Structure and Photoinduced Electron Transfer in DNA Hairpin

Oct 20, 2009 - Hairpin, and (d) Intercalated Tethered Conjugate. J. Phys. Chem. B 2009 ... Reaction of pyrene-1-carboxylic acid (1.0 g, 4.1 mmol) with...
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Structure and Photoinduced Electron Transfer in DNA Hairpin Conjugates Possessing a Tethered 5′-Pyrenecarboxamide Karsten Siegmund,† Pierre Daublain,† Qiang Wang,‡ Anton Trifonov,‡ Torsten Fiebig,*,‡ and Frederick D. Lewis*,† Departments of Chemistry, Northwestern UniVersity, EVanston, Illinois 60208-3113, and Boston College, Chestnut Hill, Massachusetts 02467 ReceiVed: July 30, 2009; ReVised Manuscript ReceiVed: September 2, 2009

The structure, spectroscopy, and photophysical behavior of a series of hairpin-forming conjugates possessing a 5′-tethered N-alkylpyrenecarboxamide chromophore have been investigated. Comparison of the NMR spectra of the conjugates and analogs lacking the tethered pyrene indicates that the pyrene does not behave as an end-capping group but rather is intercalated between the two terminal hairpin base pairs. An intercalated structure is also consistent with the thermodynamic parameters for hairpin formation and the steady state and transient spectral properties of the conjugates. Quenching of the pyrene fluorescence and transient absorption spectra is observed only when guanine is located in one of the two terminal base pairs and is attributed to hole injection from singlet pyrene to guanine. The fast component of the transient decay is more rapid when guanine is located in the second vs first base pair, consistent with an intercalated but not an end-capped geometry. Spectral broadening of ultraviolet, fluorescence, and transient absorption spectra is attributed to multiple ground state conformations. Introduction Oligonucleotide conjugates possessing flexibly tethered chromophores have been extensively employed in studies of DNA structure and properties, including conformational dynamics and charge transfer. Bis(oligonucleotide) conjugates possessing doubly tethered chromophores have been utilized for the preparation of hairpin,1–4 duplex,5 and triplex structures6 as well as structures possessing multiple chromophores.7 Hairpin structures have been widely used in investigations of photoinduced DNA charge transfer by our group and others.3,8 The position of the chromophore is clearly defined in linked hairpins possessing short flexible linkers (Chart 1a).9,10 The requirement of monoactivated, monoprotected difunctional derivatives has limited the selection of chromophores suitable for bis(oligonucleotide) conjugate synthesis.11 The use of singly tethered aromatic chromophores offers greater ease of synthesis and thus provides access to a wider range of conjugate structures. The singly tethered chromophore can be introduced on the exterior, interior (Chart 1b), or the 5′or 3′-terminal end of a duplex (Chart 1c,d). Covalent attachment to a nucleobase has been used to introduce chromophores into the duplex major or minor groove.12,13 These conjugates display complex excited state kinetics and have not been widely employed in studies of DNA charge transfer. Interior placement of single or multiple aromatic chromophores has been accomplished using synthetic nucleotides14,15 and peptide nucleic acid analogs.16 Interior placement complicates studies of photoinduced charge transfer in duplex structures by providing two competing directional pathways.13,17 Thus 5′-terminal conjugates have been employed in a majority of recent investigations of DNA charge transfer.18,19 * E-mail: [email protected];[email protected]. † Northwestern University. ‡ Boston College.

CHART 1: Structures of Several DNA Conjugates: (a) Hairpin, (b) Internal Base Pair Surrogate, (c) Capped Hairpin, and (d) Intercalated Tethered Conjugate

Duplex structures possessing flexibly 5′-tethered coordination complexes20 and acridines21 are known to intercalate between base pairs near the location of tether attachment; however, the precise location of intercalation is not uniquely determined. Smaller neutral organic chromophores are normally assumed to form capped structures (Chart 1c) in which hydrophobic association between the terminal base pair and the organic chromophore serves to stabilize the duplex structure.19,22–24 However, both intercalating and capping geometries for the tethered chromophore can account for an increase in duplex stability and fluorescence quenching by neighboring bases.25 A capped structure has been reported for a duplex possessing a 5′-tethered stilbene derivative similar to those used in our studies of photoinduced electron transfer in stilbene-capped hairpins.26 However, end-capping structures have not been established for other 5′-tethered chromophores and recent structural studies suggest that they may in fact adopt alternative conformations (Chart 1d).27 Since chromophore location can influence steady state and transient spectral data, interpretation of such data without knowledge of structure is a questionable practice.

10.1021/jp907323d CCC: $40.75  2009 American Chemical Society Published on Web 10/20/2009

Electron Transfer in DNA Hairpin Conjugates CHART 2: Structures of (a) Pyrenecarboxamide Derivatives, (b) Pya-Conjugates Shown as Capped Hairpins, and (b) Reference Hairpin Sequences Lacking Pya

We report here the results of a collaborative investigation of the excited state behavior and ground state structure of a family of hairpin-forming oligonucleotide conjugates possessing a 5′tethered pyrene-1-carboxamide (Pya, Chart 2a) and a G-C base pair separated from Pya by zero to four A-T base pairs (Chart 2b). Fluorescence quenching is observed only when G is the first or second base nearest Pya. Furthermore, the major component of 1Pya* transient decay is more rapid when G is located at the second vs first position. These observations prompted an NMR study of several hairpin-forming Pya conjugates and the corresponding unmodified hairpins (Chart 2c). This study establishes that these conjugates do not adopt stable capped-hairpin structures. A more extensive investigation of the structure of conjugate Pya-2G indicates that Pya is intercalated between the first and second base pairs, thus accounting for its seemingly anomalous photophysics. Experimental Section Materials. N-(3-Hydroxypropyl)pyrene-1-carboxamide (PyaH). Reaction of pyrene-1-carboxylic acid (1.0 g, 4.1 mmol) with excess thionyl chloride afforded the acid chloride that was reacted with (3-hydroxypropyl)amine (6.1 mmol) and triethylamine (61 mmol) in tetrahydrofuran followed by silica get chromatography (methanol) to provide Pya-H as a pale yellow solid (1.0 g, 3.3 mmol). Mp: 147-149 °C. 1H NMR (500 MHz, DMSO-d6): δ 8.4 (1H, d), 8.2 (2H, m), 8.0 (6H, m), 6.7 (1H, bs), 3.8 (4H, m), 3.3 (1H, b), 1.9 (2H, m). 13C NMR (DMSOd6): δ 172.19, 132.04, 130.86, 130.52, 128.13, 127.28, 127.0, 126.51, 126.40, 125.20, 124.89, 124.38, 123.92, 123.84, 123.51, 122.42, 60.41, 32.79, 28.73 ppm. ESI-MS (m/z) calculated 303.13, found 305.00. N-(N-Phenyl-N-methyl-(3-aminopropyl))pyrene-1-carboxamide (Pya-An). Reaction of the acid chloride obtained from pyrene-1-carboxylic acid (0.5 g, 2.0 mmol) with N-(3-amino-

J. Phys. Chem. B, Vol. 113, No. 50, 2009 16277 propyl)-N-methylbenzenamine (3.0 mmol) and triethylamine (30 mmol) in tetrahydrofuran followed by silica gel chromatography (75/1 dichloromethane/methanol) and washing with ether afforded the dyad Pya-An as a white solid (0.4 g, 1.1 mmol). Mp: 167-169 °C. 1H NMR (500 MHz, CDCl3): δ 8.5 (1H, d), 8.2 (2H, t), 8.0 (5H, m), 7.9 (1H, d), 7.2 (2H, t), 6.7 (3H, m), 6.2 (1H, b), 3.7 (2H, dd), 3.5 (2H, t), 3.0 (3H, s), 2 (2H, t). 13C NMR (CDCl3): δ 170.03, 168.71, 133.99, 132.46, 131.77, 131.10, 130.94, 130.67, 128.61, 128.56, 127.09, 126.24, 125.70, 125.66, 124.70, 124.50, 124.46, 124.36, 123.23, 36.96, 35.20, 28.53 ppm. ESI-MS (m/z) calculated 392.19, found 391.10. DNA Conjugates. Pya-H was converted to its phosphoramidite by reaction with 2-cyanoethyl diisopropylchlorophosphoramidite.1 The Pya conjugates and unmodified oligonucleotides shown in Chart 2 were synthesized by means of conventional phosphoramidite chemistry using a Millipore Expedite DNA synthesizer as previously described.15 Following synthesis, the conjugates were deprotected using 30% ammonium hydroxide and purified by RP-HPLC. The RP-HPLC purification was carried out on a Dionex chromatograph with a Hewlett-Packard Hypersil ODS-5 column (4.6 × 250 mm) and a 1% gradient of acetonitrile in 0.03 M triethylammonium acetate buffer (pH 7.0) with a flow rate of 1.0 mL/min. Molecular weights were determined following desalting by means of MALDI TOF mass spectrometry (Table S1, Supporting Information). General Procedures. UV absorption, fluorescence, and circular dichroism spectra were obtained as previously described on samples contained in 1 cm path length cuvettes.28 Quantum yields for fluorescence were determined using quinine sulfate in sulfuric acid as a reference standard.29 CD spectra are the average of two scans with a data interval of 1.0 nm and a time interval of 2 s per point. The base lines are corrected by subtraction of the spectrum of the buffer (10 mM phosphate, 0.1 M NaCl). Electrochemical measurements were performed using a CH Instruments Model 660A electrochemical workstation. The solvent was dimethyl sulfoxide containing 0.1 M tetran-butylammonium perchlorate electrolyte. A 1.0 mm diameter platinum disk electrode, platinum wire counter electrode, and Ag/AgxO reference electrode were employed. Ferrocene/ferrocinium (Fc/Fc+, 0.52 vs SCE) was used as an internal reference.30 Samples for kinetic spectroscopic measurements were prepared in 10 mM sodium phosphate, pH 7.2 buffer with 0.1 M NaCl (standard buffer). Samples were stirred during these measurements and absorption spectra checked for the occurrence of decomposition. Hairpin concentrations were adjusted to provide an absorbance of 0.2 at 354 nm in a 1 cm path length cuvette for fluorescence measurements and 0.27 in a 1 mm path length cell for pump-probe measurements. Fluorescence decay measurements, nanosecond transient absorption measurements, and femtosecond broadband pump-probe spectra were obtained as previously described.28 NMR Spectra and Structure Generation. NMR spectra were recorded using either a Varian Inova 500 spectrometer or a Varian Inova 600 spectrometer equipped with a Cold Probe. Samples of Pya-AT, Pya-1G, Pya-2G, AT, 1G, and 2G were HPLC purified as described above, but using ammonium acetate buffer with an acetonitrile gradient from 3% to 18% over 30 min. The samples were then desalted using a Sep-Pak Vac 1 mL cartridge (100 mg, Waters Corp., Milford, MA) with 0.1 M ammonium-carbonate buffer (6-10 mL), doubly distilled water (6-10 mL), and 50% acetonitrile/water until all material was eluted (typically 2-3 mL). The samples were dissolved in

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210 µL of 10 mM phosphate buffer in D2O (“100.0%”, SigmaAldrich) or H2O/D2O (90:10) containing 100 mM NaCl at pH 7 (buffer was adjusted to pH 7, lyophilized, and dissolved in an equal amount of D2O) and then transferred to an NMR microtube (Shigemi Co., Tokyo, Japan). For each sample a series of 1D spectra was recorded at temperatures between 10 and 70 °C. Two-dimensional spectra were recorded for Pya-2G and its control sequence 2G. The 2D NOESY spectra of Pya-2G were acquired using 16 scans at temperatures of 25, 30, and 40 °C and mixing times of 250 ms (40, 30, and 25 °C in D2O and 25 °C in 90% H2O), 125 ms (40 °C/D2O, 25 °C in 90% H2O). Spectra of the unmodified hairpin 2G were taken in D2O with a mixing time of 250 ms (17 °C) and with 125 and 250 ms mixing times (25 °C). The TOCSY (60 ms) and COSY spectra were acquired for each of the conditions NOESY spectra were taken using 8 scans. Some additional TOCSY spectra were acquired in an attempt to further optimize the temperature for line width and peak overlap in the pyrene region. Spectra in D2O were acquired with presaturation of the water protons. Spectra in 90% H2O were acquired using the WATERGATE pulse sequence for water suppression.31 Distance constraints were derived by integration of NOESY cross-peaks at a mixing time of 250 ms. The results at different temperatures were compared and yielded similar distances. Therefore the average of all values was chosen. The distanceintensity calculation was calibrated using fixed distances within the oligonucleotide (cytidine H5/H6, thymidine H7*/H6 and H2′/H2′′, H5′/H5′′) and calculated using a formula that takes spin diffusion into account.32 The constraints including exchangeable protons were entered using very loose boundaries of (1.3 Å. Additional hydrogen bonding and weak base planarity constraints were applied to all base pairs whose exchangeable protons were visible in the spectra acquired in 90% water. Calculations were run both with and without base pairing constraints for the remaining terminal AT base-pair. The protocol used for the molecular dynamics calculations had the default values provided with the program CNS.33 Topology and parameter files for DNA were used as provided with CNS; the initial parameter and topology files for the pyrene were created with the help of the program XPLO-2D34 and then edited. The link between the pyrene residue and the next oligonucleotide was created by employing a modified NUC patch, which allows for different types of carbon atom next to the phosphate. Initial structures were obtained from relatively few constraints, but include constraints from and between exchangeable protons. Refining these initial structures, constraints were visualized using a self-made script,35 incorrect constraints were removed and new constraints added using this visualization. Due to a lack of a unique assignment of the proton resonances in the pyrene moiety and the low intensity of pyrene-nucleotide cross-peaks caused by the line-broadening, addition of constraints and refinement of the structure was only pursued up to a total of ca.180 NOE constraints. These were then used to provide models for intercalation or end stacking. Results and Discussion Electronic Spectra and Thermodynamic Stability of Pyrene-1-carboxamide Conjugates. Methods employed for the synthesis and characterization of the hydroxypropyl-carboxamide Pya-H, the dyad Pya-An, the conjugates Pya-AT and PyanG, and the unmodified conjugates AT, 1G, and 2G (Chart 2) are described in the Experimental Section. The absorption and fluorescence spectra of Pya-H in methanol are shown in Figure 1a. The presence of the amide substituent has little effect on

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Figure 1. UV (solid line) and fluorescence spectra (dashed line, excitation wavelength ) 350 nm) of (a) Pya-H in methanol and (b) Pya-AT in standard buffer (10 mM phosphate, pH 7.2, 100 mM NaCl).

the appearance of these spectra, which display vibronic structure similar to that of unsubstituted pyrene.29 The weak absorption maximum at 380 nm is assigned to the pyrene band 1 and the stronger maximum at 348 nm to pyrene band 2. The cyclic voltammogram for Pya-H in acetonitrile displays irreversible reduction and oxidation waves with one-electron reduction and oxidation potentials of -1.99 and +1.53 V, respectively (referenced to SCE using ferrocene as an internal redox standard). The absorption and fluorescence spectra of the dyad Pya-An in methanol are similar to those of Pya-H. Both the Pya-modified and unmodified conjugates possess a GC base pair adjacent to a ACC loop, a base sequence known to promote the formation of stable mini-hairpins.23,36 Thermodynamic parameters and melting temperatures obtained by analysis of the thermal dissociation profiles for several of the conjugates are reported in Table 1. The increase in Tm values for the series AT < 1G < 2G parallels the results obtained using a web-based calculator for these hairpins.37 Introduction of the 5′-Pya group results in a similar increase in Tm for all three sequences, the largest increase being observed for Pya-2G. The slightly greater stability of hairpins with G in the second vs first position is entropic rather than enthalpic in origin (Table 1). The value of ∆Tm for Pya-1G vs 1G (11.5 °C) is somewhat larger than those reported for duplexes possessing 5′-tethered pyrenes.25,38 However, the value of ∆Tm for Pya-AT vs AT is approximately half as large as that reported for introduction of a 5′-stilbenecarboxamide group to a similar mini-hairpin.15 As previously noted by Mann et al., an increase in Tm for conjugated versus non-conjugated duplexes is consistent with either capping or intercalated geometries, but not with a groove-bound geometry.25 The absorption and fluorescence spectra of the Pya conjugates in aqueous buffer (Figures 1b and S1 (Supporting Information))

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TABLE 1: Thermodynamics and Melting Points of Pya-DNA Conjugates and Their Unmodified Controls sequencea

∆H/(kcal/mol)b

∆S/(cal · K-1 · mol-1)b

∆G/(kcal · /mol, 37 °C)

Tm/°C ( 1 °Cb

∆Tm/°Cc

AT 1G 2G Pya-AT Pya-1G Pya-2G

-34 ( 3 -45 ( 1 -39 ( 1 -36 ( 1 -40 ( 3 -38 ( 3

-104 ( 9 -138 ( 1 -118 ( 3 -108 ( 3 -118 ( 8 -109 ( 8

-1.2 ( 0.1 -2.6 ( 0.1 -2.7 ( 0.1 2.6 ( 0.1 -3.6 ( 0.1 -3.8 ( 0.5

48.9 56.0 59.9 60.6 67.1 73.0

11.7 11.1 13.1

a Structures are shown in Chart 2. b Determined by Meltwin56 from the UV absorption at 260 nm during the thermal dissociation in aqueous 10 mM sodium phosphate buffer containing 0.1 M NaCl. Errors are standard deviation errors between heating and cooling. c Difference between strand with and without pyrene.

TABLE 2: Fluorescence Quantum Yields and Decay Times for Pya-H and Pya-DNA Conjugates pyrenea

Φflb

Pya-H Pya-An Pya-1G Pya-2G Pya-3G Pya-4G Pya-AT

0.30 0.003 0.022 0.082 0.36 0.36 0.38

τs, ns (amp)c 37 8.6 2 (35), 9.6 (65) 2 (40), 10.4 (60) 2 (42), 10.3 (42) 3.9 (27), 10.3 (42)

a Structures shown in Chart 2. b Fluorescence quantum yields determined using quinine sulfate as an actinometer.29 c Fluorescence decay times for Pya-H and Pya-An in methanol and for Pyd conjugates in aqueous buffer (ca. 5 µM conjugate in 10 mM sodium phosphate, pH 7.2 with 0.1 M NaCl). Fast decay components are not resolved from the lamp profile for Pya-1G and are poorly resolved for the other Pya-nG conjugates.

are broadened and red-shifted by several nanometers, when compared to the spectra of Pya-H in methanol. Fluorescence quantum yields (Φfl) and decay times for Pya-H, Pya-An, and the Pya-nG conjugates are reported in Table 2. Values of Φfl for Pya-H and Pya-AT, and Pya-nG (n g 3) are all between 0.30 and 0.38. Lower values of Φfl are observed for Pya-1G, Pya-2G, and Pya-An. The fluorescence decay of Pya-H is best fit as a single exponential with τs ) 37 ns. Its fluorescence quantum yield (Table 2) is similar to that of pyrene (Φfl ) 0.32); however, its singlet lifetime is significantly shorter than that of pyrene (τs ) 450 ns).29 Rate constants for both radiative and nonradiative decay are increased by electronic interactions with the carboxamide substituents, as is the case for other conjugated pyrene derivatives.9,39 In the case of the very weakly fluorescent Pya-1G only a single fluorescence decay component is resolved. The fluorescence decays of other conjugates are best fit as dual exponentials having a shorter-lived component with τs ∼ 2 ns and a longer-lived component with τ ∼ 10 ns (Table 2). The circular dichroism spectrum of Pya-AT (Figure S2, Supporting Information) consists of a weak broad negative band around 350 nm attributed to induced CD of the pyrene chromophore and strong bands between 200-300 nm similar to those for the duplex poly(dA):poly(dT) and thus attributed to the base pair domain.40 The low values of Φfl for Pya-An, Pya-1G, and Pya-2G are attributed to electron transfer quenching of Pya singlet by aniline and guanine, respectively. The energetics of photoinduced electron transfer for the Pya-An dyad can be estimated using Weller’s equation (eq 1),

∆Get)-Es - (Erdn - Eox) + (2.6/ε - 0.13 eV)

(1)

where Es and Erdn are the singlet energy and reduction potential of Pya-H (+3.30 eV and -1.95 V, respectively), Eox is the

Figure 2. Transient absorption spectra for (a) Pya-H (0-5 ps) and (b) Pya-An (0-100 ps) in methanol solution. Blue spectra are earlier times (0-1 ps) and red spectra later times (>10 ps).

oxidation potential of N,N-dialkylaniline (-0.78 V), and the final term is an empirical correction for the solvent-dependent Coulomb attraction energy.41 The estimated value of ∆Get for Pya-An is -0.6 eV. Similarly, the estimated values of ∆Get for oxidation of guanine and adenine (Eox ) 1.24 and 1.69 eV for the nucleosides in acetonitrile vs SCE42) are 0.0 ( 0.1 V and 0.5 ( 0.1 V, respectively. Reduction of thymine is estimated to be endergonic by ca. 0.4 ( 0.1 V using the oxidation potential of Pya-H (1.53 V) and the reduction potential of T (-2.26 V). The calculated values of ∆Get are consistent with the observation of extensive fluorescence quenching for Pya-An and Pya-1G and the absence of fluorescence quenching for Pya-AT (Table 2). Femtosecond Pump-Probe Spectra. Pump-probe measurements employed a Ti-Sp-based system with a white-light continuum, which provides a usable probe source between 300 or 350 and 750 nm with a spectral resolution of ca. 10 nm and a time resolution of ca. 150 fs.13,43 The transient absorption spectra of Pya-H and Pya-An (Figure 2) display formation of a band at ca. 575 nm attributed to pyrene 2S* during the exciting laser pulse. Rapid decay of this band within the first picosecond following excitation is accompanied by the formation of two bands at shorter wavelengths assigned to pyrene 1S*. The appearance of the transient spectra assigned to 2S* and 1S* and the occurrence of ultrafast internal conversion are similar to the behavior reported for pyrene and several of its derivatives.39,44 The transient spectrum of Pya-H assigned to pyrene 1S* (Figure 2a) does not decay appreciably during the 1.9 ns time window of our measurements, in accord with its long fluorescence decay time (Table 2). The transient absorption spectra of

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Siegmund et al. TABLE 3: Assignments and Decay Times for Pya-H, Pya-An, and Pya-DNA Conjugates Obtained from Transient Absorption Dataa pyreneb Pya-H Pya-An Pya-AT Pya-1G Pya-2G Pya-3G Pya-4G Pya-5G

1

S* relax, ps

4.1, 34 5.9 3.4 3.1, 15 3.5, 24 3.0, 18

1

S* decay, ps >2000 73 >2000 65c 48c >2000 >2000 >2000

Pya•- rise (decay), ps 102 (>2000)

a Assignments based on band shape analysis as described in the text. Decay times obtained from single wavelength transient decays and preexponentials reported in Table S2, Supporting Information. b Structures shown in Chart 2. c Absence of Pya•- transient absorption attributed to inverted kinetics.

Figure 3. Transient absorption spectra (a) of conjugates Pya-AT (0-5 ps) and Pya-2G, (b) at 0-10 ps, and (c) at 0.2-1.9 ns, in aqueous buffer. Blue spectra are earlier times, and red spectra, later times.

Pya-An (Figure 2b) display the delayed growth of a narrow band at 510 nm attributed to the anion radical Pya•- on the basis of comparison with the transient absorption spectra reported for an alkane-linked pyrene-aniline dyad45 and a urea-linked pyreneaniline dyad.39 The formation and decay of the Pya•- 510 nm band (Figure S4, Supporting Information) provide values of τcs ∼ 100 ps and τcr > 2 ns (Tables S2 (Supporting Information) and 3). The transient absorption spectra of the Pya conjugates PyaAT, Pya-2G, and Pya-4G (Figures 3 and S3 (Supporting Information)) are broader than those of the dyad Pya-An and are assigned to Pya 1S*. Their 525 nm bands sharpen during the first 100 ps following the laser pulse, resulting in an increase in peak intensity attributed to structural and solvent relaxation of 1S*. The rises for Pya-1G and Pya-2G can be fit as single exponentials with τrise ∼ 5 ps (Tables S2 (Supporting Information) and 3). The rises for the other conjugates are best fit as dual exponentials with τrise ca. 4 and 20 ps. The transient absorption spectra of Pya-AT, Pya-3G, and Pya-4G (Figures 3, S3, and S4 (Supporting Information)) do not decay appreciably during the 1.9 ns time window of our measurements, in accord with their relatively long fluorescence decay times and large fluorescence quantum yields (Table 2). More rapid transient decay is observed for Pya-1G and Pya-2G (Figure S4, Supporting Information), in accord with their lower fluorescence quantum yields (Table 2). The absence of a narrow 510 nm Pya•- band similar to that observed for Pya-An (Figure 2b) is indicative of charge recombination occurring with a rate constant comparable to or faster that for charge separation (inverted kinetics). Inverted kinetics have previously reported for a covalently linked adduct of benzo[a]pyrene diol epoxide and guanine46 and for perylene- and pyrenedicarboxamide-linked DNA hairpins having an adjacent G-C base pair.4,11 The dominant 345 and 525 nm transient decay components for Pya-

1G and Pya-2G (ca. 65 and 45 ps, Table 3) are attributed to charge separation (formation of Pya•-). Faster charge separation for Pya-2G vs Pya-1G is contrary to the distance dependence expected for intramolecular photoinduced charge separation and led us to question the assumption that the Pya conjugates adopt capped hairpin structures. Influence of Pyrene on the 1D 1H NMR Spectra. To gain insight into the influence of the pyrene moiety on the structures of the Pya conjugates, one-dimensional (1D) 1H NMR were acquired for Pya-AT, Pya-1G, and Pya-2G and for the reference hairpins AT, 1G, and 2G (Chart 2b,c). The aromatic and deoxyribose regions of the 1D spectra are highly congested, as shown for Pya-2G in Figure S5, Supporting Information. However, the thymine-methyl protons are the only protons with chemical shifts in the region from 1.2 to 1.8 ppm, permitting interpretation of the 1D spectra in this region (Figure 4). Selective line-broadening for the signals of protons close to the pyrene moiety is observed for all of the Pya conjugates, whereas no such broadening is observed for the corresponding signals in the reference hairpins. For the hairpins 1G and 2G, the methyl proton signal of the T connected 3′ to guanosine can readily be assigned on the basis of their chemical shifts.47 In Pya-1G, the peak for the methyl protons of thymidine T2 broadens considerably at temperatures below 30 °C, although this methyl group would be separated by one GC base-pair from an end-capping pyrene. The broadening is much more pronounced than that of the signal of T1-methyl in Pya-AT or Pya2G. For Pya-AT, line broadening is less predominant and its observation is hampered by overlapping peaks at lower temperatures. The methyl chemical shift changes induced by incorporation of the pyrene moiety are relatively small, with values of ∆ ppm e 0.1 ppm. Imino and amino protons of the nucleobases can be assigned from the imino region of the NOESY spectra of Pya-2G in 90:10 water:D2O, with water suppression employing the WATERGATE pulse sequence (Figure 5).31 The 1D spectra of PyaAT, Pya-1G, and Pya-2G all display signals for five of the six exchangeable imino protons. These signals become sharper with increasing distance between the corresponding protons and the pyrene moiety. The H3 imino proton of the thymidine groups T1 in Pya-AT and Pya-2G and the H1 imino proton of G1 in Pya-1G could not be observed, presumably because of rapid exchange with water. This suggests that the terminal base-pair is not protected from water by a pyrene capping group. Two-Dimensional NMR Spectra of Pya-2G. The 2D spectra of Pya-2G were recorded at 25, 30, and 40 °C, temperatures well below the hairpin Tm of 73 °C (Table 1). The pyrene

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Figure 4. Methyl region of the 1D NMR spectra at temperatures between 10 and 70 °C of Pya-AT, Pya-1G, Pya-2G, and their unmodified controls AT, 1G, and 2G. Assignments are written above the spectrum at 20 °C for Pya-2G (from 2D-spectra) and Pya-1G (assigned from chemical shift and line broadening data). The triplet at 1.3 ppm present in some of the spectra is caused by residual triethylamine used in HPLC purification.

Figure 5. Imino region of Pya-AT, Pya-1G, and Pya-2G at 25 °C in 90:10 D2O:H2O. Assignments are given next to the peaks. Assignments for Pya-2G were made from information in 2D spectra; assignments of the peaks in Pya-AT and Pya-1G are based on their chemical shifts and line broadening.

protons exhibit significantly sharper peaks at 40 °C than at lower temperatures, unlike protons remote from pyrene. Spectra at lower temperatures also avoid the increased T1 noise experienced at 40 °C and take advantage of chemical shift changes. Two-dimensional spectra in 9:1 water:D2O were taken at 25 °C because the signal intensity of the amino and imino protons is much reduced at higher temperatures (Figure S6, Supporting Information). A complete assignment of the Pya-2G DNA backbone could be obtained from NOESY, TOCSY, and COSY spectra using H1′-H6/H8 connectivities (Figure 6a),48 along the H6/H8 and the thymidine methyl groups (Figure 6b), as well as along H2′/ H2′′ and H6/H8 (data not shown). Neither all pyrene protons nor G2H2 could be successfully assigned because of their overlapping resonances at 40 °C and increased line width at lower temperatures. Comparison of the spectra at different temperatures reveals five of the six expected TOCSY peaks for the pyrene moiety. A tentative assignment was made on the basis of the spectrum obtained at 30 °C (Figure S7, Supporting Information); however,

Figure 6. (a) Intersection between signals of the thymidine methyl groups and the aromatic protons for Pya-2G. Due to the presence of several sequential thymidines in the sequence, an alternative NOESY walk is possible along the methyl and aromatic protons for an oligonucleotide in B-form DNA. (b) Intersection between signals of the aromatic protons and H1′ protons of the ribose moieties. Sequence of peaks, the “NOESY walk” is connected by lines. The cross peaks of a TOCSY spectrum are drawn in blue above the red NOESY cross peaks. Assignments are shown in the form of the nucleotide name and number next to the intranucleotide cross peaks. Cross peaks between the pyrene moiety P0 and oligonucleotide H1′ protons are indicated by boxes.

no assignment consistent with the signals observed in the spectra at all temperatures was found. Signals for the pyrene moiety are found within a 0.5 ppm spectral region, with eight out of the nine protons in an even narrower 0.22 ppm region, a much narrower region than for other reported pyrene-DNA conjugates.49,50

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Figure 8. (a) Structures of Pya-2G with pyrene intercalated between first and second base pair calculated with a constraints from NOE data. Left: side view with the base pair G2-C14 shown on top (drawn with filled bases) with the pyrene and base pair T1-A14 below. The rest of the double strand is omitted for clarity. Right: end-on view of the same conformation. (b) Structures of Pya-2G with an end-capped pyrene generated by omitting NMR constraints between the pyrene and the G2-C14 base pair. For clarity only the first two base pairs are shown: left, viewed from the side, and right, viewed from the end.

Figure 7. View of the conjugate Pya-2G with intercalated pyrene. The trace of the backbone is shown as a transparent tube.

The line broadening for protons in proximity to the pyrene moiety and the narrow range of pyrene proton chemical shifts suggest that the pyrene adopts multiple conformations within the NMR time scale, where the averaging between conformations causes the chemical shifts of all pyrene protons to converge to similar values. Such averaging can also explain the small influence of the pyrene moiety on methyl chemical shifts (Figure 4) when compared to the reference hairpins AT, 1G, and 2G. All cross peaks between the pyrene moiety and the oligonucleotide are weak compared with the internucleotide cross peaks used for the sequential assignment. The pyrene moiety shows cross peaks to the terminal and to the penultimate base pairs, indicating that it is intercalated between these base pairs in its major conformation. Spectra taken in 9:1 water:D2O at 25 °C display NOESY cross-peaks of equal intensity from the amide NH of the linker to resonances at 7.85 and 7.52 assigned to pyrene protons H1 and H9 (Chart 2a). This suggests that the pyrene-amide bond exists as a mixture of syn and anti pyrene-carbonyl rotamers (the rotamer with N-H syn to H1 is shown in Chart 2a). A structure for Pya-2G calculated using a subset of the constraints from the NMR data is shown in Figure 7. In this structure pyrene is intercalated between the terminal A-T base pair and the G-C base pair. The structure of the hairpin loop region is similar to that reported for a related mini-hairpin.36 End-on and side views of pyrene and the two terminal base pairs are shown in Figure 8a. These structures show good π-overlap between pyrene with T1 and G2 but only partial overlap with bases in the complementary strands. Similar structures can be generated by flipping of the pyrenecarboxamide and rotation about the pyrene-carbonyl bond. Alternative endcapped structures were generated by using NMR constraints between the pyrene and the terminal base pair, but not the penultimate base pair. End-on and side views of an end-capped structure are shown in Figure 8b. In this structure one of the

hydrophobic faces of pyrene is exposed to water and the other face shows only partial overlap with the terminal A-T base pair. Omission of constraints to the penultimate base-pair should favor end-capped vs intercalated structures. However, the molecular dynamics calculations also yielded some low-energy intercalated structures. We have not investigated the structures of Pya-AT and Pya-1G but assume they are similar to that for Pya-2G on the basis of similarities in their thermodynamic parameters (Table 1) and 1D NMR spectra. Comparisons with Related Conjugate Structures. As mentioned previously, Pya was intended to serve as a hydrophobic hairpin capping group similar to the pyrenedeoxyribonucleoside single base overhangs studied by Kool and co-workers.51 Tuma et al. have reported that a duplex possessing tethered stilbenecarboxamide has a capped hairpin structure (Chart 1c).26 Unlike the case of Pya-nG in which a fast exchange leads to line broadening, the exchange between the stilbene conformations is slow, resulting in two sets of peaks in the NMR spectra. In our case, the broadened lines at 600 MHz indicate that exchange occurs on the millisecond time scale.52 We note the magnitude of duplex stabilization by tethered Pya (Table 1) is much smaller than that reported for either the pyrene nucleosides or tethered stilbenecarboxamide, both of which adopt end-capped structures.26,51 Conjugates possessing 5′-tethered 1-alkylpyrenes employed in studies of DNA electron transfer and as DNA hybridization probes have been assumed to adopt capped duplex structures.22,53 However, Richert et al. have recently suggested that duplexes formed by conjugates possessing a 3′-(pyrenylmethyl)deoxyadenosine adopt intercalated structures.38 This group has also reported a 5′-tetherred oxolinic acid and a 2′-linked nalidixic acid moiety do not act as duplex end-caps but instead disrupt the terminal base pair, capping the penultimate base pair and displacing an adenine or uracil nucleobase into the minor groove.27 There are several reported NMR structures for duplexes formed by conjugates possessing an intercalated, covalently attached pyrene. Smirnov et al. reported the structures of two duplexes containing a pyrene deoxyriboside on one strand and an abasic site on the other.49 In one of these the pyrene nucleotide is in the middle of the strand and indeed replaces

Electron Transfer in DNA Hairpin Conjugates the base pair. The other duplex has a terminal pyrene nucleotide on one strand and a tetranucelotide overhang on the other. In this duplex the pyrene serves as a capping group but is protected from water by the overhang. Nakamura et al. have reported an interesting duplex structure in which (hydroxymethyl)pyrene is linked to the 2′-ribose of a midstrand uracil.50 The adjacent bases remain paired even though the duplex structure is distorted by the intruding intercalated pyrene. The solution NMR structures of several duplexes possessing adducts formed from the reaction of the carcinogen 9,10-epoxy7,8,9,10-tetrahydrobenzo[a]pyrene with the amino group of adenosine or guanine have also been reported.54 In most of these structures pyrene is intercalated next to the base pair to which it is attached. These duplexes exhibit intact, if severely distorted base pairs, despite their very short linker to pyrene, an unfavorable point of attachment, and in some cases purinepurine mismatches. The nature of these structures indicates that there is a very strong driving force for pyrene to intercalate while retaining largely intact base pairs, even in cases involving a base pair mismatch. Concluding Remarks. Our NMR investigation of the solution structure of the conjugate Pya-2G indicates that it forms a hairpin structure in which the tethered Pya is intercalated between the first two base pairs rather than capping the terminal base pair. This structure is similar to those of two other 5′tethered aromatics27 but is different from that of a tethered 5′stilbenecarboxamide which forms a capped hairpin structure.26 We suspect the difference in geometry for the pyrene vs stilbene conjugates is a consequence of the larger hydrophobic surface for pyrene and its more compact dimensions, which result in poorer overlap with a terminal base pair than is possible with the more elongated stilbene. These results call into question the usual assumption of capped duplex structures for 5′- and 3′tethered pyrenes and other aromatic chromophores without explicit structural evidence. Our structural studies also provide a plausible explanation for the unusual charge separation dynamics for Pya-nG conjugates (Table 3). Intercalated and end-capped structures for Pya-2G can account, respectively, for the fast (major) and slow (minor) components of transient decay assigned to charge separation. The absence of transient lifetime or fluorescence intensity quenching at longer distances (Pya-nG, n g 3) indicates that effective quenching is observed only for nearest neighbor guanines. Both capping and intercalated structures for Pya-1G provide nearest neighbor interactions with guanine. The small components of longer-lived transient decays and low fluorescence quantum yields for Pya-1G and Pya-2G may arise from minor conformations. Multiple conformations can also account for the spectral broadening observed in the absorption, fluorescence, and transient absorption spectra of the Pya conjugates. Finally, it is important to note the synergy between the photophysical and structural components of this investigation. Photophyscial studies provided results that were not consistent with the normal distance dependence expected for photoinduced electron transfer in what was initially assumed to be a cappedhairpin structure. These studies also suggested that Pya-1G and Pya-2G adopt a major and minor kinetically nonequivalent structures. Our structural studies revealed an unexpected preference for intercalated structures in which the tethered pyrene is located between the two terminal base pairs. These studies also indicate the likely presence of minor structures with interconversion on the NMR time scale, which is much slower than the picosecond time scale of the electron transfer processes. The

J. Phys. Chem. B, Vol. 113, No. 50, 2009 16283 synergy between these studies motivated our decision to incorporate both into one report rather than separate them, as has been done in the few cases where structures have been obtained for intercalated or tethered chromophores employed in studies of DNA electron transfer.9,55 Acknowledgment. We thank Arun K. Thazhathveetil for assistance in the revision of this manuscript. This research is supported by grants from the Chemical Sciences, Geosciences and Biosciences Division, Office of Basic Energy Sciences, Office of Science, U.S. Department of Energy (DE-FG0296ER14604 to FDL) and the National Science Foundation (CHE-0628130 to TF). Supporting Information Available: Two tables containing MALDI-TOF data and single wavelength transient data and seven figures containing conjugate fluorescence, CD, and transient absorption spectra, single wavelength transient traces, 1D NMR and 2D NOSEY spectra for Pya-2G. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Letsinger, R. L.; Wu, T. J. Am. Chem. Soc. 1995, 117, 7323–7328. (2) Lewis, F. D.; Liu, X.; Wu, Y.; Miller, S. E.; Wasielewski, M. R.; Letsinger, R. L.; Sanishvili, R.; Joachimiak, A.; Tereshko, V.; Egli, M. J. Am. Chem. Soc. 1999, 121, 9905–9906. (3) (a) Manetto, A.; Breeger, S.; Chatgilialoglu, C.; Carell, T. Angew. Chem., Int. Ed. 2006, 45, 318–321. (b) Ito, T.; Hayashi, A.; Kondo, A.; Uchida, T.; Tanabe, K.; Yamada, H.; Nishimoto, S. Org. Lett. 2009, 11, 927–930. (4) Lewis, F. D.; Zhang, L.; Kelley, R. F.; McCamant, D.; Wasielewski, M. R. Tetrahedron 2007, 63, 3457–3464. (5) (a) Letsinger, R. L.; Wu, T. J. Am. Chem. Soc. 1994, 116, 811– 812. (b) Langenegger, S.; Ha¨ner, R. Chem. Commun. 2004, 2792–2793. (c) Zheng, Y.; Long, H.; Schatz, G. C.; Lewis, F. D. Chem. Commun. 2005, 4795–4797. (d) Bayer, J.; Radler, J. O.; Blossey, R. Nano Lett. 2005, 5, 497–501. (6) (a) Bevers, S.; O’Dea, T. P.; McLaughlin, L. W. J. Am. Chem. Soc. 1998, 120, 11004–11005. (b) Rahe, M.; Rinn, C.; Carell, T. Chem. Commun. 2003, 2120–2121. (c) Trkulja, I.; Ha¨ner, R. Bioconj. Chem. 2007, 18, 289–292. (d) Trkulja, I.; Ha¨ner, R. J. Am. Chem. Soc. 2007, 129, 7982– 7989. (7) Malinovskii, V. L.; Samain, F.; Ha¨ner, R. Angew. Chem., Int. Ed. 2007, 46, 4464–4467. (8) (a) Lewis, F. D.; Letsinger, R. L.; Wasielewski, M. R. Acc. Chem. Res. 2001, 34, 159–170. (b) Takada, T.; Kawai, K.; Fujitsuka, M.; Majima, T. Angew. Chem., Int. Ed. 2006, 45, 120–122. (9) Egli, M.; Tereshko, V.; Mushudov, R.; Sanishvili, R.; Liu, X.; Lewis, F. D. J. Am. Chem. Soc. 2003, 125, 10842–10849. (10) Tuma, J.; Tonzani, S.; Schatz, G. C.; Karaba, A. H.; Lewis, F. D. J. Phys. Chem. B 2007, 111, 13101–13106. (11) Daublain, P.; Siegmund, K.; Hariharan, M.; Vura-Weis, J.; Wasielewski, M. R.; Lewis, F. D.; Shafirovich, V.; Wang, Q.; Raytchev, M.; Fiebig, T. Photochem. Photobiol. Sci. 2008, 7, 1501–1508. (12) (a) Telser, J.; Cruickshank, K. A.; Morrison, L. E.; Netzel, T. L. J. Am. Chem. Soc. 1989, 111, 6966–6976. (b) Gaballah, S. T.; Collier, G.; Netzel, T. L. J. Phys. Chem. B 2005, 109, 12175–12181. (13) Kaden, P.; Mayer-Enthart, E.; Trifonov, A.; Fiebig, T.; Wagenknecht, H.-A. Angew. Chem., Int. Ed. 2005, 44, 1636–1639. (14) (a) Kool, E. T. Chem. ReV. 1997, 97, 1473–1487. (b) Wilson, J. N.; Kool, E. T. Org. Biomol. Chem. 2006, 4, 4265–4274. (15) Zhang, L.; Zhu, H.; Sajimon, M. C.; Stutz, J. A. R.; Siegmund, K.; Richert, C.; Shafirovich, V.; Lewis, F. D. J. Chinese Chem. Soc. (Taipei) 2006, 53, 1501–1507. (16) (a) Amann, N.; Wagenknecht, H. A. Tetrahedron Lett. 2003, 44, 1685–1690. (b) Ito, T.; Rokita, S. E. J. Am. Chem. Soc. 2003, 125, 11480– 11481. (c) Wagner, C.; Wagenknecht, H. A. Org. Lett. 2006, 8, 4191– 4194. (d) Baumstark, D.; Wagenknecht, H. A. Angew. Chem., Int. Ed. 2008, 47, 2612–2614. (e) Fukui, K.; Tanaka, T. Angew. Chem., Int. Ed. 1998, 37, 158–161. (17) (a) Fukui, K.; Tanaka, K.; Fujitsuka, M.; Watanabe, A.; Ito, O. J. Photochem. Photobiol. B 1999, 50, 18–27. (b) Wan, C.; Fiebig, T.; Schiemann, O.; Barton, J. K.; Zewail, A. H. Proc. Natl. Acad. Sci. U.S.A. 2000, 97, 14052–14055. (c) Zeidan, T. A.; Carmieli, R.; Kelley, R. F.; Wilson, T. M.; Lewis, F. D.; Wasielewski, M. R. J. Am. Chem. Soc. 2008, 130, 13945–13955. (d) Hess, S.; Go¨tz, M.; Davis, W. B.; Michel-Beyerle, M. E. J. Am. Chem. Soc. 2001, 123, 10046–10055.

16284

J. Phys. Chem. B, Vol. 113, No. 50, 2009

(18) (a) Murphy, C. J.; Arkin, M. R.; Ghatlia, N. D.; Bossmann, S.; Turro, N. J.; Barton, J. K. Proc. Natl. Acad. Sci. U.S.A. 1994, 91, 5315– 5319. (b) Kawai, K.; Majima, T. Top. Cur. Chem. 2004, 236, 117–137. (19) Schuster, G. B. Acc. Chem. Res. 2000, 33, 253–260. (20) Erkkila, K. E.; Odom, D. T.; Barton, J. K. Chem. ReV. 1999, 99, 2777–2795. (21) Assetline, U.; Delarue, M.; Lancelot, G.; Toulme´, F.; Thuong, N. T.; Montenaygarestier, T.; He´le`ne, C. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 3297–3301. (22) Zahavy, E.; Fox, M. A. J. Phys. Chem. B 1999, 103, 9321–9327. (23) Lewis, F. D.; Zhang, L.; Liu, X.; Zuo, X.; Tiede, D. M.; Long, H.; Schatz, G. S. J. Am. Chem. Soc. 2005, 127, 14445–14453. (24) Kawai, K.; Kodera, H.; Osakada, Y.; Majima, T. Nature Chem. 2009, 1, 156–159. (25) Mann, J. S.; Shibata, Y.; Meehan, T. Bioconjugate Chem. 1992, 3, 554–558. (26) Tuma, J.; Paulini, R.; Stu¨tz, J. A. R.; Richert, C. Biochemistry 2004, 43, 15680–15687. (27) (a) Tuma, J.; Connors, W. H.; Stitelman, D. H.; Richert, C. J. Am. Chem. Soc. 2002, 124, 4236–4246. (b) Siegmund, K.; Maheshwary, S.; Narayanan, S.; Connors, W.; Riedrich, M.; Printz, M.; Richert, C. Nucleic Acids Res. 2005, 33, 4838–4848. (28) Lewis, F. D.; Zhu, H.; Daublain, P.; Fiebig, T.; Raytchev, M.; Wang, Q.; Shafirovich, V. J. Am. Chem. Soc. 2006, 128, 791–800. (29) Berlman, I. B. Handbook of Fluorescence Spectra of Aromatic Molecules, 2nd ed.; Academic Press: New York, 1971. (30) Astruc, D. In Electron Transfer in Chemistry; Balzani, V., Ed.; Wiley-VCH: Weinheim, 2001; Vol. 2. (31) Piotto, M.; Saudek, V.; Sklena´r, V. J. Biomol. NMR 1992, 2, 661– 665. (32) Wijmenga, S. S.; van Buuren, B. N. M. Prog. Nucl. Magn. Reson. Spectrosc. 1998, 32, 287–387. (33) Bru¨nger, A. T.; Adams, P. D.; Clore, G. M.; DeLano, W. L.; Gros, P.; Grosse-Kunstleve, R. W.; Jiang, J. S.; Kuszewski, J.; Nilges, M.; Pannu, N. S.; Read, R. J.; Rice, L. M.; Simonson, T.; Warren, G. L. Acta Crystallogr. Sect. D-Biol. Crystallogr. 1998, 54, 905–921. (34) Kleywegt, G. J.; Jones, T. A. Macromol. Cryst. Pt B 1997, 277, 208–230. (35) Siegmund, K. Diplomarbeit, Konstanz, 2001. (36) Yoshizawa, S.; Kawai, G.; Watanabe, K.; Miura, K.; Hirao, I. Biochemistry 1997, 36, 4761–4767. (37) (a) Markham, N. R.; Zuker, M. Nucleic Acids Res. 2005, 33, W577– W581. (b) Markham, N. R.; Zuker, M. In Bioinformatics, Volume II. Structure, Functions and Applications; Keith, J. M., Ed.; Humana Press: Totowa, NJ, 2008; pp 3-31. (38) Printz, M.; Richert, C. Chem.sEur. J. 2009, 15, 3390–3402. (39) Lewis, F. D.; Daublain, P.; Delos Santos, G.; Liu, W.; Asatryan, A. M.; Markarian, S. A.; Fiebig, T.; Raytchev, M.; Wang, Q. J. Am. Chem. Soc. 2006, 128, 791–800. (40) Johnson, W. C. In Circular Dichroism, Principles and Applications; Berova, N., Nakanishi, K., Woody, R. W., Eds.; Wiley-VCH: New York, 2000; pp 741-768.

Siegmund et al. (41) Weller, A. Z. Phys. Chem. Neue. Folg. 1982, 133, 93–98. (42) Seidel, C. A. M.; Schulz, A.; Sauer, M. H. M. J. Phys. Chem. 1996, 100, 5541–5553. (43) Raytchev, M.; Mayer, E.; Amann, N.; Wagenknecht, H. A.; Fiebig, T. Chem. Phys. Chem. 2004, 5, 706–712. (44) Raytchev, M.; Pandurski, E.; Buchvarov, I.; Modrakowski, C.; Fiebig, T. J. Phys. Chem. A 2003, 107, 4592–4600. (45) Okada, T.; Migita, M.; Mataga, N.; Sakata, Y.; Misumi, S. J. Am. Chem. Soc. 1981, 103, 4715–4720. (46) O’Conner, D.; Shafirovich, V. Y.; Geacintov, N. E. J. Phys. Chem. 1994, 98, 9831–9839. (47) Arter, D. B.; Schmidt, P. G. Nucleic Acids Res. 1976, 3, 1437– 1447. (48) Scheek, R. M.; Boelens, R.; Russo, N.; van Boom, J. H.; Kaptein, R. Biochemistry 1984, 23, 1371–1376. (49) Smirnov, S.; Matray, T. J.; Kool, E. T.; de los Santos, C. Nucleic Acids Res. 2002, 30, 5561–5569. (50) Nakamura, M.; Fukunaga, Y.; Sasa, K.; Ohtoshi, Y.; Kanaori, K.; Hayashi, H.; Nakano, H.; Yamana, K. Nucleic Acids Res. 2005, 33, 5887– 5895. (51) Ren, R. X. F.; Chaudhuri, N. C.; Paris, P. L.; Rumney, S.; Kool, E. T. J. Am. Chem. Soc. 1996, 118, 7671–7678. (52) Millet, O.; Loria, J. P.; Kroenke, C. D.; Pons, M.; Palmer, A. G. J. Am. Chem. Soc. 2000, 122, 2867–2877. (53) Narayanan, S.; Gall, J.; Richert, C. Nucleic Acids Res. 2004, 32, 2901–2911. (54) (a) Feng, B.; Gorin, A.; Hingerty, B. E.; Geacintov, N. E.; Broyde, S.; Patel, D. J. Biochemistry 1997, 36, 13769–13779. (b) Feng, B.; Gorin, A.; Kolbanovskiy, A.; Hingerty, B. E.; Geacintov, N. E.; Broyde, S.; Patel, D. J. Biochemistry 1997, 36, 13780–13790. (c) Gu, Z.; Gorin, A.; Krishnasamy, R.; Hingerty, B. E.; Basu, A. K.; Broyde, S.; Patel, D. J. Biochemistry 1999, 38, 10843–10854. (d) Mao, B.; Gu, Z.; Gorin, A.; Chen, J.; Hingerty, B. E.; Amin, S.; Broyde, S.; Geacintov, N. E.; Patel, D. J. Biochemistry 1999, 38, 10831–10842. (e) Pradhan, P.; Tirumala, S.; Liu, X.; Sayer, J. M.; Jerina, D. M.; Yeh, H. J. Biochemistry 2001, 40, 5870– 5881. (f) Volk, D. E.; Rice, J. S.; Luxon, B. A.; Yeh, H. J.; Liang, C.; Xie, G.; Sayer, J. M.; Jerina, D. M.; Gorenstein, D. G. Biochemistry 2000, 39, 14040–14053. (g) Volk, D. E.; Thiviyanathan, V.; Rice, J. S.; Luxon, B. A.; Shah, J. H.; Yagi, H.; Sayer, J. M.; Yeh, H. J. C.; Jerina, D. M.; Gorenstein, D. G. Biochemistry 2003, 42, 1410–1420. (55) (a) Gasper, S. M.; Armitage, B.; Shui, X. Q.; Hu, G. G.; Yu, C. J.; Schuster, G. B.; Williams, L. D. J. Am. Chem. Soc. 1998, 120, 12402– 12409. (b) Kielkopf, C. L.; Erkkila, K. E.; Hudson, B. P.; Barton, J. K.; Rees, D. C. Nature Struct. Biol. 2000, 7, 117–121. (c) von Feilitzsch, T.; Tuma, J.; Neubauer, H.; Verdier, L.; Haselsberger, R.; Feick, R.; Gurzadyan, G.; Voityuk, A. A.; Griesinger, C.; Michel-Beyerle, M. E. J. Phys. Chem. B 2008, 112, 973–989. (56) McDowell, J. A. MeltWin 3.5, 2001.

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