NANO LETTERS
Precise Positioning of Individual DNA Structures in Electrode Gaps by Self-Organization onto Guiding Microstructures
2004 Vol. 4, No. 4 607-611
Gunter Maubach† and Wolfgang Fritzsche* Institute for Physical High Technology, P.O. Box 100239, Jena, Germany Received January 5, 2004; Revised Manuscript Received February 13, 2004
ABSTRACT The ability to precisely position (bio)molecules onto microstructured substrates is a core technology critical to the long-term goal of molecular nanotechnology: the integration of molecular units in technical devices. Therefore, parallel processes have to be established to facilitate in a first step large-scale scientific investigations with statistically significant numbers, and later on technically relevant integration densities. DNA molecules were aligned by a receding meniscus generated by a drying droplet. Micromachined structures on the surface facilitated the alignment of DNA in the receding meniscus. The result was immobilized DNA following the electrode structures and spanning gaps. This self-assembly process presents a further step toward highly parallel processes for the precise positioning of molecular structures on chip surfaces.
Due to its unique properties, DNA is a potential building block for molecular nanotechnology. It provides highly specific connectivity, an established toolbox of enzyme-based (restriction, ligation) and technical (gel electrophoresis) manipulation techniques, and easy access to highly defined molecules in large numbers (custom synthesis, cloning). Experimental proofs of the possibilities include the creation of artificial 2D and 3D DNA structures,1,2 the positioning of DNA molecules in microelectrode gaps,3 the metallization of DNA,4-8 and the demonstration of a carbon nanotube transistor enabled by a DNA scaffold providing both the precise localization as well as the template for a metal wiring.9 There are two key aspects in immobilization of DNA on substrates: the extension of the rather flexible molecule, and the precision of positioning regarding location and orientation. The preparation of extended DNA was originally developed for in-situ hybridization for applications such as highresolution restriction maps of isolated chromatin and DNA molecules.10 Thereby, the released DNA binds along its length onto glass substrates that are either unmodified or chemically modified in order to enhance the adsorption process. The process can be supported by a controlled receding meniscus.11 Although extended DNA molecules provide an interesting system for molecular nanotechnology, their integration into * Corresponding author. † Present address: IBN Singapore, 10 Medical Drive, CRC #04-25, Singapore 117597. 10.1021/nl049968+ CCC: $27.50 Published on Web 03/06/2004
© 2004 American Chemical Society
technical setups such as microelectrode arrays is still hampered due to the undefined location of adsorption. The pattern of the adsorbed molecules does not match any predefined surface structures such as electrodes. On the other hand, procedures have been developed to bind long DNA molecules specifically on their ends. Typically, the specific interaction between complementary DNA sequences3 or between the negatively charged ends and a positively charged surface12 is used so the DNA molecules are pinned down between two predefined locations (e.g., chemically modified surface regions) in an extended state. These approaches allow a much more precise control of the location (and sometimes even the orientation) of immobilization. Refining the line electrode approach3 even further to electrodes of limited (lower micrometer range) size, the controlled immobilization of exactly one DNA molecule per gap has been achieved.12 However, the yield was rather low. In addition to the microstructured electrodes, chemical modifications of the electrode as well the substrate surface were required and a custom-built flow-through chamber had to be applied in a rather complicated process. The individual molecule in the gap was relatively fragile. Frequently, experiments (e.g., metallization prior to electrical characterization) failed because of damage to this molecule. The technique presented in this paper enhances this immobilization process considerably: The technology was improved so that it is more generally applicable and so the yield could be significantly increased, reaching often 100% of gaps with immobilized molecules. Now, a bundle of DNA
Figure 1. Droplet method to obtain aligned immobilization of long (lambda) DNA onto functionalized glass substrates. DNA-specific fluorescence. Overview (a), zoom in a peripheral region (b), and scanning electron micrograph (c) after metal particle labeling, (d) scheme of DNA orientation in the receding meniscus of a drying droplet.
is spanning the gap, providing a higher stability for further experiments and making detection, e.g., by fluorescence labeling, easier. Microstructured gold electrodes (100 nm thickness, about 1 µm width) on a silicon oxide substrate were prepared by sputtering and a standard photolithographic lift-off process. The silicon oxide surface was modified by silanization with ODTS (octadecyltrichlorosilane).13 Commercial available lambda-DNA was precipitated with 2-propanol, washed with 70% ice-cooled ethanol, and the pellet was then dissolved in sterile filtered water at a concentration of 250 ng/µL. To establish the method, preliminary trials were conducted using glass substrates modified with APTES ((3-aminopropyl)triethoxysilane) in 1 mM acetic acid.14 Droplets of 0.1 µL, with a diameter of about 1 mm, were spotted using a conventional pipet.15 The DNA stock solution (250 ng/µL) was 500 and 1000-fold diluted for these experiments. Droplets of 1.5 µL were applied to the substrates with microstructured electrodes. The DNA was labeled using the fluorescence dye YOYO-1 (Molecular Probes) either before or after application. A labeling with positively charged nanoparticles16 was conducted for SEM imaging. Fluorescence imaging was conducted using a microscope Axiotech (Carl Zeiss, Jena, Germany), equipped with a CCD camera Sensicam (PCO Computer Optics, Kehlheim, Ger608
many). High-resolution topographic images were obtained with a scanning force microscope NanoScopeIII with a Dimension 3100 measurement head (Digital Instruments, Santa Barbara, CA) in tapping mode. Scanning electron microscopy (SEM) imaging was performed with a digital scanning electron microscope DSM 960 (Carl Zeiss, Jena, Germany). In a first set of experiments, techniques for DNA stretching were adapted from the literature. We decided to follow the droplet procedure as described in the literature.16 The dried droplets showed typically an inhomogeneous image in the fluorescence (DNA) contrast (Figure 1a). The center of the droplet appeared bright (large concentration of DNA that could not be resolved), then a zone with radial extended fibers could be observed, and finally a surrounding bright line apparently caused by the droplet boundaries. Magnification of the fiber region by fluorescence or electron microcopy revealed the nearly parallel arrangement of bundled DNA structures (Figure 1b,c). A branching behavior was observed that was oriented toward the periphery of the droplet. The orderly arrangement of the fibers on this homogeneous substrate was a promising result for the further steps toward a controlled alignment of the fibers along predefined patterns. Nano Lett., Vol. 4, No. 4, 2004
Figure 2. Droplet incubation onto microstructured chip substrates with metal electrodes. DNA-specific fluorescence contrast. (a, b) Typical appearance of DNA immobilization by the droplet method on microstructured substrates; only occasionally the DNA and the electrode structures show some interaction resulting in changes of the DNA orientation (overview (a), zoom - (b)). (c, d) In this experiment, the local orientation of the DNA regarding the electrodes is controlled by the point of droplet deposition, and a parallel alignment of the DNA was achieved (overview (c), zoom (d)). Please compare the image in (b) for the electrode arrangement. (e) Occasionally, DNA structures appeared in a negative contrast as dark thread-like features. We assume that DNA was immobilized during the fluorescence labeling but accidentally removed during washing steps. The images show the background signal from unspecifically adsorbed dye molecules on the surface, and only the areas previously protected by DNA show no signal (dark). (f) Fluorescence signal from fibers crossing electrodes correlates with fiber thickness: Only thicker fibers exhibit sufficient signal to be detectable on such crossing points.
The method established on glass substrates was transferred onto chip substrates with microstructured electrodes. Again, the droplet procedure resulted in DNA immobilized in an oriented pattern reflecting the droplet boundaries (Figure 2a, DNA appears bright, the electrodes dark). A variation of the fiber thickness with the distance to the center can be observed. A zoom into the region with finer electrodes reveals DNA fibers (bright due to DNA-specific fluorescence) spread over the substrate surface with electrode structures (black). An even closer look shows regions where the DNA crosses the electrodes nearly perpendicular (Figure 2b, arrow). Here, the electrodes have apparently no influence on the DNA orientation. No interaction is observed, the DNA immobilization appears independent from the orientation of the underlying electrodes. Although the DNA fibers seem to cross the electrodes (see, e.g., Figure 2b, arrow), the brightness disappears as soon as the DNA reaches the electrode and reappears when the DNA leaves the electrode structure on the other side. Occasionally an influence of such crossing events on the orientation of DNA is observed. Then, the fibers seem to be pinned down by the electrodes, changing their orientation after crossing (e.g., arrowheads in Figure 2b). In these cases, the DNA crosses the electrodes in an angle much smaller than 90°. One explanation for this observation could be a weak interaction between the DNA and the electrodes during the process of immobilization. When the DNA is rather parallel to the electrodes, it can partially align and interact. For DNA perpendicular to the electrodes, the interaction is minimal and has no effect on the orientation. If this explanation is valid, it should be possible to align the DNA along the electrodes. Therefore, the droplet has to be positioned so that a region with parallel oriented DNA fibers is aligned with an electrode array. This Nano Lett., Vol. 4, No. 4, 2004
approach was tested; an overview of the immobilized DNA (bright) on an electrode array (similar to that shown in Figure 2a) is shown in Figure 2c. In the central region, the DNA is aligned parallel to an electrode structure (similar to the one visualized in Figure 2b). A zoom of this region is shown in Figure 2d. Clearly visible are the bright DNA fibers; only some electrode structures in the upper part are still visible as dark features. Comparing this image with the original electrode geometry (as, e.g., visible in Figure 2b), a high degree of co- localization of electrodes and DNA becomes apparent. The majority of the visible DNA is aligned on top of the underlying electrodes. The electrodes used for the alignment experiments contain a 2 µm gap (along the line of horizontal metal structures visible in Figure 2b and d). If the theory of the aligned immobilization due to attractive interaction between DNA and electrodes is valid, a small gap should not interfere significantly with the alignment process. The DNA is expected to adsorb on the electrodes above and below the gap, thereby spanning this region. To check for this behavior, the gap regions of the substrate shown in Figure 2c and d were studied in higher resolution. The zoomed fluorescence image of an electrode arrangement with oriented DNA is shown in Figure 3a. The bright fibrous structures are visible between the electrodes and usually aligned (parallel to the electrodes) on the substrate. Four electrode gaps are visible, and in every gap a small (about 2 µm length) but bright feature can be observed. What is the origin of this feature? The bright signal points to fluorescent labeled DNA. On the other hand, DNA is usually observed as fibrous structures with lengths of some tens of microns. However, as already mentioned, the DNA signal can be subdued when the DNA crosses the electrodes. Based on the orientation of the visible 609
Figure 3. DNA positioned in electrode gaps of 2 µm width. (a) Overview in DNA-specific fluorescence contrast. The gold electrode structures appear dark and the DNA bright, respectively. Four electrode gaps are visible: two of them are marked by arrows. (b, c) AFM (height mode) zooms of the two gaps marked in (a). The DNA appears as thread-like structures (arrows).
DNA, the possibility of low visibility of DNA on the electrodes, and the observed features in the gap, we assumed that some DNA fibers are immobilized on top of the electrodes, but are visible only in the short region spanning the gap. The remaining parts of the DNA are not detected due to quenching effects between the fluorescent dyes along the DNA and the electrode material (gold). To test this hypothesis, electrode gaps characterized by fluorescence microscopy were subsequently scanned by an AFM. Figure 3b shows a zoom of the left gap from Figure 3a. The DNA causing a bright signal in the fluorescence contrast appears now as a rod-like structure. In contrast to the fluorescence image, the AFM image shows that the DNA extends over the electrode structures. However, the AFM contrast for DNA is much higher in the gap compared to the contrast on top of the electrodes. An AFM zoom of the second gap (right arrow in Figure 3a) is presented in Figure 3c. Here, the DNA appears above and below the central electrode, which is compatible with the two bright features at these locations in Figure 3a. If one looks at the geometry of the electrode structures (upper part in Figure 3a), there are in general two possibilities for adsorption along the electrodes. These electrodes consist of two parts: an inner electrode that could be used for applying an electrostatic potential and a second electrode surrounding the first one. Adsorption of parallelaligned DNA could happen to both structures. If the DNA follows the outer electrode, it will cross only one gap, as is the case in the left gap. For DNA oriented along the inner electrode, two gaps have to be crossed, as visible in the second gap (also Figure 3c). We were interested in the elucidation of the course of the DNA at the electrodes. The AFM images in Figure 3b and c do not fully resolve DNA on top of the electrodes. An increased surface roughness pointing to deposited materials is apparent. So we decided to study a gap with DNA in 610
Figure 4. Detailed comparative ultramicroscopic characterization of DNA in an electrode gap. The same gap was visualized by DNAspecific fluorescence (a), height mode AFM (b), and SEM after metal particle labeling (c).
greater detail using a parallel approach by fluorescence microscopy, AFM and SEM (Figure 4). The fluorescence image (Figure 4a) shows two bright features spanning the gaps between the electrodes (dark structures). The AFM image (Figure 4b) resolves clearly thread-like structures between the electrodes at the locations of the strong Nano Lett., Vol. 4, No. 4, 2004
fluorescence signals. Following the directions of these molecules onto the electrodes, the molecule disappears. Because the surface roughness on the electrodes is higher than in the gap and the area of interest shows a higher density of amorphous structures, the AFM images do not exclude the presence of DNA on the electrodes. However, it cannot be resolved. So the DNA was labeled with nanoparticles and visualized by SEM (Figure 4c). The resulting image showed clearly the electrodes together with a DNA bridging both gaps. A closer view reveals that the DNA is only partially visible. AFM imaging (Supporting Information) yielded that this effect is due to incomplete coverage of the DNA with nanoparticle labels. This time, the course of the DNA can be followed over the electrodes, but again hampered by low resolution at the metal surfaces. What is the mechanism for the observed differences in the contrast of DNA on microelectrodes? Gold films are able to subdue fluorescent signals due to quenching processes. Assuming similar gold structures (about 100 nm thick) in all experiments, it alone cannot explain the variations. Another factor is the background fluorescence signal, which is probably influenced by the surface modification protocol and could reflect variations in this step. This background signal is documented in Figure 2e. The black thread-like structures are probably induced by DNA fibers that were adsorbed on (and thereby shadowing) this location in the step of DNA labeling. The background also binds fluorescence dyes, with the exception of the regions protected by the adsorbed molecules. These molecules were later removed, probably in washing steps involving shear stress. Maybe the thickness of the DNA fibers also plays a role, because quenching is a highly localized process, and thicker fibers would be less influenced in the parts more distant to the electrodes. This theory is supported by images such as the one shown in Figure 2f, where fibers cross the electrode structures, and the fluorescence signal in these points is apparently correlated with the fiber thickness. So thicker fibers are still visible in these areas, but thinner fibers give no detectable signal. We explain the binding of the DNA along the electrodes by a preferred adsorption of the previously aligned DNA. The alignment is probably due to occasional terminal binding of DNA17 and the induced fluid flow by the receding droplet meniscus. When the DNA has a higher affinity to the gold electrodes, its primary binding will occur there and therefore also the alignment and later the adsorption along the molecule. This process is probably supported by differences in hydrophobicity between the electrode surface and the silanized substrate surface. Further experiments will show if a passivation of the substrate, by functionalization in order to minimize DNA binding there, results in a higher ratio of DNA binding on the electrodes versus DNA binding on the substrate. We could demonstrate a new technique for highly paralleled, precise positioning of DNA in microelectrode gaps
Nano Lett., Vol. 4, No. 4, 2004
based on guided self-organization by the DNA. Different microscopic tools contributed to the elucidation of the final structure. The technique combines the experiences and reliability of the droplet spreading method with a prealignment to microstructured electrode arrays in order to achieve a molecular positioning in a technical environment as a key step for any use of molecular nanotechnology today. Acknowledgment. We thank A. Csaki for valuable discussions and the establishment of DNA-nanoparticle techniques in our group, A. Steinbru¨ck and A. Wolff for Figure 2e, F. Jahn for assistance with SEM characterization, D. Born for design and provision of the microstructured substrates, and J. M. Ko¨hler for his contribution to this research area at the IPHT. This work was supported by the Volkswagen Foundation (Priority Area: Physics, Chemistry and Biology with Single Molecules). Supporting Information Available: Scanning force micrograph (height mode) of nanoparticle-labled DNA (as shown in Figure 4c by SEM) revealing the inhomogenity of particle distribution along the DNA that is highlighted by arrowheads. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Seeman, N. C. Trends Biotechnol. 1999, 17, 437-443. (2) Hu, J.; Zhang, Y.; Gao, H.; Li, M.; Hartmann, U. Nano Lett. 2002, 2, 55-57. (3) Braun, E.; Eichen, Y.; Sivan, U.; Ben-Yoseph, G. Nature 1998, 391, 775-778. (4) Keren, K.; Krueger, M.; Gilad, R.; Ben-Yoseph, G.; Sivan, U.; Braun, E. Science 2002, 297, 72-75. (5) Mertig, M.; Colomb Ciachi, L.; Seidel, R.; Pompe, W.; De Vita, A. Nano Lett. 2002, 2, 841-844. (6) Monson, C. F.: Woolley, A. T. Nano Lett. 2003, 3, 359-363. (7) Patolsky, F.; Weizmann, Y.; Lioubashevski, O.; Willner, I. Angew. Chem., Int. Ed. Eng. 2002, 41, 2323-2327. (8) Yan, H.; Park, S. H.; Finkelstein, G.; Reif, J. H.; LaBean, T. H. Science 2003, 301, 1882-1884. (9) Keren, K.; Berman, R. S.; Buchstab, E.; Sivan, U.; Braun, E. Science 2003, 302, 1380-1382. (10) Heng, H. Q. H.; Squire, J.; Tsui, L.-C. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 9509-9513. (11) Bensimon, D.; Simon, A. J.; Croquette, V.; Bensimon, A. Phys. ReV. Lett. 1995, 74, 4754-4757. (12) Maubach, G.; Csaki, A.; Seidel, R.; Mertig, M.; Pompe, W.; Born, D.; Fritzsche, W. Nanotechnology 2003, 14, 546-550. (13) Vallant, T.; Brunner, H.; Mayer, U.; Hoffmann, H.; Leitner, T.; Resch, R.; Friedbacher, J. G. J. Phys. Chem. B 1998, 102, 7190-7197. (14) Fang, Y.; Spisz, T. S.; Wiltshire, T.; D’Costa, N. P.; Bankman, I. N.; Reewes, R. H.; Hoh, J. H. Anal. Chem. 1998, 70, 2123-2129. (15) Jing, J.; Reed, J.; Huang, J.; Hu, X.; Clarke, V.; Edington, J.; Housman, D.; Anantharaman, T. S.; Huff, E. J.; Mishra, B.; Porter, B.; Shenker, A.; Wolfson, E.; Hiort, C.; Kantor, R.; Aston, C.; Schwartz, D. C. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 8046-8051. (16) Warner, M. G.; Hutchison, J. E. Nature Materials 2003, 2, 272277. (17) Allemand, J. F.; Bensimon, D.; Jullien, L.; Bensimon, A.; Croquette, V. Biophys. J. 1997, 73, 2064-2070.
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