Nucleic Acid Complexing Glycosyl Nucleoside-Based Amphiphile

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Bioconjugate Chem. 2005, 16, 864−872

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Nucleic Acid Complexing Glycosyl Nucleoside-Based Amphiphile Jerome Arigon,† Carla A. H. Prata,§ Mark W. Grinstaff,§ and Philippe Barthe´le´my*,† Faculte´ des sciences d’Avignon, 33 rue Louis Pasteur, F-84000 Avignon, France, and Departments of Chemistry and Biomedical Engineering, Metcalf Center for Science and Engineering, Boston University, Boston, Massachusetts 02215. Received February 4, 2005; Revised Manuscript Received April 25, 2005

A neutral amphiphile derived from uridine featuring two oleyl chains and one glucose for DNA binding was prepared using a convenient four-step synthetic route. The nucleic acid binding capabilities of this amphiphile were investigated by UV-vis, quasi-elastic light scattering (QELS), transmission electronic microscopy (TEM), gel electrophoresis, 31P NMR, IR, and circular dichroism (CD). Amphiphile-nucleic acid complex formation is a consequence of the amphiphilic character of the molecule, phosphate-sugar, and nucleobase-nucleobase interactions. This work presents for the first time a glyco-nucleo-amphiphile capable of binding efficiently the nucleic acid double helix structure.

INTRODUCTION

The supramolecular assemblies observed between nucleic acids and different amphiphiles represent elegant examples of the role multiple noncovalent interactions have in governing structure and even function. These assemblies are of fundamental interest, as well as clinical importance, since some of these assemblies are used for the in vitro and in vivo delivery of DNA. The delivery of therapeutic nucleic acid offers the potential to treat diseases by replacing or inhibiting defective genes. The vast majority of DNA supramolecular assemblies are formed with molecular or macromolecular cationic amphiphiles including lipids, linear polymers, and dendritic polymers (1-6). One caveat with these cationic amphiphiles is that these systems are likely to interact with other biologics, since most proteins are also negatively charged at neutral pH. However, there are a few examples of DNA supramolecular assemblies formed with noncationic amphiphiles (7). Recently, neutral sugar- and poly(ethylene glycol)/uridine-based amphiphiles have been reported that complex DNA (8-10). The neutral glycosyl amphiphiles are particularly interesting since these compounds may provide a means to develop amphiphiles for selective biologic complexation. Such amphiphiles could be of interest for medical and biotechnological applications involving the manipulation or use of DNA or proteins. With regards to the glycosyl amphiphiles, there are known examples of natural saccharides that bind nucleic acid. For example, S. Shinkai et al. have reported the molecular recognition between adenine, cytosine, and uracil in a single-stranded RNA and schizophyllan, a natural polysaccharide (12). It is known that when a given polyol structure, like a saccharide, is added to an aqueous solution containing DNA, the stability of the double helix can be affected. The effects observed depend on the polyol influence on the forces that stabilize the double helix, namely, hydrogen bonding, van der Waals forces, and electrostatic interactions. Consequently, the polyol can either stabilize or destabilize the duplex as * To whom correspondence should be addressed. E-mail: [email protected]. † Faculte ´ des sciences d’Avignon. § Boston University.

Figure 1. Chemical structures of 1-(2′,3′-dioleylcarbamoyluridine-5′)-β-D-glucopyranoside (DOUGluc).

evident by a change in the DNA denaturation temperature (Tm). G. Barone and co-workers have studied the effects of different small molecule polyols (i.e., glycerol, arabitol, D-mannitol, D-sorbitol, myo-inositol, etc.) on the thermal denaturation of DNA (13). The double helix destabilization was mainly attributed to the polyols interacting with the polynucleotide solvation sites. The polyols displace water molecules and alter the electrostatic interactions between the polynucleotide and its surrounding counterions. Consequently, when the concentration of polyols increases, the Coulombian repulsion between the negative charges of the phosphate groups increases leading to greater destabilization of the DNA double helix. Stabilizing contributions (polyols are less efficient than water in competing with the base-base interstrand hydrogen bonds) are counterbalanced by destabilizing electrostatic repulsions. A stabilization of the DNA double helix has been observed in the presence of a maltoheptaose amphiphile (14), where the saccharide promotes stabilization of the double helix versus single strand by the formation of DNA aggregates. This is likely due to the amphiphilic character of the polyols. To further explore the interactions between DNA and a neutral amphiphile, we have designed a noncationic amphiphile possessing both nucleoside and glycosyl moieties (Figure 1). Herein, we report the synthesis and physicochemical properties of this amphiphile as determined by UV-vis, quasi-elastic light scattering (QELS), NMR, IR, circular dichroism (CD), gel electrophoresis, and transmission electronic microscopy (TEM).

10.1021/bc050029y CCC: $30.25 © 2005 American Chemical Society Published on Web 06/18/2005

Nucleic Acid Complexing Glyco-nucleo-amphiphile MATERIALS AND METHODS

General Experiments and Analytical Conditions. Unless noted otherwise, all starting materials were obtained from commercial suppliers and were used without further purification; the solvents were redistilled on calcium chloride, calcium hydride, potassium hydroxide, or sodium according to the solvent used. All compounds were characterized using standard analytical and spectroscopic data such as 1H, 13C NMR (apparatus BRUKER Avance DPX-300, 1H at 300.13 MHz, 13C at 75.46 MHz, and 31P at 121.49 MHz) and mass spectrometry (instrument JEOL SX 102, NBA matrix). The NMR chemical shifts are reported in ppm relative to tetramethylsilane using the deuterium signal of the solvent (CDCl3, MeOD) as a heteronuclear reference for 1H and 13C. The 1H NMR coupling constants, J, are reported in hertz. TEM microscopy experiments were performed on a Philips CM 10 (negative staining with ammonium molybdate 1% in water, Cu/Pd carbon-coated grids). Fluorescence spectra were recorded on a Hitachi F2500 using λex ) 320 nm and λem ) 350-450 nm. The hydrodynamic radii (Rh) were determined at 25 °C using a Wyatt Mini-Dawn quasi-elastic light scattering (QELS) instrument. UV-visible studies were performed on a Perkin-Elmer Lambda 25 Peletier equipped, and circular dichroism was performed on a JASCO J-810 spectropolarimeter. The 31P NMR experiments were performed on a BRUKER AC 250 at 121.49 MHz. The 31P NMR chemical shifts (δP) are reported in ppm relative to phosphoric acid. FTIR spectra were recorded using a NICOLET Fourier transform spectrophotometer. Silica gel 60 (particle size 40-60 µm) was employed for flash chromatography. Thin layer chromatograms were performed with aluminum plates coated with silica gel 60 F254 (Merck). Varian silica gel reverse phase C18 MEGA BE-C18, 2GM, 12 mL was used. Abbreviation. DOUGluc, 1-(2′,3′-dioleylcarbamoyluridine-5′)-β-D-glucopyranoside; HEPES, N-(2-hydroxyethyl)piperazine-N′-(2-ethanesulfonic acid); M.S., molecular sieves; EtBr, ethidium bromide; DMAP, (dimethylamino)pyridine; DCM, dichloromethane; DMF, dimethylformamide; AcOEt, ethyl acetate; SUV, small unilamellar vesicles; polyA-polyU, polyadenylic acid-polyuridylic acid; polyC-polyG, polycytidylic acid-polyguanylic acid; Tm, melting temperature; RT, room temperature; CDI, carbonyl diimidazole. Materials. Calf thymus DNA (CAS no. 91080-16-9), pBR322 plasmid DNA (CAS no. 93384-17-9), polyadenylic acid-polyuridylic acid (CAS no. 24936-38-7], and polycytidylic acid-polyguanylic acid (CAS no. 90385-88-9) were obtained from Sigma. Buffer (0.02 M HEPES, 0.01 M NaCl) was prepared with distilled and filtered (0.22 µm) water, and the pH was adjusted to 7.4 with NaOH 0.1 M. Synthetic Procedures. 2′,3′-Dioleylcarbamoyluridine (2). 5′-Dimethoxytrityluridine 1 (1 g, 1 equiv), carbonyldiimidazole (1.22 g, 2.2 equiv), and a catalytic amount of DMAP were dissolved in 20 mL of anhydrous DMF. After 1 h at room temperature, oleylamine (4 mL, 7 equiv) was added, and then the reaction mixture was stirred for 12 h. DMF was removed under reduced pressure, and 20 mL of DCM was added. To this solution an excess of a 3% trichloroacetic acid solution in DCM was added, and the reaction mixture was stirred for 30 min at room temperature. After addition of methanol (3 mL), the organic layer was washed three times with 10 mL of water and dried over sodium sulfate. Compound 2 (0.54 g) was obtained after chromatography (DCM/MeOH 95/

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5). Yield: 36%. Rf: 0.32 (DCM/MeOH 95/5). 1H NMR (CDCl3) δ in ppm: 0.96 (s, CH3), 1.2 (m, CH2), 2.3 (m, CH2), 3.2 (m, H5′), 3.92 (m, H4′), 4.2 (m, H3′), 4.95 (m, NH), 5.4 (m, H2′), 5.8 (d, CH, J ) 8.1 Hz), 6.1 (d, H1′, J ) 3.3 Hz), 7.85 (d, CH, J ) 8.0 Hz), 8.6 (s, NH). 13C NMR (CDCl3) δ in ppm: 14 (s, CH3), 27 (s, CH2β), 30 (m, CH2), 33 (m, CH2R), 42 (s, C-NH), 70 (m, C2,C3), 100 (s, CH), 131 (m, CH dCH), 133 (m, CH), 164 (m, CO carbamates), 170 (m, CO). MS FAB+: [M + Na], m/z ) 853. 1-(2′,3′-Dioleylcarbamoyluridine-5′)-2,3,4,6-tetra-O-acetylβ-D-glucopyranoside (3). 2,3,4,6-Tetra-O-acetyl-R-D-glucopyranosyl trichloroacetimidate (0.2 g, 1.5 equiv) (1518), 2′,3′-dioleylcarbamoyluridine 2 (0.2 g, 1 equiv), molecular sieves, and BF3‚Et2O (0.18 g, 1.5 equiv) were dissolved in 10 mL of DCM. The reaction mixture was stirred for 4 h at room temperature. The product was obtained as a white solid (0.15 g) after chromatography (DCM/MeOH 95/5). Yield: 55%. Rf ) 0.72 (DCM/MeOH 95/5). 1H NMR (CDCl3) δ in ppm: 0.96 (s, CH3), 1.2 (m, CH2), 2.0 (m, CH3 acetyl), 3.1 (m, CH2R), 3.9 (m, H5′), 4.0 (m, H4′), 4.1 (m, H5), 4.2 (m, H6, H6), 4.7 (m, H3′), 4.85 (m, H2), 5.1 (m, H3), 5.5 (m, H4), 5.6 (d, H1β, J ) 8.2 Hz), 5.65 (d, CH, J ) 8.2 Hz), 6.1 (d, H1′, J ) 3.0 Hz), 7.9 (d, CH, J ) 8.1 Hz). 13C NMR (CDCl3) δ in ppm: 14 (s, CH3), 20 (s, OAc), 27 (s, CH2β), 30 (m, CH2), 33 (m, CH2R), 42 (s, CH2NH), 61-75 (m, C5,C3,C2,C3′,C4′,C5′), 97 (s, C1), 105 (s, CH), 131 (m, CHdCH), 133 (s, CH), 164 (m, CO carbamates), 170 (m, CO). MS FAB+: [M + H], m/z ) 1162. 1-(2′,3′-Dioleylcarbamoyluridine-5′)-β-D-glucopyranoside (4). 1-(2′,3′-Dioleylcarbamoyluridine-5′)-2,3,4,6-tetraO-acetyl-β-D-glucopyranoside (3, 0.15 g) and a catalytic amount of sodium methoxide were dissolved in 10 mL of methanol. The reaction mixture was stirred for 4 h at room temperature. After evaporation of methanol under reduced pressure, the crude was purified by silica gel reverse phase column (DCM/MeOH 3/7) to afford the compound as a white solid (0.12 g). Yield: quantitative. Rf ) 0.52 (DCM/MeOH reverse phase). 1H NMR (CD3OD) δ in ppm: 0.96 (s, CH3), 1.3 (m, CH2), 1.5 (m, CH2β), 3.1 (m, CH2R), 3.3 (m, H6, H6), 3.7 (m, H3′, H4′), 3.8 (m, H5′, H2′), 4.3 (m, H5), 4.6 (s, H4), 4.8 (s, H3), 5.3 (m, CHdCH), 5.7 (d, H′β, J ) 8.2 Hz), 5.8 (d, CH, J ) 8.0 Hz), 6.2 (d, H1′, J ) 3.0 Hz), 8.0 (d, CH, J ) 8.1 Hz). 13C NMR (CD3OD) δ in ppm: 15 (s, CH3), 27 (s, CH2β), 30 (m, CH2), 33 (m, CH2R), 42 (s, CH2NH), 61-75 (m, C5,C3,C2,C3′,C4′,C5′), 97 (s, C1), 105 (s, CH), 131 (m, CHd CH), 133 (s, CH), 162 (m, CO carbamates), 170 (m, CO). MS FAB+: [M + Na], m/z ) 1015. HRMS [M + Na] m/ztheoritical ) 1015.6551, m/zobserved ) 1015.6559. [R]20 D ) -7.26 (CH3OH). Critical Aggregation Concentration. For the measurements of critical micellar concentration, a stock solution of pyrene (0.5 mM) in ethanol was added to test tubes. The ethanol was evaporated by a stream of nitrogen and thereafter under vacuum for 1 h. Varying amounts of amphiphile stock solution in buffer were added to the tubes to give a final volume of 1 mL and a final pyrene concentration of 0.5 µM. The samples were incubated in the dark at 25 °C for 24 h with intermittent vigorous mixing. The fluorescence intensity (I1, λ ) 373 nm) was plotted as a function of the lipid concentration. Thermal Denaturation and Circular Dichroism Studies. Samples were prepared in the following buffers: (i) 0.02 M HEPES, 0.01 M NaCl, pH ) 7.4, and (ii) 0.02 M HEPES, 0.01 M NaCl, 0.01 M MgCl2, pH ) 7.4. Solutions at different ratios of polyA-polyU (1 mg/mL) or CT-DNA (1 mg/mL) to 4 were prepared in either sodium (i) or magnesium (ii) buffer and stirred for 10 min.

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Scheme 1. Synthetic Route to Glucosyl Dicarbamate Uridine Amphiphilea

a Conditions: (a) CDI/DMAP in DMF 2 h, then oleylamine 12 h; (b) trichloroacetic acid in CH Cl (yield 2 2 a+b ) 36%); (c) 2,3,4,6tetra-O-acetyl-R-D-glucopyranosyl trichloroacetimidate, M.S., and BF3-Et2O in CH2Cl2 (yield ) 55%); (d) MeONa in MeOH (yield ) quantitative).

The temperature was increased from 15 to 95 °C at 1 °C/min, and the absorbance at 260 nm was measured every 1 °C increments. Hyperchromicity ) A - A0, where A is the absorbance measured at λ ) 260 nm and A0 is the initial absorbance (λ ) 260 nm) at 20 °C. Tm values were determined from first-order derivatives. CD spectra were recorded at 25 °C in rectangular quartz cuvettes of 0.1 cm; each spectrum was averaged from three successive accumulations and was baseline-corrected and smoothed using the software supplied by JASCO. Quasi-Elastic Light Scattering (QELS). The Rh was measured at a 90° angle and λ ) 685 nm using a flow cell. The solvent was filtered through a 0.02 µm membrane. The lipid and DNA solutions of desired concentrations were prepared and filtered through 0.2 µm pore size filter (Whatman) into the scattering cell. Rh measurements were realized at t ) 0, 24, and 36 h of 4 (50 equiv per DNA phosphate) in the absence or presence of CTDNA (1 mg/mL). Rh reported corresponds to the average of three experiments in aqueous HEPES buffer. Astra software (version 4.90.80) was used for data acquisition and analysis. 31 P NMR Studies and Job Plot. Na2HPO4 (3 mM) solution in D2O-DMSO-d6 (8:2) was added with different amounts of 4, 2, or D-glucose, and the mixtures were vortexed during 30 s. The chemical shifts (δP) of Na2HPO4 were measured by 31P NMR. For the Job plot, the concentration of the mixture was kept constant at 3 mM. Infrared Spectroscopy. PolyA-polyU (0.5 mg/mL) was mixed with DOUGluc 4 (1.5 mg) in H2O and vortexed for 30 s; then the solution was freeze-dried. The solid obtained was mixed with KBr, and 64 scans were accumulated. In the case of D2O experiments, the polyApolyU/DOUGluc 4 and polyC-polyG/DOUGluc 4 mixtures are prepared in D2O in the same conditions as for solid FTIR experiments, and the IR spectra are obtained directly from the D2O solutions. Agarose Gel Electrophoresis. Plasmid DNA (2 µL, 0.02 g/L in HEPES buffer) was added to various amounts of 4 at RT, and the mixture was vortexed during 30 s and left at RT for 36 h. Then the solutions were loaded into 1% (w/v) agarose gel at 60 V/cm in HEPES buffer. DNA was revealed with EtBr and visualized under UV light.

Transmission Electronic Microscopy (TEM). The samples were prepared with a solution of calf thymus DNA (1 mg/mL in HEPES buffer) and extruded (50 nm filter) HEPES buffer solution containing 50 equiv of DOUGluc 4 (for 1 equiv of phosphate DNA). This solution was vortexed and left for 36 h at RT prior to examination. RESULTS AND DISCUSSION

The formation of amphiphile-DNA supramolecular assemblies is well-known in the case of cationic lipids; however, only a few examples of neutral amphiphiles that complex with DNA have been reported (7, 10, 14, 19). To further increase the known types of DNA-binding neutral amphiphiles, we are designing amphiphiles that are based on a nucleoside and a sugar moiety. In this approach, the nucleoside and sugar derivative can provide sufficient interaction forces with DNA to form nucleic acid-amphiphile supramolecular assemblies. Consequently, we have synthesized the 1-(2′,3′-dioleylcarbamoyluridine-5′)-β-D-glucopyranoside and evaluated the ability of this neutral amphiphile to bind to nucleic acids using UV-vis, QELS, TEM, gel electrophoresis, 31P NMR, FTIR, and CD. Synthesis. The synthetic strategy is shown in Scheme 1 and involves four steps starting from 5′-dimethoxytrityluridine. The lipophilic intermediate 2 was obtained by coupling oleylamine to the 2′ and 3′ ribose secondary alcohols using CDI to afford carbamate linkages. The dimethoxytrityl protecting group was removed using trichloroacetic acid. Finally, the amphiphilic dioleyl nucleoside derivative 4 was obtained after addition of the glucose moiety via a trichloroacetimidate glycosylation and a deprotection step to remove the acetyl groups. Amphiphile 4 is soluble in water, whereas 2 possesses limited aqueous solubility and is soluble in DMSO. Physicochemical Studies. The critical aggregation concentration (CAC) of 4 was determined using a fluorescence titration method (20, 21). Thus, to determine the CAC value of our glycosyl nucleoside amphiphile, a stock solution of 4 was added to a pyrene-saturated solution (4 × 10-7 mol/L) at 298.2 K and stirred for 24 h to reach complete equilibrium between amphiphile and pyrene. The fluorescence emission was monitored at 394

Nucleic Acid Complexing Glyco-nucleo-amphiphile

Figure 2. Measurements of pyrene fluorescence at 394 nm (λext ) 320 nm) versus log(c) in mol/L of compound 4. CAC obtained was determined to be 1 × 10-5 mol/L.

nm (λext was 320 nm). The CAC value for amphiphile 4 is 10-5 mol/L (see Figure 2). Note, that this value is similar in magnitude to CACs reported in the literature for mono- and dioleyl amphiphilic polyols or cationic derivatives of 4 suggesting that the nucleoside moiety does not significantly affect the CAC (22-25).

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Nucleic Acid Thermal Denaturation. The nucleic acid binding capability of 4 was next evaluated by studying the thermal denaturation profile of a polyApolyU double helix in the presence of varying amounts of glycosyl nucleoside amphiphile 4. The denaturation profiles and corresponding first derivatives obtained in the absence or presence of magnesium are presented in Figure 3. Specifically, panels Ia, IIa, and IIIa of Figure 3 show the thermal denaturation profiles of polyA-polyU at 260 nm in the presence of compound 4, glucose, and uridine, respectively. The first derivatives of the corresponding absorbances versus temperature are presented in panels Ib, IIb, and IIIb of Figure 3. Likewise, thermal denaturation experiments with polyA-polyU and amphiphile 4 in the presence of a magnesium buffer are shown in Figure 3, panel IVa. The corresponding first derivative is plotted in Figure 3, panel IVb. The DNA thermal denaturation results are summarized in Table 1. In a HEPES buffer (HEPES 0.02 M, pH 7.4, NaCl 0.01 M), the Tm measured for polyA-polyU is 43.4 °C. A positive shift, ∆Tm [∆Tm ) Tm(4 + polyA-polyU) - Tm-

Figure 3. Hyperchromicity at 260 nm versus temperature of polyA-polyU in the presence of compound 4, glucose, and uridine (respectively, panels Ia, IIa, and IIIa), first derivatives of absorbance versus temperature (respectively, panels Ib, IIb, and IIIb) in aqueous HEPES buffer 0.02 M, pH 7.4, NaCl 0.01 M). Same experiments with polyA-polyU and amphiphile 4 in the presence of magnesium (IVa) and their first derivatives (IVb) are also shown (performed in aqueous HEPES magnesium buffer 0.02 M, pH 7.4, NaCl 0.01 M, MgCl2 0.01 M).

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Table 1. DNA Thermal Denaturation Results ∆Tma (°C) amount of compd (equiv)

4+ polyA-polyUb

4+ CT-DNAc

4+ polyA-polyU + Mgd

uridine + polyA-polyUb

glucose + polyA-polyUb

1 4 8 10

1.3 2.2 2.6 3.5

1.5 2.1 2.3 e

-0.5 -0.7 -0.8 e

-0.6 -0.7 -1.7 e

-0.5 -0.9 -1.2 e

a ∆T (°C) ) T of polyA-polyU + compound mixture - T of polyA-polyU (average of three experiments). The error in melting m m m temperature (Tm) does not exceed 0.2 °C. b Aqueous HEPES buffer (HEPES 0.02 M, pH 7.4, NaCl 0.01 M). Tm measured for polyA-polyU c in this buffer is 43.4 °C. Aqueous HEPES buffer (HEPES 0.02 M, pH 7.4, NaCl 0.01 M). Tm measured for CT-DNA in this buffer is 72.3 °C. d Aqueous HEPES magnesium buffer (HEPES 0.02 M, pH 7.4, NaCl 0.01 M, MgCl2 0.01 M). Tm measured for polyA-polyU in this buffer is 69.6 °C. e Not available.

(polyA-polyU)], of the polyA-polyU melting temperature is observed when compound 4 is added to the solution (Figure 3I) indicating a stabilization of the double helix (∆Tm ) 2.6 °C, 8 equiv of 4 for 1 equiv of RNA phosphate). Note that the same phenomenon was observed with a calf thymus (CT) DNA (∆Tm ) 2.3 °C, 8 equiv of 4 for 1 equiv of DNA phosphate). The ∆Tm increases as the number of amphiphile equivalents of 4 is increased suggesting that the stabilization of the double helix is not due to a one to one complex. Since these thermal denaturation experiments were performed with concentrations of amphiphile 4 higher than the CAC (1 equiv of 4 is 1.6 × 10-5 mol/L), the stabilization of the DNA double helix can be attributed to a DNA aggregation resulting from interactions between DNA and amphiphile aggregates. As expected, in the presence magnesium the Tm(polyA-polyU) is enhanced with a new value of 69.6 °C. It is well-known that magnesium ions increase Tm by binding to the phosphate backbone and stabilizing the double helix (26). When amphiphiles 4 is added to the Mg solution a decrease of the Tm(polyA-polyU) is noted. This small destabilization is consistent with the strong association of diols with the phosphate (27). In this case, sugar-phosphate associations compete with the Mgphosphate complex inducing Mg displacement and consequently a decrease in double helix stability. To estimate the impact of the nucleoside and sugar moieties on the stability of polyA-polyU double helix, the denaturation experiments were performed in the presence of uridine and glucose (without added Mg). A negative ∆Tm was observed for both compounds (Figure 3, panels II and III). These negative values can be rationalized by the polyol interactions with DNA inducing an enhancement of the repulsion between the negative charges of the phosphate groups (13). These results underline the importance of the amphiphilic character and the aggregate formation in stabilizing the DNA double helix. Quasi-Elastic Light Scattering (QELS) Studies. We evaluated the impact of the CT-DNA on amphiphile aggregation. The hydrodynamic radius (Rh) of the aggregates was measured at three time points (t ) 0, 24, and 36 h), and Table 2 summarizes the values obtained. Amphiphiles (typical concentration 0.6 mM) were extruded through a 50 nm filter in buffer (HEPES 0.02 M, pH 7.4, NaCl 0.01 M), DNA was then added to the aqueous solutions (DNA phosphate concentration 0.006 mM) containing the amphiphiles. Extruded amphiphile solutions exhibit stable aggregates with hydrodynamic radius of roughly 20 nm even after 36 h (Rh ) 19.8 ( 0.3) In the presence of CT-DNA after 36 h, the observed Rh increases to 42.6 ( 1.8 nm. It is reasonable to suggest that this size enhancement is due to the formation of new structures composed of amphiphiles and DNA.

Table 2. Hydrodynamic Radius (Rh, nm) of Amphiphiles and CT-DNA/Amphiphile Aggregatesa t (h)

DOUGluc (4)

DOUGluc (4) + CT-DNA

0 24 36

23.1 ( 1.2 21.2 ( 0.3 19.8 ( 0.3

22.1 ( 0.4 22.2 ( 0.8 42.6 ( 1.8

At t ) 0, 24, and 36 h of 4 in the absence or presence of CTDNA. Average of three experiments realized in aqueous buffer (HEPES 0.02 mM, pH 7.4, NaCl 0.01 mM). Value measured for CT-DNA alone (type XV activated, Sigma) is 3.0 ( 0.5 nm.

Figure 4. Transmission electronic microscopic (TEM) images of (a) small unilamellar vesicles obtained after extrusion of 4 through a 50 nm filter (bar ) 50 nm) and (b) large aggregates (4 equiv/CT-DNA ) 50, bar ) 100 nm).

TEM Studies. To confirm the impact of nucleic acids on amphiphile self-assembly, aqueous solutions of 4 were studied either in the absence or in the presence of CTDNA by transmission electronic microscopy (TEM). As shown in Figure 4a, the extrusion of 4 through a 50 nm filter gives small unilamellar vesicles (SUV). The same solution was also studied after addition of calf thymus DNA. In agreement with the results obtained by QELS experiments, the presence of CT-DNA affects the supramolecular organization. A new population of larger aggregates is present in the solution. TEM images show a decrease in vesicle population and larger aggregates of roughly 100 nm (Figure 4b). Gel Electrophoresis Studies. Electrophoresis gel shift is a convenient assay of DNA complexation. Thus, in light of the UV-vis, QELS, and TEM studies, agarose gel electrophoresis experiments were conducted by using plasmid pUC18 DNA. Increasing amounts (∼0, 5, 10, 15, 25, 50, and 75 equiv based on the amphiphile 4/phosphate, which correspond to the following concentrations: 0, 0.3, 0.6, 0.8, 1.4, 2.7, and 4.0 mM, respectively) of 4 were mixed with plasmid DNA (plasmid DNA phosphate concentration 0.06 mM) in HEPES buffer (HEPES 0.02 M, pH 7.4, NaCl 0.01 M). After incubation for 36 h at room temperature, the mixtures (20 µL) were loaded into a 1% agarose gel (HEPES 0.02 M, pH ) 7.4, NaCl 0.01 M, EtBr 500 ng/mL) using a pipettor. The gel was run at constant power (60 V) and at room temperature for 2.5 h. The DNA-amphiphile complexation is readily monitored under UV. High amphiphile ratios

Nucleic Acid Complexing Glyco-nucleo-amphiphile

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Figure 5. Electrophoretic gel shifts for the complex plasmid4: from lane 1 to lane 7, decreasing amounts of 4 from 75 to 0 equiv.

(50-75 amphiphile/phosphate equiv) behave similarly, and the DNA is immobilized as typically shown in Figure 5. The glycosylated nucleoside amphiphile 4 immobilizes plasmid pUC18 DNA demonstrating the masking of DNA charges by the uncharged amphiphile aggregates. 31 P NMR and FTIR Studies. To determine whether the stabilization of the DNA double helix depends on both glucose-phosphate interactions and amphiphile selforganization, the binding of Na2HPO4 (guest, [G]t ) total guest concentration) with glucose, 2, and 4 (hosts, [H]t ) total host concentration) were studied by 31P NMR spectroscopy model systems. NMR titration of phosphate with sugars was performed by keeping the total phosphate concentration constant ([Na2HPO4] ) 3.0 × 10-3 M) and gradually increasing the sugar derivative concentrations. These studies reveal that the complexation of the phosphate anion is occurring only in the case of the amphiphile. The 31P NMR signal undergoes upfield shifts in the presence of 4 host until saturation is reached at [H]t/[G]t ratios higher than 1 (Figure 6a), whereas no significant chemical induced shift is observed for glucose and compound 2. Such a result indicates that both the glucose moiety and amphiphilic character are needed to induce the formation of a glucose-phosphate complex in aqueous media. Fifty percent complexation occurs for 4 host at [H]t/[G]t ) 0.5 (Figure 6a), that is, at host concentration of [H]50 ) 1.5 mM ([G]t ) 3 mM). The implication of [H]50 is that Kmin ) 1/[H]50 ≈ 6 × 102 M-1 represents the lower limit of the binding constant of H for a molecule of G (where the equilibrium constant K ) [H-G]/[H][G], H + G a H-G).22 The binding affinity, Kmin, obtained for 4 is slightly lower than those for the oligosaccharide clusters-phosphate complexes (103 M-1) in aqueous media (28). A Job plot (29) for the binding of 4 and phosphate (Figure 6b) showed a maximum between X ) 0.5 and X ) 0.6, indicating a preference for 1:1 complexes over 2:1 complexes, which coexist in solution to a minor degree. From the NMR studies, the chemical induced shifts observed are due to phosphate anion coordination incorporated in the aggregate pools via hydrogen bonding. Indeed, in view of its amphiphilic nature, it is not surprising that host 4 forms aggregates, where the hydrogen bond forming anion may link to the sugar moieties. This is because the phosphate ion is associated with the sugar surface of the aggregates and a chemical induced shift is observed. Compound 2 does not possess a glucose moiety and therefore cannot bind phosphate anion. Consequently, the phosphate is not coordinated, and no significant chemical induced shifts are observed.

Figure 6. 31P NMR titration (a) of phosphate (guest, [G] ) 3 mM) with hosts ([H]) glucose (2, red), compound 2 (b, blue), and compound 4 (9, black) at 25 °C in D2O/DMSO-d6 with δp in reference to external H3PO4 in D2O (δp ) 0) and Job plot (b) for the binding of 4 and phosphate. Concentration of 4 and of phosphate was held constant at 3 mM.

Figure 7. FTIR spectra of (a) polyA-polyU, (b) 4, and (c) polyA-polyU/4 1:1 equiv in the region of 1350-800 cm-1.

The infrared spectra of the polyA-polyU and the polyA-polyU/4 (1:1 equiv) complexes in the region of

870 Bioconjugate Chem., Vol. 16, No. 4, 2005

Arigon et al.

Figure 9. CD spectra of polyA-polyU (blue) and 4/polyApolyU (4:1) (red) at 25 °C.

Figure 8. FTIR spectra in solution of (a) 4/polyA-polyU (1:1), (b) polyA-polyU, (c) 4, (d) 4/polyC-polyG (1:1), and (e) polyCpolyG in the region of 1500-1800 cm-1.

1350-800 cm-1 are shown in Figure 7. The band around 1080 cm-1 in the IR spectrum (Figure 7a) has been assigned to the symmetric PO2- stretching mode coupled with the C5′-O5′ vibrations (30), whereas the antisymmetric PO2- stretching vibrational band is observed around 1255 cm-1 (31). In the same region, 4 exhibits three bands at 1076 (C-O stretching primary alcohol), 1112 (C-O stretching secondary alcohol), and 1260 cm-1 (C-N-CdO vib) (32). The phosphate-sugar interactions are characterized by a shift of the antisymmetric PO2stretch at 1216 cm-1, which confirms the amphiphileRNA complex formation (33). FTIR spectra recorded in D2O solutions in the region 1500-1800 cm-1 are presented in Figure 8. In the case of the mixture 4/polyA-polyU (1:1) (Figure 8a), two additional bands at 1710 and 1683 cm-1 are observed. These absorptions are not observed for polyA-polyU (Figure 8b) and 4 (Figure 8c). According to Dagneaux et al., the absorption at 1712 cm-1 is assigned to a reverse Hoogsteen C2dO2 (thymidine, dT*dA-dT) stretching vibration (34). The band at 1683 cm-1 has been assigned to a C2dO2 stretch of additional base paired uridine (35). These results suggest that the base is also involved in the stabilization of the amphiphile/polyA-polyU complex via either Hoogsteen or reverse Hoogsteen nucleobase pairing with the uracil part of the amphiphile. A negative control using a mixture of 4/polyC-polyG (1:1) is shown in Figure 8d. This FTIR spectrum, which does not show

Figure 10. Interaction of (a) DNA-amphiphile, in which the presence of both hydrophobic interactions (amphiphiles-amphiphiles) and hydrogen bonding (sugars-phosphates) lead to a DNA double helix stabilization and (b) DNA-sugar, in which the sugar induces hydrogen bonding only, which gives a double helix destabilization.

any change compared to polyC-polyG (Figure 8e), confirms the Hoogsteen or reverse Hoogsteen nucleobase pairing observed in the case of the 4/polyA-polyU (1:1) mixture. Circular Dichroism Studies. The CD spectrum of polyA-polyU was recorded in the presence of 4. The CD characteristics of the polyA-polyU/4 complex are different from those of the polyA-polyU duplex. The CD spectrum of the polyA-polyU/4 complex (red curve) and that of the polyA-polyU duplex (blue curve) are depicted in Figure 9. The RNA-4 complex is characterized by a positive band at 260 nm and a negative band in the 245 nm region, and both of these bands have lower ellipticity values compared to that of the polyA-polyU duplex, which has a positive band in the 270 nm region, followed by a negative band at 245 nm. These results suggest that the polyA-polyU double helix structure is affected by the complexation with 4. According to the circular dichroism investigations on U-A duplex and U*A-U triplex reported by Ray et al. (36), the lower ellipticity values observed in the case of 4/polyA-polyU compared to

Nucleic Acid Complexing Glyco-nucleo-amphiphile

polyA-polyU duplex could be attributed to amphiphile U*A-U complex. Proposed 4-DNA Complex. The molecular model for DNA-amphiphile assemblies formed with the glucosyl uridine amphiphile is proposed in Figure 10. In the case of amphiphile 4, the stabilization of the double helix is due to concomitant phosphate-sugar and hydrophobic interactions (Figure 10a), whereas only phosphate-sugar interactions are responsible for DNA destabilization (37) by sugar or uridine (Figure 10b). CONCLUSION

We describe here a new neutral DNA amphiphile for binding to nucleic acid and subsequent supramolecular assembly formation. The synthetic route to the glucosyl dioleyl uridine amphiphile is reported, which should provide easy access to other nucleoside-based sugar amphiphiles. A critical aggregation concentration of 10-5 mol/L for 4 was determined by using fluorescence probing methods. Thermal denaturation experiments of a polyApolyU double helix in the presence of different amounts of 4 show a positive shift of ∆Tm indicating a double helix stabilization. The formation of amphiphile-nucleic acid complexes was confirmed by QELS, TEM, and gel electrophoresis studies. The 31P NMR investigations reveal that the phosphate anion is associated with the sugar surface of the glucosyl dioleyl uridine amphiphile aggregates. Similarly, the PO2- stretch at 1216 cm-1 of the polyA-polyU/4 (1:1 equiv) complex shows the presence of phosphate-sugar interactions. The bands at 1712 and 1683 cm-1 observed in aqueous solutions by FTIR and the lower ellipticity values measured by CD suggest that the uridine moiety of 4 is also involved in the stabilization of the amphiphile/polyA-polyU complex. All together, the results strongly support amphiphile-nucleic acid complex formation as a consequence of the amphiphilic character of the molecule, nucleoside, and phosphatesugar interactions. This glycosyl nucleoside-based amphiphile demonstrates an alternative approach to forming stable nucleic acids supramolecular assemblies. These results expand the current repertoire of DNA complexing amphiphiles and provide further motivation for the design and evaluation of new amphiphiles. ACKNOWLEDGMENT

This research was supported by the Army Research Office (Grant DAAD 19-02-1-0386). The authors would like to thank Dr. Stephen J. Lee, Estelle Champion, Prof. Catherine Vieillescazes, Michael Hovaneissian, and Prof. Mohamed El Maataoui. LITERATURE CITED (1) Felgner, P. L., Gadek, T. R., Holm, M., Roman, R., Chan, H. W., Wenz, M., Northrop, J. P., Ringold, G. M., and Danielsen, M. (1987) Lipofection: a highly efficient, lipid mediated DNA-transfection procedure. Proc. Natl. Acad. Sci. U.S.A. 84, 7413-7417. (2) Leventis, R., and Silvius, J. R. (1990) Interactions of mammalian cells with lipid dispersions containing novel metabolizable cationic amphiphiles. Biochim. Biophys. Acta 1023, 124-132. (3) Gao, X., and Huang, L. (1991) A novel cationic liposome reagent for efficient transfection of mammalian cells. Biochem. Biophys. Res. Commun. 179, 280-285. (4) Lynn, D. M., Anderson, D. G., Putmann, D., and Langer, R. J. (2001) accelerated discovery of synthetic transfection vector: parallel synthesis and screening of a degradable polymer library. J. Am. Chem. Soc. 123, 8155-8156.

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