthalene quenching by an adjacent dA of 3 × 109 s-1. Dark-state formation is less important in hairpin vs single strand structures, presumably due to the more rigid hairpin structure. Duplexes formed between complementary naphthalene-linked conjugates display naphthalene excimer emission. The base-sequence dependence of the excimer emission quantum yields indicates that distancedependent electron-transfer quenching by dG may occur. Acknowledgment. This research is supported by the Division of Chemical Sciences, Office of Basic Energy Sciences, U.S. Department of Energy under contract DE-FG02-96ER14604. We thank T. L. Netzel for helpful discussions. References and Notes (1) Goodchild, J. Bioconjugate Chem. 1990, 1, 165-191. (2) Durand, M.; Chevrie, K.; Chassignol, M.; Thuong, N. T.; Maurizot, J. C. Nucleic Acids Res. 1990, 18, 6353-6359. (3) Durand, M.; Peloille, S.; Thuong, N. T.; Maurizot, J. C. Biochemistry 1992, 31, 9197-9204.

Lewis et al. (4) Cload, S. T.; Schepartz, A. J. Am. Chem. Soc. 1991, 113, 63246326. (5) Kessler, D. J.; Pettitt, B. M.; Cheng, Y. K.; Smith, S. R.; Jayaraman, K.; Huynh, M. V.; Hogan, M. E. Nucleic Acids Res. 1993, 21, 4810-4815. (6) Rumney, S.; Kool, E. T. J. Am. Chem. Soc. 1995, 117, 56355646. (7) Gao, H.; Yang, M.; Cook, A. F. Nucleic Acids Res. 1995, 23, 285292. (8) Salunkhe, M.; Wu, T.; Letsinger, R. L. J. Am. Chem. Soc. 1992, 114, 8768-8772. (9) Letsinger, R. L.; Wu, T. J. Am. Chem. Soc. 1995, 117, 73237328. (10) Lewis, F. D.; Wu, T.; Zhang, Y.; Letsinger, R. L.; Greenfield, S. R.; Wasielewski, M. R. Science 1997, 277, 673-676. (11) Lewis, F. D.; Letsinger, R. L. J. Biol. Inorg. Chem. 1998, 3, 215221. (12) Letsinger, R. L.; Wu, T. J. Am. Chem. Soc. 1994, 116, 811-812. (13) Lewis, F. D.; Wu, T.; Burch, E. L.; Bassani, D. M.; Yang, J. S.; Schneider, S.; Ja¨ger, W.; Letsinger, R. L. J. Am. Chem. Soc. 1995, 117, 8785-8792. (14) Waldeck, D. H. Chem. ReV. 1991, 91, 415-436. (15) Berlman, I. B. Handbook of Fluorescence Spectra of Aromatic Molecules, 2nd ed.; Academic Press: Ltd.: London, 1971. (16) Lewis, F. D. AdV. Photochem. 1986, 13, 165-235. (17) Chandross, E. A.; Dempster, C. J. J. Am. Chem. Soc. 1970, 92, 3586-3593. (18) Lim, E. C. Acc. Chem. Res. 1987, 20, 8-17. (19) Yanagidate, M.; Takayama, K.; Takeuchi, M.; Nishimura, J.; Shizuka, H. J. Phys. Chem. 1993, 97, 8881-8888. (20) Mendicuti, F.; Saiz, E.; Mattice, W. L. Polymer 1992, 33, 49084912. (21) Yang, C. P.; Chen, W. T. J. Polym. Sci., Part A: Polym. Chem. 1993, 31, 1571-1578. (22) Ohya, Y.; Noro, H.; Komatsu, M.; Ouchi, T. In One-Dimensional Polymeric Supramolecular Assemblies Using Oligonucleotides; Ohya, Y., Noro, H., Komatsu, M., Ouchi, T., Eds.; American Chemical Society: Orlando, FL, 1996. (23) James, D. R.; Siemiarczuk, A.; Ware, W. R. ReV. Sci. Instrum. 1992, 64, 1710. (24) Burdett, K. A. Synthesis 1991, 441-442. (25) Lui, Y. H.; McGlynn, S. P. J. Mol. Spectrosc. 1974, 49, 214-225. (26) Shafirovich, V. Y.; Courtney, S. H.; Ya, N.; Geacintov, N. E. J. Am. Chem. Soc. 1995, 117, 4920-4929. (27) Seidel, C. A. M.; Schulz, A.; Sauer, M. H. M. J. Phys. Chem. 1996, 100, 5541-5553. (28) Lewis, F. D.; Bassani, D. M. J. Photochem. Photobiol. A: Chem. 1992, 66, 43-52. (29) Cadet, J.; Vigny, P. In Photochemistry of Nucleic Acids; Cadet, J., Vigny, P., Eds.; Wiley-Interscience: New York, 1990; Vol. 1, pp 1-272. (30) Weller, A. Zeit. Phys. Chem. Neu. Folg. 1982, 133, 93-98. (31) Steenken, S.; Jovanovic, S. V. J. Am. Chem. Soc. 1997, 119, 617618. (32) Seidel, C.; Rittinger, K.; Cortes, J.; Goody R. S.; Koellner, M.; Wolfrum, J.; Gruelich, K. O. Proc. SPIE-Int. Soc. Opt. Eng. 1991, 1432, 105-116. (33) Va´mosi, G.; Gohlke, C.; Clegg, R. M. Biophys. J. 1996, 71, 972994. (34) SantaLucia, J., Jr. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 14601465. (35) Schweitzer, B. A.; Kool, E. T. J. Am. Chem. Soc. 1995, 117, 18631872. (36) Hayashi, T.; Mataga, N.; Sakata, Y.; Misumi, S.; Morita, M.; Tanaka, J. J. Am. Chem. Soc. 1976, 98, 5910-5913. (37) Lewis, F. D.; Burch, E. L. J. Photochem. Photobiol. 1995, 96, 1923. (38) Caldwell, R. A.; Creed, D.; DeMarco, D. C.; Melton, L. A.; Ohta, H.; Wine, P. H. J. Am. Chem. Soc. 1980, 102, 2369-2377. (39) Majima, T.; Tojo, S.; Takamuku, S. J. Phys. Chem. 1997, 101, 1048-1055.