A Novel Strategy to Construct a Flat-Lying DNA Monolayer on a Mica

May 12, 2006 - Flat-lying, densely packed DNA monolayers in which DNA chains are well organized have been successfully constructed on a mica surface b...
33 downloads 0 Views 1MB Size
Download PDF
10792

J. Phys. Chem. B 2006, 110, 10792-10798

A Novel Strategy to Construct a Flat-Lying DNA Monolayer on a Mica Surface Yonghai Song, Zhuang Li,* Zhiguo Liu, Gang Wei, Li Wang, Lanlan Sun, Cunlan Guo, Yujing Sun, and Tao Yang State Key Laboratory of Electroanalytical Chemistry, Changchun Institute of Applied Chemistry, Graduate School of the Chinese Academy of Sciences, Chinese Academy of Sciences, Changchun 130022, Jilin ProVince, People’s Republic of China ReceiVed: NoVember 8, 2005; In Final Form: April 13, 2006

Flat-lying, densely packed DNA monolayers in which DNA chains are well organized have been successfully constructed on a mica surface by dropping a droplet of a DNA solution on a freshly cleaved mica surface and subsequently transferring the mica to ultrapure water for developing. The formation kinetics of such monolayers was studied by tapping mode atomic force microscopy (TMAFM) technique. A series of TMAFM images of DNA films obtained at various developing times show that before the sample was immersed into water for developing the DNA chains always seriously aggregated by contacting, crossing, or overlapping and formed large-scale networks on the mica surface. During developing, the fibers of DNA networks gradually dispersed into many smaller fibers up to single DNA chains. At the same time, the fibers or DNA chains also experienced rearrangement to decrease electrostatic repulsion and interfacial Gibbs free energy. Finally, a flat-lying, densely packed DNA monolayer was formed. A formation mechanism of the DNA monolayers was proposed that consists of aggregation, dispersion, and rearrangement. The effects of both DNA and Mg2+ concentration in the formation solution on DNA monolayer formation were also investigated in detail.

Introduction Molecular monolayers are attracting considerable interests in a variety of interface studies, including wetting, lubrication, and molecular recognition,1-5 owing to their well-ordered organization, high homogeneity, and molecular dimensions.6-8 The preparation and formation mechanisms of molecular monolayers on a solid surface have been extensively investigated.9-19 These studies are mostly focused on small molecules because they can spontaneously form molecular monolayers on a solid surface by direct adsorption from a solution. For long flexible macromolecules such as polymer and DNA molecules, it is difficult to spontaneously form molecular monolayers on a solid surface by simple adsorption. DNA is a negatively charged biomolecule. For long DNA molecules, it has an extraordinary large length-to-width ratio. Its ideal duplex-helical structure (stacked base pairs on the inside and charged phosphate backbone on the outside20) enables it to recognize its complementary strand and transfer electrons.21 The monolayers of such molecules may have potential applications as biomaterials in medicine and engineering. However, some factors hinder DNA macromolecule from spontaneously forming molecular monolayers on a solid surface by simple adsorption. First, DNA always exists in a variety of randomly coiled conformations in a solution.22 When the molecule is deposited on a solid surface, the aggregation and entanglement of DNA chains are hardly avoidable,23 which makes it difficult to form molecular monolayers. Second, high DNA concentration is indispensable to ensure enough DNA molecules for forming densely packed monolayers. However, under high DNA concentrations, DNA molecules tend to contact, cross, or overlap each other and form networks when they are deposited on a solid surface. * Author to whom correspondence should be addressed. Phone/Fax: +86-431-5262057. E-mail: [email protected].

Third, salt cations in a DNA solution can enhance aggregation of DNA chains during adsorption by partially screening electrostatic repulsions between the negatively charged phosphate backbones.24 Last, some sticky ends, which are created by breaking the ends of linear DNA and DNA chains during sample handling, facilitate the aggregation of DNA chains by binding other chains to form triple-stranded DNA or hybridizing complementary bases of other ends,25,26 although the effect is very weak. As a result, large-scale, two-dimensional DNA networks have been extensively observed on a solid surface.27-36 Recently, flat-lying, well-ordered DNA monolayers have been constructed on cationic lipid membranes.37-41 The formation of excellent monolayers was mainly attributed to the fluidity of the membranes. The diffusive movements of lipids give rise to membrane-induced, attractive interactions that balance the direct repulsive interactions between adjacent molecules, forming a well-ordered organization of DNA molecules. According to this mechanism, analogous structures are difficult to form on a solid surface, since the solid surface has no fluidity. In this paper, we develop a simple and easy method to fabricate flat-lying, densely packed DNA monolayers on a solid mica surface. This strategy includes dropping a droplet of DNA solution with Mg2+ on a freshly cleaved mica surface and subsequently immersing the sample in ultrapure water to develop. The DNA monolayers obtained by this method have some similarity to those on lipid membranes.37-41 The formation process of the DNA monolayers was investigated by tapping mode atomic force microscopy (TMAFM). The possible formation mechanisms and the effects of both DNA and Mg2+ concentration on the DNA monolayer formation were also discussed in detail. The obtained information may improve our understanding on how to prepare long flexible macromolecular monolayers on a solid surface and will also accelerate the application of DNA monolayers.

10.1021/jp0564344 CCC: $33.50 © 2006 American Chemical Society Published on Web 05/12/2006

Construction of a DNA Monolayer on a Mica Surface Experimental Sections Chemicals. λ-DNA (48 502 bp) was purchased from SinoAmerican Biotechnology Company (Beijing, China) and used without further purification. Magnesium acetate (Mg(OAc)2) was purchased from Aldrich. Other chemicals were obtained from Beijing Chemical Reagent Factory (Beijing, China). Ultrapure water (18.2 MΩ cm) was prepared with a quartz distillatory below its boiling point and then purified with a Milli-Q water purification system (Millipore Co. Ltd.). Muscovite mica (KAl2(AlSi3)O10(OH)2,V-1 grade) purchased from Linhe Street Commodity Marketplace (Changchun, China) was cut into about 1.2 × 1.2 cm2 square pieces as substrates. Both sides of the mica surface were freshly cleaved before use. Solution Preparation. The 400 ng/µL stock solution of λ-DNA (10 mM Tris-HCl, 1mM EDTA, pH 7.5) was diluted to different concentrations with ultrapure water. And then the diluted DNA solutions were mixed with Mg(OAc)2 solutions in equal volumes as DNA monolayer forming solutions. Formation of DNA Monolayers. After these forming solutions were incubated at 38 °C for 1 h, a droplet of the forming solution (20 µL) was dropped onto a freshly cleaved mica surface and remained there for about 8 min for DNA adsorption. Then the sample was immersed in ultrapure water for DNA monolayers developing for different times. After the sample was taken out of the ultrapure water, it was rinsed with anhydrous ethanol, which both immediately stopped the monolayers developing because of rapid diffusion of water molecules into anhydrous ethanol and enhanced stabilization of the DNA films.42 Finally, the sample was dried under air and prepared for TMAFM imaging. In a control experiment, to confirm the effect of the removal of salt ions on the formation of DNA monolayers, the sample was immersed in a concentrated salt solution composed of 10 mM Tris-HCl, 1 mM EDTA, and 1 M NaCl (pH 8.05) to develop for 24 h. The longer developing time was used here to ensure the balance of salt ions in the system. Atomic Force Microscopy Meausurements. All AFM experiments were carried out using a Digital Instruments Nanoscope IIIa (Santa Barbara, CA). Typical AFM images were acquired in tapping mode with silicon (Si) cantilevers (spring constant, 20-100 N/m) below their resonance frequency (typically, 200-400 kHz) at room temperature under ambient conditions. All AFM images were raw data except for flattening. The surface coverage and height of the DNA monolayers were carefully measured by using Bearing and Section analysis software. All average values were measured at least from five different AFM images. Results and Discussion AFM Images of DNA Monolayer Formation. AFM is one of the most adequate techniques for direct observation of molecular monolayer formation due to its extraordinary resolution and precision.43,44 TMAFM was used here to monitor the DNA monolayer formation process because it exerts a minimum scanning force to samples and can keep the integrity of the samples.45,46 Figure 1 is a series of TMAFM images of DNA films obtained at various developing times in ultrapure water. All of the images are typical of those obtained from at least five separated samples. Figure 1a is a TMAFM result of a control experiment in which the sample was not treated by immersion in ultrapure water. The DNA chains obviously aggregated by contacting, crossing, or overlapping each other and formed large-scale DNA networks. The crossing has been believed to be a common structure for simple adsorption, since

J. Phys. Chem. B, Vol. 110, No. 22, 2006 10793 statistically it bares more weight than the parallel arrangement of DNA.38 Such DNA networks have many big meshes that appeared as the dark regions in the image, thus exposing large parts of the mica surface. Figure 1b is a TMAFM image of a sample that was treated by immersion in ultrapure water for 40 s, which indicates that the mesh size is significantly reduced and the surface area of bare mica is also reduced. For the sample that was treated by immersion in ultrapure water for 5 min, the TMAFM image as shown in Figure 1c indicates a further decrease of the mesh size and bare mica surface area. After 30 min of immersion, a DNA monolayer appeared (Figure 1d, it was proved to be a monolayer in the following text). As shown in the enlarged image (Figure 1e), DNA chains in the monolayers are densely packed and well organized, with only a few strands still crossed or overlapped. No obvious change was observed as the immersion time was further elongated to 1 h (Figure 1f) or longer. The heights of the fibers in these DNA films were measured from cross-section analysis of TMAFM images, and the histogram of their distribution is shown in Figure 2. Without immersion in ultrapure water, the height distribution of the fibers is scattered, and the main height is approximately 2.44 ( 0.25 nm (Figure 2a). This value is 5 times that of the single DNA chain height that is reported to be about 0.5 nm by TMAFM in air.47,48 Thus these fibers might be overlapped DNA bundles with five or more DNA chains. With the immersion in ultrapure water carried out, the height distribution of the fibers became narrower, and its main height gradually decreased to 1.01 ( 0.10 nm (Figure 2b) when the immersion time was 40 s. Such changes suggested that the bundles in DNA networks as shown in Figure 1a were dispersing into smaller bundles composed of fewer DNA chains. Especially, after 5 min of immersion, the main height was decreased to 0.56 ( 0.06 nm (Figure 2c), indicating the appearance of many single double-stranded DNA chains. Finally, about 90% of the height was located at around 0.56 ( 0.06 nm after immersion for 30 min (Figure 2d). It can be concluded that a flat-lying DNA monolayer was formed, and only a few chains crossing or overlapping were left in this stage. After that, the height distribution of the fibers and its main height had no distinguishable change when the immersion time was further increased (Figure 2e). It is also noticed that about 10% of the height is still higher than 0.56 ( 0.06 nm even after 24 h of immersion. It might be caused by some DNA molecules that failed to rearrange themselves into separate ones and some sticky ends created by breaking the DNA chains during sample handling that bound other chains to form triple-stranded DNA.25,26 The rearrangement of adsorbed DNA molecules will be discussed in detail in the following. Figure 3 shows the plot of the estimated surface coverage of the DNA films as a function of developing or immersion time. The surface coverage was estimated as below

Γe )

Γm Ctb

where Γe is the estimated surface coverage, Γm is the measured surface coverage, and Ctb is coefficient of tip broadening (in this case, Ctb is estimated to be 5.9). The surface coverage rapidly increased from 5.33 ( 0.5% to 7.72 ( 0.8% in the initial 40 s. In the following 5 min of developing, the surface coverage continuously and gradually increased to 9.10 ( 0.9%. After that, the surface coverage approximately varied only 2.97%. The surface coverage was finally maintained at 12.07 ( 1.2%, regardless the increase of immersion time. These data suggest

10794 J. Phys. Chem. B, Vol. 110, No. 22, 2006

Song et al.

Figure 1. Typical TMAFM images of DNA films formed in ultrapure water for various developing times: 0 s (a), 40 s (b), 5 min (c), 30 min (d), and 60 min (f). Part e is an enlarged image of part d. The vertical scales are 10 nm (a), 5 nm (b), and 3 nm (c-f), respectively. The DNA and Mg2+ concentrations used in this study are 12.5 ng/µL and 1 mM, respectively.

that the formation rate of DNA monolayers is very rapid in the initial stage and then slows down gradually. Possible Formation Mechanism of DNA Monolayers. The formation of molecular monolayers is governed by intermolecular and interfacial interactions, and these interactions drive the system toward a minimum of the overall free energy. The formation of the DNA monolayers is just a result of equilibrium of all kinds of forces between DNA and mica or DNA. To immobilize negatively charged DNA onto negatively charged and hydrophilic mica surface,49 the Mg2+ is used here as a bridge ion between the phosphate groups of DNA and the mica surface.50,51 The immobilization force is the sum of the electrostatic attraction and the hydrophilic force between the mica surface and DNA.50,52 The force also induces the attractions between DNA molecules by a similar mechanism. Especially, when the DNA and salt ion concentrations are high, the

attraction force usually induces the formation of DNA networks on the mica surface. Moreover, some sticky ends created by breaking the DNA chains also enhanced the aggregation of DNA chains on the mica surface,27 although its effect is very weak. All these interactions were randomly exerted on DNA molecules and the mica surface. Thus the DNA chains in DNA networks are highly disordered, and the height distribution of fibers in DNA networks is very scattered. These fibers are usually composed of several DNA chains. However, the so-called electrical double-layer force53 repels these surfaces between DNA and mica or DNA. This force is the sum of the electrostatic repulsion between the same charged ions and thermal pressure.50 The impetus for the dispersion of DNA bundles just comes from these forces after the sample was immersed in ultrapure water for developing. The salt ions that adsorbed on the DNA molecules and the mica surface or

Construction of a DNA Monolayer on a Mica Surface

J. Phys. Chem. B, Vol. 110, No. 22, 2006 10795

Figure 4. Typical TMAFM image of the DNA network formed by immersing the sample in 10 mM Tris-HCl, 1 mM EDTA, and 1 M NaCl buffer (pH 8.05) to develop for 24 h. The DNA network was almost maintained. The vertical scale is 5 nm. The DNA and Mg2+ concentrations used in this study are 12.5 ng/µL and 1 mM, respectively.

Figure 2. Histograms of the height distribution for the fibers in DNA films formed in ultrapure water for various developing times: 0 s (a), 40 s (b), 5 min (c), 30 min (d), and 60 min (e).

Figure 3. Plot of the estimated surface coverage (Γe) of DNA films as a function of developing time in ultrapure water. The DNA and Mg2+ concentrations used in this study are 12.5 ng/µL and 1 mM, respectively.

bridged DNA-mica and DNA-DNA surfaces gradually diffuse into water due to the concentration difference between the DNA-DNA and DNA-mica interfaces and the bulk water. The removal of salt ions increases the repulsion forces24 and rapidly decreases the electrostatic attraction force between the DNA chains. When the repulsion forces become the main forces between them, the DNA bundles gradually disperse into many smaller bundles up to a single molecular chain. This conclusion was further confirmed by the fact that the DNA networks were maintained after 24 h of immersion in a high salt concentration solution (as shown in Figure 4). According to this assumption, the formation rate of the DNA monolayers should be fast in

the initial stage and then gradually become slow since the diffusion of salt ions is rapid at the initial stage because of the high concentration difference, which is very consistent with our TMAFM observation (Figure 3). However, it is hard to form a flat-lying, well-organized monolayer only by the simple diffusion of salt ions because of the crossing and overlapping of DNA chains, suggesting another mechanism might be also involved in the formation of DNA monolayers. Previous works37-41 have proved that the fluidity of the membrane is essential to induce the formation of the densely packed, flat-lying DNA monolayers on cationic lipid membranes. While our experiment showed that the densely packed, well-organized DNA monolayers can be formed on the mica surface, the mica surface does not possess fluidity. It might be possible that the formation of DNA monolayers on the solid surface is due to the DNA’s lateral movement. A statistical polymer chain analysis and high charge density of DNA have also proved that under the appropriate conditions the DNA molecules can arrange themselves on a solid surface before they are captured in the lowest energy conformations.23,54 Thus, in the process of monolayer formation, the adsorbed DNA molecules or fibers may experience rearrangement to decrease the electrostatic repulsion and interfacial Gibbs free energy due to weak binding to the mica surface by Mg2+ bridges in water.51 During reorientation and equilibration, some segments of DNA chains or fibers may detach from the mica surface, which facilitates the whole chain or fibers rebinding in a new conformation at which the electrostatic repulsions are decreased to minimum value. For long times of immersion, the disorganized DNA chains have enough time to rearrange, finally arriving at the most stable configuration: well-organized DNA monolayers (thermodynamically stable state). So we tentatively assume that DNA monolayer formation exhibits a mechanism of aggregation, dispersion, and rearrangement. It must be pointed out that there are some disadvantages in our ex situ TMAFM observations. For example, we cannot follow specific regions of the surface as they evolve so that the observed morphology of the uncompleted monolayers may not represent the actual structure in ultrapure water. The process of removal, rinsing, and drying can dramatically affect the morphology of film, which causes some differences between

10796 J. Phys. Chem. B, Vol. 110, No. 22, 2006

Song et al.

Figure 5. Typical TMAFM images of DNA monolayers prepared from various DNA concentrations: 2.5 ng/µL (a), 5 ng/µL (b), 7.5 ng/µL (c), 12.5 ng/µL (d), 15 ng/µL (e), and 25 ng/µL (f). The vertical scales are 3 nm. The Mg2+ concentration used in this study is 1 mM. The developing time for all these samples was 30 min.

the structures that we have observed and that of the actual DNA monolayers. However, a lot of information on the formation process of the DNA monolayers can be still revealed by ex situ TMAFM studies.15,17,55,56 Effect of DNA Concentration on DNA Monolayer Formation. The study of the effect of DNA concentration on DNA monolayer formation is very important because low DNA concentrations cannot produce densely packed DNA monolayers due to a lack of sufficient DNA molecules. Figure 5 is a series of typical TMAFM images of DNA monolayers constructed by using DNA solutions with different concentrations. Under low DNA concentrations (2.5 ng/µL), the DNA molecules did not cover the whole mica surface, and the compactness of the DNA monolayers was very low (as shown in Figure 5a). The DNA chains were loosely and randomly dispersed on the mica surface, and the distances between the chains were uncertain. With DNA

concentrations increasing to 5, 7.5, 12.5, and 15 ng/µL, the compactness of the DNA monolayers gradually increased (as shown in Figures 5b-e). The interchain distances decreased accordingly and tended to be uniform. With a further increase of the DNA concentration to 25 ng/µL, the compactness of the DNA monolayers and the interchain distances kept on changing (Figure 5f). The interchain distances have transformed into uniformity, which was confirmed by the diffraction ring in the Fourier transform image, as shown in Figure 6 (the average radii of the ring are 7.46 nm in the vertical axis and 7.51 nm in the horizontal axis, respectively). The uniformity of the interchain distances is the result of the equilibrium of all interactions between DNA chains. Such an arrangement decreased interactions between DNA chains to a minimum value and produced the most stable configuration: well-organized DNA monolayers. It is obvious that the uniformity of the interchain distances is

Construction of a DNA Monolayer on a Mica Surface

J. Phys. Chem. B, Vol. 110, No. 22, 2006 10797

Figure 7. Plot of the estimated surface coverage (Γe) obtained at different DNA concentrations versus DNA concentration (ng/µL). The Mg2+ concentration used in this study is 1 mM. The developing time was 30 min.

Figure 6. Enlarged image of Figure 5f (a) and its corresponding Fourier transform image (its range from 500 nm (center) to 1.95 nm (periphery)) (b). The arrows indicated that the average radii of the ring are 7.46 nm in the vertical axis and 7.51 nm in the horizontal axis, respectively.

strongly dependent on DNA concentrations, since the compactness of the DNA monolayers increased as DNA concentrations increased. Some chains became indistinguishable because of small interchain distances and the size limitation of the AFM tip. Therefore, the effect of higher concentration on the packing density of DNA monolayers cannot be further discussed in the present study. The estimated surface coverage of DNA monolayers as function of DNA concentration was plotted in Figure 7. The surface coverage of DNA monolayers rapidly increased as DNA concentration increased in the range of 2.5-15 ng/µL. Above 15 ng/µL, the effect of DNA concentration on the compactness of DNA monolayers becomes very weak, suggesting that 15 ng/µL is the minimal concentration for the formation of densely packed monolayers. Although higher concentration (higher than 15 ng/µL) slightly affects the compactness of DNA monolayers, it is very indispensable for the ordered monolayer formation. It is noticeable that the surface coverage of DNA monolayers only reaches about 20% when the DNA concentration is 25 ng/µL. It is assumed that the electrical double-layer force between DNA chains is the main factor to prevent the formation of more densely packed monolayers or multilayers.

Figure 8. Plot of the estimated surface coverage (Γe) of DNA monolayers formed in solutions with different Mg2+ concentrations versus the Mg2+ concentration. The DNA concentration used in this study is 7.5 ng/µL. The developing time was 1 h.

Optimal Mg2+ Concentration for DNA Monolayer Formation. To ascertain the optimal Mg2+ concentration for DNA monolayer formation, a series of samples prepared in solutions with different Mg2+ concentrations (varied from 0.5 to 6 mM in this study) were imaged by TMAFM (images were not shown here). The estimated surface coverage as a function of Mg2+ concentration was plotted in Figure 8. The surface coverage slightly increased as Mg2+ concentrations increased from 0.5 to 1 mM, which suggests that the surface coverage is sensitive to Mg2+ concentrations among this region. When Mg2+ concentrations were above 1 mM, no significant change was observed, indicating that 1 mM Mg2+ is enough for DNA monolayer formation. It is similar to the previous results that the divalent cationic concentration for maximal DNA binding onto the mica surface is 1 mM.51 Hence, the optimal Mg2+ concentrations for DNA monolayer formation should be 1 mM. Conclusions In summary, flat-lying, densely packed DNA monolayers have been constructed on the mica surface by dropping a droplet of a DNA solution on freshly cleaved mica and subsequently

10798 J. Phys. Chem. B, Vol. 110, No. 22, 2006 transferring the mica to ultrapure water for developing. TMAFM images of partially formed monolayers created at various developing times show that the monolayer formation is controlled by thermodynamics. The formation mechanism of DNA monolayers consists of aggregation, dispersion, and rearrangement of the DNA chains. It is seemed that 30 min of developing is enough to form well-organized monolayers. The uniformity of the interchain distances strongly depends on DNA concentrations. The compactness of DNA monolayers rapidly increases as the DNA concentration increases from 2.5 to 15 ng/µL and then becomes slow. The optimal Mg2+ concentration for DNA monolayer formation is 1 mM. This information may help to fabricate long flexible macromolecules monolayers and improve the knowledge of the mechanisms. Further studies will be focused on the effects of other factors such as temperature, divalent cations, and species of DNA on monolayer formation. Acknowledgment. This work was supported by the National Natural Science Foundation of China and the State Key Laboratory of Electroanalytical Chemistry. We are grateful to Dr. Li Wang (Tohoko University, Japan) for helpful discussions. We especially appreciate our two anonymous referees for their valuable comments and suggestions. References and Notes (1) Rabe, J. P.; Buchholz, S. Science 1991, 253, 424-427. (2) Morita, M.; Koga, T.; Otsuka, H.; Takahara, A. Langmuir 2005, 21, 911-918. (3) Zhang, Q.; Archer, L. A. J. Phys. Chem. B 2003, 107, 1312313132. (4) Houston, J. E.; Kim, H. I. Acc. Chem. Res. 2002, 35, 547-553. (5) Katz, E.; Willner, I. Angew. Chem., Int. Ed. 2000, 39, 1180-1218. (6) Swalen, J. D.; Allara, D. L.; Andrade, J. D.; Chandross, E. A.; Graoff, S.; Israelachvili, J.; McCarthy, T. J.; Murray, R.; Pease, R. F.; Rabolt, J. F.; Wynne, K. J.; Yu, H. Langmuir 1987, 3, 932-950. (7) Godinez, L. A. Res. Soc. Quim. Mex. 1999, 43, 219-229. (8) Ulman, A. An Introduction to Ultrathin Organic Films; Academic Press: San Diego, CA, 1991. (9) Wang, H.; Chen, S.; Li, L.; Jiang, S. Langmuir 2005, 21, 26332636. (10) Peng, Z.; Dong, S. Langmuir 2001, 17, 4904-4909. (11) Love, J. C.; Wolfe, D. B.; Haasch, R.; Chabinyc, M. L.; Paul, K. E.; Whitesides, G. M.; Nuzzo, R. G. J. Am. Chem. Soc. 2003, 125, 25972609. (12) Tian, Y.; Ng, Q. Y.; Fendler, J. H. Langmuir 1998, 14, 30673070. (13) Lim, J. C.; Neuman, R. D.; Park, S. Langmuir 2002, 18, 61256132. (14) Zamlynny, V.; Zawisza, I.; Lipkowski, J. Langmuir 2003, 19, 132145. (15) Wang, L.; Song, Y. H.; Han, X. J.; Zhang, B. L.; Wang, E. K. Chem. Phys. Lipids 2003, 123, 177-185. (16) Sparr, E.; Eriksson, L.; Bouwstra, J. A.; Ekelund, K. Langmuir 2001, 17, 164-172. (17) Wang, L.; Jiang, J. G.; Song, Y. H.; Zhang, B. L.; Wang, E. K. Langmuir 2003, 19, 4953-4957. (18) Wang, L.; Wang, E. K. Langmuir 2004, 20, 2677-2682. (19) Wang, L.; Song, Y. H.; Zhang, B. L.; Wang, E. K. Thin Solid Films 2004, 1-2, 197-202. (20) Watson, J. D.; Crick, F. H. L. Nature 1952, 171, 737-738. (21) Boon, E. M.; Jackson, N. M.; Wightman, M. D.; Kelley, S. O.; Hill, M. G.; Barton, J. K. J. Phys. Chem. B 2003, 107, 11805-11812.

Song et al. (22) Xiao, Z. W.; Xu, M. X.; Sagisaka, K.; Fujita, D. Thin Solid Films 2003, 438-439, 114-117. (23) Xiao, Z. W.; Xu, M. X.; Ohgi, T.; Sagisaka, K.; Fujita, D. Superlattices Microstruct. 2002, 32, 215-220. (24) Saenger, W. Principles of Nucleic Acid Structure; Cantor, Ch. R., Ed.; Springer Advanced Texts in Chemistry; Springer-Verlag: New York, 1984. (25) Re´vet, B.; Fourcade, A. Nucleic Acids Res. 1998, 26, 2092-2097. (26) Cohen, S. N.; Chang, A. C.; Boyer, H. W.; Helling, R. B. Proc. Natl. Acad. Sci. U.S.A. 1973, 70, 3240-3244. (27) Wu, A. G.; Li, Z.; Yu, L. H.; Wang, H. D.; Wang, E. K. Anal. Sci. 2001, 17, 583-584. (28) Wu, A. G.; Li, Z.; Yu, L. H.; Zheng, J. P.; Wang, E. K. Analyst 2002, 127, 585-587. (29) Ye, J. Y.; Umemura, K.; Ishikawa, M.; Kuroda, R. Anal. Biochem. 2000, 281, 21-25. (30) Jo, Y. S.; Lee, Y.; Roh, Y. Mater. Sci. Eng., C 2003, 23, 851855. (31) Jiang, X. H.; Lin, X. Q. Electrochem. Commun. 2004, 6, 873879. (32) Oliveira Brett, A. M.; Chiorcea, A. M. Langmuir 2003, 19, 38303839. (33) Chiorcea, A. M.; Oliveira Brett, A. M. Bioelectrochemisty 2004, 63, 229-232. (34) Kanno, T.; Tanaka, H.; Miyoshi, N.; Kawai, T. Appl. Phys. Lett. 2000, 77, 3848-3850. (35) Kanno, T.; Tanaka, H.; Miyoshi, N.; Kawai, T. Jpn. J. Appl. Phys. 2000, 39, L269-L270. (36) Cai, L.; Tabata, H.; Kawai, T. Appl. Phys. Lett. 2000, 77, 31053106. (37) Clausen-Schaumann, H.; Gaub, H. E. Langmuir 1999, 15, 82468251. (38) Fang, Y.; Yang, J. J. Phys. Chem. B 1997, 101, 441-449. (39) Fang, Y.; Yang, J. J. Phys. Chem. B 1997, 101, 3453-3456. (40) Mou, J.; Czajkowsky, D. M.; Zhang, Y.; Shao, Z. FEBS Lett. 1995, 371, 279-282. (41) Dan, N. Biophys. J. 1996, 71, 1267-1272. (42) Nakayama, Y.; Tanaka, H.; Kawai, T. Jpn. J. Appl. Phys. 2001, 40, L824-L825. (43) Zhou, D. J.; Sinniah, K.; Abell, C.; Rayment, T. Angew. Chem., Int. Ed. 2003, 42, 4934-4937. (44) Sennato, S.; Bordi, F.; Cametti, C.; Coluzza, C.; Desideri, A.; Rufini, S. J. Phys. Chem. B 2005, 109, 15950-15957. (45) Stroh, C.; Wang, H.; Bash, R.; Ashcroft, B.; Nelson, J.; Gruber, H.; Lohr, D.; Lindsay, S. M.; Hinterdorfer, P. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 12503-12507. (46) Dahlgren, P. R.; Lyubchenko, Y. L. Biochemistry 2002, 41, 1137211378. (47) Lyubchenko, Y. L.; Shlyakhtenko, L. S. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 496-501. (48) Thundat, T.; Allison, D. P.; Warmack, R. J. Nucleic Acids Res. 1994, 22, 4224-4228. (49) Sasou, M.; Sugiyama, S.; Yoshino, T.; Ohtani, T. Langmuir 2003, 19, 9845-9849. (50) Pastre´, D.; Pie´trement, O.; Fusil, S.; Landousy, F.; Jeusset, J.; David, M. O.; Hamon, L.; Cam, E. L.; Zozime, A. Biophys. J. 2003, 85, 25072518. (51) Hansma, H. G.; Laney, D. E. Biophys. J. 1996, 70, 1933-1939. (52) Arenzon, J. J.; Stilck, J. F.; Levin, Y. Eur. Phys. J. B. 1999, 12, 79-82. (53) Israelachvili, J. N. Intermolecular and Surface Force; Academic Press: San Diego, CA, 1992. (54) Rivetti, C.; Guthold, M.; Bustamante, C. J. Mol. Biol. 1996, 264, 919-932. (55) Xiao, X. D.; Liu, G. Y.; Charych, D. H.; Salmeron, M. Langmuir 1995, 11, 1600-1604. (56) Bierbaum, K.; Grunze, M.; Baski, A. A.; Chi, L. F.; Schrepp, W.; Fuchs, H. Langmuir 1995, 11, 2143-2150.