A Sensitive Multispectroscopic Probe for Nucleic Acids - The Journal

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7958

J. Phys. Chem. B 2010, 114, 7958–7966

A Sensitive Multispectroscopic Probe for Nucleic Acids Xin Sonia Gai, Edward E. Fenlon,* and Scott H. Brewer* Department of Chemistry, Franklin & Marshall College, Lancaster, PennsylVania 17604-3003 ReceiVed: February 12, 2010; ReVised Manuscript ReceiVed: April 30, 2010

Azides have recently been used as vibrational probes of proteins, but their incorporation into nucleic acids has been limited to photo-cross-linking or click chemistry applications. The utility of 2′-azido-2′-deoxyuridine (N3-dU, 1) as an IR and 15N NMR spectroscopic probe of the sugar phosphate backbone region of nucleic acids was investigated by measuring the effects of solvent, heterodimer formation, and temperature on peak frequencies and IR bandwidth. The azide IR asymmetric stretching band (ν˜ N3) of N3-dU was sensitive to its environment, undergoing a blue shift of 13.5 cm-1 when changing the solvent from THF to water. The solvent effects on 15N chemical shifts (δ15N) of each of the nitrogen atoms in the azido group was studied, and the terminal nitrogen atom was the most sensitive to solvent, shifting downfield by 3.8 ppm when changing the solvent from THF-d8 to D2O. Formation of a base-pair-like heterodimer between 3 (a silyl ether analogue of 1) and 2,6-diheptanamidopyridine (4) in chloroform resulted in minimal changes in the IR and 15N NMR spectral frequency and chemical shift, respectively, as expected given the location of the azido moiety. The intrinsic temperature dependence of ν˜ N3 and δ15N were found to be minimal over the temperature range studied especially compared to the solvent dependence of these spectral observables. The analysis of the experimental studies was complemented by density functional theory (DFT) calculations on model systems. Introduction Azides (R-N3) have outstanding potential as vibrational probes of biomolecules.1-5 Several factors contribute to this, including (1) the azide IR absorption is narrow, intense, and in a clear region of the spectrum; (2) azides are sensitive to changes in local environment; (3) azides contain only three atoms, making them smaller than many other probes; and (4) aliphatic azides are stable under most conditions. The utility of an azido group as a spectral probe of environment is enhanced by 15N enrichment of the nitrogen atoms. Specifically, 15N enriched azides have the additional potential as NMR spectroscopic probes. Moreover, azides have the unique ability to undergo highly selective and useful “click chemistry”sHuisgen cycloaddition reactions with alkynes.6,7 Thus, click chemistry can be used to conjugate probes such as fluorescent dyes or nitroxide spin labels to biomolecules modified with an azido functional group. Two recent reports have shown the great potential of the azido group as an IR spectroscopic probe in peptides and proteins.3,5 The first of these utilized β-azidoalanine (N3-A), which exhibited great sensitivity to its environment as indicated by a solvent-induced blue shift of ∼14 cm-1 in going from DMSO to water. This azide was also shown to have an oscillator strength approximately 19 times greater than the corresponding nitrile-modified alanine unnatural amino acid. The Cho group3 further demonstrated the usefulness of this IR probe to monitor the environment of the aggregated amyloid β-peptide. The second study utilized p-azidophenylalanine (N3F) to investigate the light-induced conformational changes in the G protein-coupled receptor rhodopsin. Codon suppression was used to incorporate N3-F into five different positions in rhodopsin, and IR difference spectra of the dark minus photoproduct spectrum showed distinct azide frequency shifts among * Corresponding authors. E.E.F.: e-mail, [email protected]; phone, (717) 291-4201; fax, (717) 291-4343. S.H.B.: e-mail, scott.brewer@ fandm.edu; phone, (717) 358-4766; fax, (717) 291-4343.

the various sites. Polar to less polar transitions were observed for N3-F at position 250 (N3-F250), the reverse transition was found for N3-F227, and the environment around N3-F102 was essentially unchanged in going from dark to photoproduct, as expected. Other groups have used one- and two-dimensional IR (2D IR) of the azide anion to probe protein dynamics.9,10 Azide has also been used as an IR probe of methane monooxygenase11 and photosystem II.12 Nitrile probes8 represent smaller perturbations to a system and less conformational flexibility compared to azide probes; however, the greater sensitivity to environment and larger extinction coefficient of the azido group suggests that azide probes can rival the utility of nitrile probes. The choice between these two probes will depend upon the specific system of interest, but these studies suggest that azides should at least be considered as the spectroscopic probe of choice compared to nitriles. In terms of nucleic acids, azides have not been extensively employed perhaps because of incorporation difficulties. The obsolete phosphotriester oligonucleotide synthesis method is compatible with the azide functional group and short oligomers containing 2′-azido-2′-deoxyuridine (N3-dU, 1) have been prepared in this way.13 However, azides are incompatible with phosphorus(III) compounds such as phosphoramidites, presumably due to a Straudinger reduction.6,13,14 Thus, standard solidphase synthesis of DNA oligomers containing an azide is not possible. A satisfactory solution to this problem has yet to be devised. Early work by Evans and Haley15 showed that a triphosphate of 5-azidodeoxyuridine could be incorporated into DNA enzymatically; however, the DNA constructs were photoactive and unstable. Azide incorporation into oligomers prepared by the phosphoramidite method has been done by postsynthetic reaction of 5′-amino or amino-base modified oligomers14,16-18 with a tethered azide. The lengthy tethers employed placed the azide group between eight and 26 atoms away from the nucleoside. These tethered azides were subsequently used for photo-cross-linking17,18 or click chemistry.14,16

10.1021/jp101367s  2010 American Chemical Society Published on Web 05/24/2010

Sensitive Multispectroscopic Probe for Nucleic Acids

Figure 1. Thymidine-adenine base pair. R1 ) major groove probe; R2 ) phosphate sugar region probe.

None of these previous studies on azido-DNA used the azido group as a vibrational or 15N NMR spectroscopic probe. We have undertaken a program to develop minimally perturbative, multispectroscopic site-specific probes for all nucleic acid microenvironments; i.e., major groove, minor groove, and phosphate sugar region (Figure 1). Previous work has shown that the nitrile group of 5-cyano-2′-deoxyuridine (2) is an excellent vibrational2,19,20 and 15N NMR21 major grooVe probe, while the nitrile group of N2-cyano-2′-deoxyguanosine is an effective vibrational probe of the minor grooVe.2 In terms of a phosphate sugar region probe, the nitrile group may not be suitable because 2′-cyano-nucleosides are susceptible to backbone cleavage and/or base elimination due to the high acidity of the proton alpha to the nitrile group.22 The much lower acidity of protons alpha to an azido group makes 2′-azido-nucleosides23 and oligonucleotides13 stable. The greater environmental sensitivity and oscillator strength3 of azides relative to nitriles also strongly suggests that N3-dU has excellent potential as a phosphate sugar region probe. Our approach to azido-DNA uses the commercially available phosphoramidite of 2′-amino-2′-deoxyuridine24 to selectively incorporate a 2′-amino group(s) which can subsequently be converted to the desired 2′-azido group(s) by a postsynthetic diazotransfer reaction.25-27 Preliminary results in our laboratory indicate that this reaction is selective28-31 and will successfully incorporate azido groups into DNA and RNA. In this paper the utility of N3-dU (and its silyl ether analogue, 3) as a phosphate sugar region probe was investigated by IR and 15N NMR spectroscopy. The effects of solvent, heterodimer formation, and temperature were investigated experimentally and computationally with these two spectroscopic techniques and density functional theory (DFT) calculations, respectively. Experimental Section Synthetic Chemistry. General. All reagents were ACS reagent quality and were used without further purification unless otherwise noted. Sodium azide 15N labeled in the terminal position was purchased from Icon Isotopes (99%+). The following compounds were prepared according to literature procedures: 2′-amino-2′deoxyuridine (6) and 2,2′-anhydrouridine (7) by the method of McGee et al.32 and 2,6-diheptanamidopyridine (4) by the method of van Doorn et al.33 Synthetic methods for 2′-azido-2′-deoxyuridine (1) and 3′,5′-di-O-acetyl-2′-azido-2′-deoxyuridine (8) are known23,34 but alternate syntheses are provided. All reactions were stirred with a magnetic stir bar and conducted under a dry nitrogen or argon atmosphere. Analytical thin layer chromatography (TLC) was performed on 0.2 mm silica plastic coated sheets (Selecto Scientific) with F254 indicator. Flash column chromatography was performed on 230-400 mesh silica gel. NMR spectra for characterization were obtained at the following frequencies: 1H (499.7 MHz) and 13C (125.7 MHz) with a Varian INOVA 500 multinuclear Fourier transform NMR

J. Phys. Chem. B, Vol. 114, No. 23, 2010 7959 spectrometer. Chemical shifts are reported in parts per million (ppm), and coupling constants are reported in hertz (Hz). 1H spectra taken in CDCl3 were referenced to tetramethylsilane (TMS ) 0.0 ppm) as an internal standard. 13C NMR spectra taken in CDCl3 were referenced to the solvent peak at 77.0 ppm. IR spectra for characterization purposes were obtained with a Thermo-Nicolet Avatar 370 FTIR spectrometer with 4 cm-1 resolution and 16 scans using an attenuated total reflectance (ATR) attachment. The samples were evaporated on the ATR crystal resulting in a thin film. The frequencies are reported in wavenumbers (cm-1). Melting points were measured on a MelTemp melting point apparatus and are uncorrected. Abbreviations: DMAP (4-dimethylaminopyridine); DMF (dimethylformamide); ESI (electrospray ionization); FC (flash column chromatography using silica gel); HMPA (hexamethylphosphoramide); H2O (deionized water); MeOH (methanol). Procedures. Internal and Terminal Labeled (1:1) Trifluoromethanesulfonic Azide (Triflic Azide). 35 15N labeled sodium azide (83.5 mg, 1.46 mmol) was dissolved in CH3CN (1.5 mL), the mixture was cooled in an ice-water bath, and trifluoromethanesulfuric anhydride (285.6 mg, 1.01 mmol) was added dropwise. The mixture was stirred at low temperature for 2 h. The crude product solution was used for the next step without further purification. (CAUTION: Triflic azide should always be used as a solution due to its potentially explosive nature.) 15 N Middle Labeled and Unlabeled (1:1) 2′-Azido-2′-deoxyuridine (1m and 1). 2′-Amino-2′-deoxyuridine (6)32 (201.2 mg, 0.83 mmol) was dissolved in H2O (1.5 mL), and CH3CN (1.5 mL), K2CO3 (229.7 mg, 1.66 mmol), and CuSO4 · 5H2O (10.4 mg, 41.65 µmol) were added. The mixture was cooled in an ice-water bath, and the solution of 15N labeled trifluoromethanesulfuric azide (177.7 mg, 1.01 mmol) in CH3CN (1.5 mL) from above was added dropwise. The cold mixture was allowed to slowly warm to ambient temperature and was stirred for 21 h. The mixture was coevaporated with silica gel and purified by FC (10% MeOH/CHCl3) to give 182 mg (81%) of 1m/1 as a white foam. The spectral data of the product matched the literature values.23,34 15 N Internal and Terminal Labeled (1:1) 3′, 5′-Bis-O(acetyl)-2′-azido-2′-deoxyuridine (8i and 8t). 2,2′-Anhydrouridine (7)32 (211.7 mg, 0.94 mmol) was dissolved in HMPA (2.5 mL) (CAUTION: HMPA is a known carcinogen), and 15N labeled sodium azide (315.5 mg, 4.8 mmol) was added. The mixture was heated to 150 °C for 15 min, during which time most of the solids dissolved and then benzoic acid (127.2 mg, 1.04 mmol) (CAUTION: Toxic and explosive hydrazoic acid may be formed) was added. The mixture was stirred at 150 °C for 1 h. The reaction mixture quickly cooled to ambient temperature and diluted with H2O (10 mL) and extracted with CHCl3 (50 mL). The organic layer was back-extracted with H2O (2 × 10 mL), and the combined aqueous layers were washed with CHCl3 (5 × 25 mL). The aqueous layer was concentrated down under reduced pressure to give the crude product, which was then acetylated directly. The crude residue was dissolved in DMF (7 mL), and acetic anhydride (5 mL) and DMAP (121 mg, 0.98 mmol) were added. The mixture was stirred at ambient temperature for 24 h. The reaction mixture was concentrated under reduced pressure. The resultant solids were dissolved in CH2Cl2 and washed with H2O. The aqueous layer was backextracted with CH2Cl2 (2 × 10 mL). The combined organic layers were washed with H2O (3 × 10 mL), dried (Na2SO4), and then concentrated. The crude product was purified by FC (5% MeOH/CH2Cl2) to give 8i/8t as a light yellow gel. The spectral data of the product matched the literature values.23

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15

N Internal and Terminal Labeled (1:1) 2′-Azido-2′deoxyuridine (1i and 1t). 8i/8t from above was stirred overnight with NH3 in MeOH (7M, 25 mL), first in an ice-water bath and then at ambient temperature. The solvent was concentrated under reduced pressure and the crude product was purified by FC (10 f 20% MeOH/CHCl3) to give 146 mg (58% for three steps) of 1i/1t as a white foam. The spectral data of the product matched the literature values.23,34 3′,5′-Bis-O-(tert-butyldiphenylsilyl)-2′-azido-2′-deoxyuridine (3). 2′-Azido-2′-deoxyuridine (1) (117.0 mg, 0.43 mmol) was dissolved in DMF (2.0 mL) and tert-butyldiphenylchlorosilane (0.7 mL, 2.7 mmol) and imidazole (151.3 mg, 2.23 mmol) were added. The mixture was stirred at ambient temperature for 47 h. The reaction mixture was diluted with water and extracted with diethyl ether. The combined organic layers were dried (Na2SO4) and concentrated under reduced pressure. The crude product was purified by FC (1% MeOH/ CH2Cl2) to give 99 mg (41%) of 3 as a white foam: mp 81-82 °C; IR ν˜ 2931.6, 2110.4, 1693.5, 1462.2, 1427.6, 1263.4, 1112.6, 822.0, 739.2, 701.8; 1H NMR δ 9.10 (s, 1H), 7.59 (d, J ) 7.6, 1H), 7.46 (d, J ) 7.6, 1H), 7.35 (m, 20H), 6.37 (d, J ) 7.8, 1H), 5.26 (d, J ) 8.1, 1H), 4.37 (d, J ) 4.3, 1H), 3.71 (s, 1H), 3.50 (d, J ) 10.8, 1H), 3.40 (m, 1H), 2.72 (d, J ) 10.8, 1H), 1.06 (s, 9H), 0.89 (s, 9H); 13C NMR δ 161.73, 149.15, 138.11, 134.73, 134.69, 134.49, 134.16, 131.94, 131.49, 131.04, 130.74, 129.25, 129.23, 129.86, 127.03, 127.0, 126.99, 101.92, 85.54, 83.94, 73.46, 64.38, 62.30, 25.99, 25.91, 18.23, 18.08; MS (ESI, neg) 744.4 (M - 1, 100). Spectroscopy and Analysis Materials. IR Measurements. Tetrahydrofuran (Acros, 99.9%) was used without further purification. Eighteen MΩ-cm water was used to prepare all aqueous solutions. Chloroform (Aldrich, 99.9%) was purified by elution through a column of basic alumina. 15 N NMR Measurements. Tetrahydrofuran (Acros, 99.9%), deuterium oxide (Cambridge Isotope Laboratories, 99.9%), tetrahydrofuran-d8 (Cambridge Isotope Laboratories, 99.8%) and 15 N formamide (Cambridge Isotope Laboratories, 98%) were used without further purification. Eighteen MΩ-cm water was used to prepare all aqueous solutions. Deuterochloroform (Cambridge Isotope Laboratories, 99.8%) was purified by elution through a column of basic alumina. Measurements and Analysis. Equilibrium FTIR Measurements. Equilibrium FTIR absorbance spectra were recorded on a Bruker Vertex 70 FTIR spectrometer equipped with a globar source, KBr beamsplitter and a liquid nitrogen cooled mercury cadmium telluride (MCT) detector. The spectra were the result of 256 or 512 scans recorded at a resolution of 1.0 cm-1. The transmission measurements were recorded using a temperaturecontrolled cell consisting of calcium fluoride windows with a path length of ∼125 µm. The temperature of the IR cell was controlled by a water bath and the sample temperature was measured by a thermocouple embedded in the cell. The FTIR SCHEME 1

Gai et al. absorbance spectra were baseline corrected. The concentration of all azide-containing molecules was 50 mM unless otherwise noted, and the spectra were recorded at 293 K except for the variable temperature spectra in water of 1. The mixed solvent systems of THF and water were prepared by volume. Global Line Shape Fitting. Global line shape analysis was used to simultaneously model the azide IR absorbance band of compound 1 in multiple water-THF solvent mixtures. The azide IR absorbance band was modeled by three line shape functions. Each line shape function consisted of a linear combination of a Gaussian and Lorentzian function as shown in eq 1:36

F(ν˜ ) )

[

1

yo + A (1 - mLorentz)

(4 ln 2) /2 1/

e-(4 ln 2)(ν˜

- ν˜ o)2/fwhm2

π fwhm 2

mLorentz

fwhm 2 π 4(ν˜ - ν˜ )2 + fwhm2 o

]

+

(1)

where mLorentz is the fraction of the Lorentzian line shape, ν˜ o is the band position, fwhm is the full-width at half-maximum for the line shape, A is the area, and yo is a baseline offset. Two of the line shapes utilized in the global analysis of the azide absorbance band corresponded to 1 in pure water and THF. The line shape parameters for these two pure solvents were then utilized in the line shape analysis in water-THF solvent mixtures where a third line shape was utilized to model the three solvent environments encountered in these solvents.20,37-39 The line shape analysis was performed in Igor Pro (Wavemetrics). 15 N NMR Measurements. 15N NMR spectra were taken on a Varian INOVA 500 multinuclear Fourier transform NMR spectrometer (11.73 T). 15N NMR spectra were obtained at 50.5 MHz frequency, 7.6 µs pulse width, and a 45° pulse angle. The sweep width was 35476.7 Hz and 185942 points were taken for each spectrum. 15N spectra were referenced to a 100 mM solution of 15 N-labeled formamide in DMSO (HCONH2 ) 0.0 ppm). The reference solution was placed in a coaxial insert inside the 5 mm NMR tubes to allow simultaneous measurement of the 15N NMR spectrum of the reference and sample. The delay time between pulses was 1 or 10 s, and 400-10,000 scans were coadded. DFT Calculations Density Functional Theory Calculations. Geometry optimizations, single-point energy calculations, and vibrational analyses were carried out on model systems using the quantum chemical software package, Gaussian 03, on a multiprocessor Mac Pro computer.40 The calculations were performed using the density function theory (DFT) method, the B3PW91 density functional,41,42 and a 6-31++G(d,p) basis set.43,44 The calculations were performed in the gas phase, with or without one explicit water molecule to simulate hydrogen-bonding between the azide group and the solvent (water). The model structures were constructed and the normal modes of vibrations were

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CHART 1

visualized using the graphical user interface, GaussView 4. (2R,4R)-2-Methyl-4-azidotetrahydrofuran (9) was used as a model of compound 1 to probe the dependence of the azide asymmetrical stretch on H-bonding with water. Compound 1 was used as a model of compound 3 and compound 5 was used to model compound 4 in the heterodimer formation study.

Results and Discussion Synthetic Chemistry. Two synthetic routes were employed to prepare unlabeled N3-dU (1) for vibrational experiments (Scheme 1).45 These methods also allowed selective incorporation of the 15N isotopic label. Opening anhydrouridine (7) with

Figure 2. (A) FTIR absorbance spectra of 1 in water-THF mixtures ranging from 20 to 80% water (v/v) in approximately 10% increments recorded at 293 K. The spectra were normalized to the same maximum absorbance. The concentration of 1 was 50 mM. (B) The dependence of the IR absorbance band position of the azide stretch of 1 on the amount of THF in the water-THF solvent mixtures (open squares). Note: The band position for 0 and 100% water has been included for comparison, although the solvent is no longer a ternary mixture in these limiting percentages.

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Figure 3. (A) FTIR absorbance spectrum (open circles) of 1 in a 60% water solution modeled by three line shapes corresponding to a THF-like (TL, solid curve), water-THF (WT, short dashed curve), and water-like (WL, long dashed curve) band. (B) Dependence of the relative band area of the three components on percent water of the solvent. Note: The relative band area for 0 and 100% water has been included for comparison, although the solvent is no longer a ternary mixture in these limiting percentages.

the terminally 15N labeled sodium azide provided a 1:1 isotopomeric mixture of N3-dU containing the 15N isotope at the internal position (1i) and the terminal position (1t) of the azido group (Chart 1). It was necessary to acylate the crude product for purification and then deprotect it using ammonia. The threestep route provided 1i/1t in 58% yield. A diazotransfer reaction25-27 of 2′-amino-2′-deoxyuridine (6) and an isotopomeric mixture of internal and terminal 15N labeled triflic azides resulted in an 81% yield of a 1:1 mixture of unlabeled 1 and middle labeled 1m, respectively. The diazotransfer reaction was the superior route because of fewer steps, higher yields, and easier purification. The diazotransfer method also has the potential for incorporation of azido groups into oligonucleotides (vide supra). Infrared Spectroscopy. The solvent dependence of the azide IR asymmetric stretching absorbance band (ν˜ N3) of N3-dU was investigated in THF, water and mixtures of these two solvents to explore the sensitivity of this IR probe to microenvironments present in nucleic acid structures. The azide stretching frequency in THF was 2110.7 cm-1 and blue-shifted by 13.5 cm-1 to 2124.2 cm-1 in water. The bandwidth (fwhm) was 21 cm-1 in

THF and 22 cm-1 in water. The frequency shift of ν˜ N3 was over 45% larger than the shift of ν˜ CN observed with 2a in going from THF to water, showing that the azide probe is more sensitive to solvent variation than the nitrile probe.20 In contrast, the Stark tuning rate of 1 was previously measured to be less than half of 2a, showing that the azide is less sensitive to the local electric field.2 The frequency shift of ν˜ N3 is presumably due to solute-solvent interactions including hydrogen bonding and solvent dynamics.20,36 To further explore the effect of hydrogen bonding on the azide asymmetric stretching frequency, DFT calculations were conducted on a model system. In order to focus on the hydrogen bond between the water and azido group, a model compound without the base and alcohol groups was used. The ν˜ N3 of the model compound, (2R,4R)-2-methyl-4azidotetrahydrofuran (9), was calculated in the gas phase with and without an explicit water molecule to interact with the azido group. Prior to the frequency calculation, the geometry of 9 and the one water molecule was optimized. The calculations show that interaction with the single water causes a blue shift of 8.1 cm-1 of the azide asymmetric stretching frequency due to a hydrogen bond with the terminal nitrogen atom of the azide.

Sensitive Multispectroscopic Probe for Nucleic Acids This result suggests that at least part of the experimental blue shift is due to hydrogen bonding of the water to the azide. These results are consistent with the solvent shifts observed with azidemodified amino acids.3,5 A series of THF-water mixtures provided a more complete analysis of the solvent dependence of ν˜ N3 of N3-dU. Figure 2A shows the ν˜ N3 IR absorbance band of N3-dU for solvents ranging from 20 to 80% water in THF. These spectra show that the azide band shifted from 2113.4 cm-1 in 20% water to 2122.5 cm-1 in 80% water. Figure 2B shows that the shift in the band position due to the variation of water in the solvent system was nonlinear with the largest shifts occurring at high water percentages. These results are consistent with literature results on the solvent dependence of ν˜ N3 of N3-A.3 The bandwidth changes were also nonlinear with the broadest band (26 cm-1) observed at 70% water.45 These results highlight the sensitivity of the azide IR absorbance band to H-bonding interactions with water. Three line shape functions, each composed of a linear combination of a Lorentzian and Gaussian function given in eq 1, were used to globally fit the azide IR absorbance band in the THF-water mixtures. The result of the global fit for the 60% water absorbance spectrum is shown in Figure 3A. Drawing from previous work,20,39 the three bands were labeled THF-like (TL), water-THF (WT), and water-like (WL), as shown in Figure 3B. The WL and TL bands were predominately Lorentzian, while the WT was predominately Gaussian. The WL and TL bands arise exclusively from interactions of water and THF with N3-dU, respectively, and the WT band is due to solute-solvent interactions involving both water and THF. The relative band area percentage of the TL band from the global line shape analysis decreases as the percentage of water in the solvent increases, while the relative band area of the WL band increases over the same range (Figure 3B). The relative area of the WT band gradually increases with increasing water percentage to a maximum between 60 and 70% water and then rapidly decreases with further increases in water content. The exact nature of the solvent-solute interactions in the WT band is unclear; however, the experimental results clearly highlight the sensitivity of the azide IR absorbance band (frequency and bandwidth) to its local environment. The dependence of ν˜ N3 on the formation of a heterodimer between 3 and 4 (Chart 1) in chloroform solution was investigated to mimic base-pair formation in nucleic acid structures. This triply hydrogen-bonded heterodimer was selected due to the relative insensitivity of the binding constant in chloroform to minor structural perturbations, remote from the hydrogen-bonding region.20,46-49 The ν˜ N3 was found to be essentially independent of “base pair” formation, showing a 0.3 cm-1 red shift. This is in contrast to the ν˜ CN of 2c which underwent a 2.6 cm-1 blue shift upon heterodimer formation with 4.20 The insensitivity of ν˜ N3 is not surprising since the azide is attached to the sugar and is remote from the H-bonding interactions involved in the heterodimer. This result suggests that the azido moiety will be selectively sensitive to changes in the phosphate sugar region. This selective sensitivity will greatly aid in the analysis of future studies of nucleic acids utilizing N3-dU since the measured frequency and bandwidth changes can be attributed essentially only to changes in the phosphate sugar region. Therefore, N3-dU and nitrile 2 are complementary in primarily sensing either the phosphate sugar region or major groove of nucleic acids, respectively. DFT calculations on a model heterodimer between N3-dU and 5 was consistent with the experimental results predicting a 0.35 cm-1 red shift in the

J. Phys. Chem. B, Vol. 114, No. 23, 2010 7963 TABLE 1: Experimental and DFT Calculated Isotopic Shift of the Azide Asymmetric Stretching Frequency in N3-dU ∆ν˜ N3 (cm-1)a compound

exptl results (cm-1)

exptl

DFT

N3-dU (1) 15 NNN-dU (1t) N15NN-dU (1m) NN15N-dU (1i)

2110.7 2089.0 2068.6 2111.3

21.7 42.1 -0.6

25.2 48.2 -3.0

a

(Unlabeled azide frequency) - (15N labeled azide frequency).

ν˜ N3 upon heterodimer formation. The DFT calculations also showed that the azide probe destabilizes the heterodimer by only ∼1% relative to the heterodimer between 2′-deoxyuridine and 5.45 The intrinsic temperature dependence of the ν˜ N3 of N3-dU was measured by heating a H2O solution from 20 to 70 °C, which represents a typical temperature range needed to melt nucleic acid structures. The temperature increase resulted in a red shift of 2.1 cm-1 in the azide asymmetric stretch. This shift likely arises due to changes in the structure of the aqueous solvent around the azido group that affects the geometry and strength of H-bonding interactions between H2O and the azido moiety. This intrinsic temperature dependence is significantly less than the solvent dependence of the azide asymmetric stretch. This feature suggests that the azide IR absorbance band can be used to probe changes in the phosphate sugar region of nucleic acids modulated by temperature without significant interference from the temperature dependence of the IR absorbance band. Infrared spectroscopy was also used to investigate the isotopic shifts for each of the isotopomers relative to unlabeled 1.45 Table 1 shows that substitution of the 15N isotope for the middle nitrogen atom (1m) yields a large 42 cm-1 isotopic red shift, whereas substitution of an internal nitrogen atom with a 15N label (1i) results in almost no shift. The isotopic shift for 1t was nearly equidistant between these extremes. These shifts were similar to those predicted by DFT calculations of these isotopically labeled variants of N3-dU (Table 1). These isotopic variants can be used for 15N NMR measurements, as described below, and to minimize solvent interference from the water combination band at 2134 cm-1.50 15 N NMR Spectroscopy. As described above, 15N isotopic labels were incorporated into each of the different positions of the azido group to give isotopomers 1i, 1m, and 1t. The assignment of the 15N isotopomers was initially based upon knowledge of the mechanism of the anhydrouridine opening reaction23 (which produced 1i and 1t) and the copper catalyzed diazotransfer reaction26 (which produced 1m and unlabeled 1). Supporting evidence came from experimental and DFT calculated isotopic IR shifts (Table 1). The 15N NMR experimental TABLE 2: Solvent Dependence of 15N NMR Chemical Shifts of N3-dU δ15Na compound

D 2O

THF-d8

∆δ15Nb

DFT gas phase calcn for 15Nd15Nd15N-dUc

NNN-dU (1t) N15NN-dU (1m) NN15N-dU (1i)

99.4 131.0 -43.9

95.6 131.9 -45.2

3.8 -0.9 1.3

104.9 125.9 -45.6

15

a Chemical shifts referenced to 15N formamide (HCONH2 ) 0.0 ppm). b δ15N(D2O) - δ15N(THF-d8). c DFT gas phase chemical shifts were calculated using ammonia as a reference (NH3 ) 0.0 ppm) and converted to the formamide chemical shift scale (HCONH2 ) 0.0 ppm) by subtracting 112 ppm. 56

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Figure 4. (A) 15N NMR chemical shift spectra of 1t in either pure THF-d8 or pure D2O. (B) 15N NMR chemical shift spectra of 1m in either pure THF-d8 or pure D2O. (C) 15N NMR chemical shift spectra of 1i in either pure THF-d8 or pure D2O. All the spectra were normalized to the same maximum absorbance. Note that the scale of the x-axis is different in the three spectra, but all three show a 5.0 ppm spectral window. Panels A and C are different regions of the same spectrum, i.e., a 1:1 mixture of 1t and 1i.

chemical shifts closely matched DFT chemical shift calculations (Table 2) and were consistent with observed chemical shifts in previous literature studies.51

The 15N chemical shift demonstrated different sensitivity to the change of the local solvent environment depending upon the position of the 15N atom within the azido probe. The

Sensitive Multispectroscopic Probe for Nucleic Acids chemical shift of 1t was found to shift most when changing the solvent from THF-d8 to D2O. The chemical shift was measured to be 95.59 ppm in THF-d8 and 99.39 ppm in D2O (Figure 4). The 3.8 ppm ∆δ15N was presumably due to the hydrogen bonding between the terminal nitrogen atom in the azido group and D2O. This interaction is stronger than the hydrogen bond between D2O and the middle (1m) or internal (1i) nitrogen atoms of the azido group, which showed ∆δ15N of -0.9 ppm and 1.3 ppm in going from THF-d8 to D2O, respectively. The reason why the middle nitrogen atom shifts in the opposite direction relative to the other two is unclear, although recent ab initio calculations done on azidomethane show that this nitrogen atom does not form H-bonds with water whereas the internal and terminal nitrogen atoms do.4 All of these solvent-induced ∆δ15N values are considerably smaller than the 15N ∆δ15N of -8.45 ppm observed for 2b in going from THF-d8 to D2O.21 Thus, the nitrile group is significantly more sensitive than the azido group in terms of solvent-induced 15N NMR chemical shifts but significantly less sensitive than the azido group in terms of solventinduced IR frequency shifts. Although the signal-to-noise ratio of these direct detect 1D spectra of monomers is excellent (Figure 4), it may be more difficult to detect the 15N nuclei in oligomers because of longer relaxation times and lower concentrations. In this context, indirect detect 2D NMR methods such as HMBC may be preferred. This technique was successfully used to obtain natural abundance 15N NMR spectra of both internal and middle nuclei of an azido group,51 which suggests these methods will be applicable to 1i or 1m incorporated into oligonucleotides. Similar to the IR results, the 15N chemical shift of the azide probe in 3m was found to remain essentially unchanged (∆δ15N ) 0.05 ppm) upon heterodimer formation with 4 in deuterochloroform. DFT calculations predicted a larger value, 0.47 ppm, for ∆δ15N. DFT calculations for ∆δ15N of 1i and 1t showed similar values upon heterodimer formation.45 A small intrinsic temperature dependence of the chemical shift of N3dU in D2O solution was observed, with the terminal 15N nucleus shifting 0.53 ppm downfield when increasing the temperature from 20 to 70 °C. Conclusions The azido group of N3-dU was shown to be an effective IR and 15N NMR spectroscopic probe and is a promising relatively nonintrusive, site-specific, multispectroscopic probe of the phosphate sugar region of nucleic acids. In terms of solvent/ dielectric response, the azido group of N3-dU was more sensitive than a previously studied20,21 nitrile modified nucleoside (2) when IR frequency shifts are compared, but less sensitive when 15 N NMR chemical shifts are compared. In terms of formation of a base-pair mimic, the IR and 15N NMR spectroscopic peak shifts of N3-dU were minimal, as expected, given the azide’s location on the sugar of the nucleoside. The terminal nitrogen atom of the azido group showed the greatest sensitivity to solvent effects as measured by 15N NMR peak shifts, and the middle nitrogen atom was the least sensitive. However, the vibrational isotopic shift induced by the 15N substitution of the middle nitrogen atom was significantly greater than that observed for the other azide positions. Recently it has been shown that a diazotransfer reaction followed by an azide-alkyne click cycloaddition is an effective one-pot procedure to functionalize amines.52,53 Since monomeric N3-dU has previously been shown to undergo click chemistry,14,47 incorporation of this probe into oligomers will also provide a selective chemical handle for incorporation of fluorescent dyes

J. Phys. Chem. B, Vol. 114, No. 23, 2010 7965 or nitroxide spin labels. The incorporation of N3-dU into DNA oligomers via a diazotransfer reaction is currently being investigated, and spectroscopic analysis of azido-DNA will be reported in due course. The extension of the 1D IR and 1D 15N NMR measurements to 2D IR9,54,55 and 2D NMR51 measurements is also being explored. Acknowledgment. We thank Beth Buckwalter for conducting the NMR experiments and Austin Luskin for research on the diazotransfer reaction. We are grateful to Carol Strausser for assistance in the editing of the manuscript and Lisa Mertzman for obtaining materials and supplies. This work was supported by an F&M Snavely Summer Research stipend and a Snavely Research Award to X.S.G., a Mellon/CPC New Tasks, New Tools grant to E.E.F., and an award from Research Corporation to S.H.B. (CCSA 7352). Supporting Information Available: Full synthetic schemes; calculated DFT structures, potential energy surfaces, and ∆δ15N for heterodimer formation; IR spectra of 1m/1 and 1i/1t; solvent dependence of fwhm. This material is available free of charge via the Internet at http://pubs.acs.org. References and Notes (1) Suydam, I. T.; Boxer, S. G. Biochemistry 2003, 42, 12050–12055. (2) Silverman, L. N.; Pitzer, M. E.; Ankomah, P. O.; Boxer, S. G.; Fenlon, E. E. J. Phys. Chem. B 2007, 111, 11611–11613. (3) Oh, K. I.; Lee, J. H.; Joo, C.; Han, H.; Cho, M. J. Phys. Chem. B 2008, 112, 10352–10357. (4) Choi, J. H.; Oh, K. I.; Cho, M. H. J. Chem. Phys. 2008, 129, 174512. (5) Ye, S. X.; Huber, T.; Vogel, R.; Sakmar, T. P. Nat. Chem. Biol. 2009, 5, 397–399. (6) Gramlich, P. M. E.; Wirges, C. T.; Manetto, A.; Carell, T. Angew. Chem., Int. Ed. 2008, 47, 8350–8358. (7) Kolb, H. C.; Finn, M. G.; Sharpless, K. B. Angew. Chem., Int. Ed. 2001, 40, 2004–2021. (8) Lindquist, B. A.; Furse, K. E.; Corcelli, S. A. Phys. Chem. Chem. Phys. 2009, 11, 8119–8132. (9) Bandaria, J. N.; Dutta, S.; Hill, S. E.; Kohen, A.; Cheatum, C. M. J. Am. Chem. Soc. 2008, 130, 22–23. (10) Lim, M. H.; Hamm, P.; Hochstrasser, R. M. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 15315–15320. (11) Lu, S.; Sazinsky, M. H.; Whittaker, J. W.; Lippard, S. J.; MoenneLoccoz, P. J. Am. Chem. Soc. 2005, 127, 4148–4149. (12) Cooper, I. B.; Barry, B. A. Biophys. J. 2008, 95, 5843–5850. (13) Polushin, N. N.; Smirnov, I. P.; Verentchikov, A. N.; Coull, J. M. Tetrahedron Lett. 1996, 37, 3227–3230. (14) Jawalekar, A. M.; Meeuwenoord, N.; Cremers, J. G. O.; Overkleeft, H. S.; van der Marel, G. A.; Rutjes, F. P. J. T.; van Delft, F. L. J. Org. Chem. 2008, 73, 287–290. (15) Evans, R. K.; Haley, B. E. Biochemistry 1987, 26, 269–276. (16) Seo, T. S.; Li, Z. M.; Ruparel, H.; Ju, J. Y. J. Org. Chem. 2003, 68, 609–612. (17) Catalano, C. E.; Allen, D. J.; Benkovic, S. J. Biochemistry 1990, 29, 3612–3621. (18) Geselowitz, D. A.; Neumann, R. D. Bioconjugate Chem. 1995, 6, 502–506. (19) Krummel, A. T.; Zanni, M. T. J. Phys. Chem. B 2008, 112, 1336– 1338. (20) Watson, M. D.; Gai, X. S.; Gillies, A. T.; Brewer, S. H.; Fenlon, E. E. J. Phys. Chem. B 2008, 112, 13188–13192. (21) Gillies, A. T.; Gai, X. S.; Fenlon, E. E.; Brewer, S. H. Unpublished results. (22) Azuma, A.; Nakajima, Y.; Nishizono, N.; Minakawa, N.; Suzuki, M.; Hanaoka, K.; Kobayashi, T.; Tanaka, M.; Sasaki, T.; Matsuda, A. J. Med. Chem. 1993, 36, 4183–4189. (23) Verheyden, J. P. H.; Wagner, D.; Moffatt, J. G. J. Org. Chem. 1971, 36, 250. (24) ChemGenes Corporation, Wilmington, MA. (25) Zaloom, J.; Roberts, D. C. J. Org. Chem. 1981, 46, 5173–5176. (26) Nyffeler, P. T.; Liang, C. H.; Koeller, K. M.; Wong, C. H. J. Am. Chem. Soc. 2002, 124, 10773–10778. (27) Lundquist, J. T.; Pelletier, J. C. Org. Lett. 2001, 3, 781–783.

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