Patterning and Characterization of Surfaces with Organic and

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Anal. Chem. 2000, 72, 3431-3435

Patterning and Characterization of Surfaces with Organic and Biological Molecules by the Scanning Electrochemical Microscope Iva Turyan,‡ Tomokazu Matsue,† and Daniel Mandler*,‡

Department of Molecular Chemistry and Engineering, Faculty of Engineering, Tohoku University, Sendai 980-77, Japan, and Department of Inorganic and Analytical Chemistry, The Hebrew University of Jerusalem, Jerusalem 91904, Israel

A novel approach for micropatterning of surfaces with organic and biological microstructures using the scanning electrochemical microscope (SECM) is described. The approach is based on the introduction of the spatial resolution by local deposition of gold particles followed by monolayer formation and functionalization. Specifically, gold patterns were deposited locally on silicon wafers with the SECM as a result of the controlled anodic dissolution of a gold microelectrode. The gold patterns were further used as microsubstrates for assembling cystamine monolayers to which either fluoresceine isothiocyanate (FIT) or glucose oxidase (GOD) were covalently attached. Characterization of the organic monolayers, as well as the biological activity of the enzyme patterns, was carried out by fluorescence microscopy and the SECM, respectively. The development of techniques for micropatterning biomolecules on solid substrates has attracted increasing consideration as a means of assembling miniaturized and integrated chemical and biosensors, such as DNA sequencing microsensors1 and multianalyte immunosensing chips.2 Patterning has been driven by different approaches, including photochemical methods, such as photolithography,1c,2a,3-7 photoimmobilization,8-10 and laser * Corresponding author. Tel: 972-2-658-5831. Fax: 972-2-658-5319. E-mail: [email protected] † Tohoku University. ‡ The Hebrew University of Jerusalem. (1) (a) Fodor, S. P. A.; Read, J. L.; Pirrung, M. C.; Stryer, L.; Lu, A. T.; Solas, D. Science (Washington, D.C.) 1991, 251, 767-73. (b) Chee, M.; Yang, R.; Hubbell, E.; Berno, A.; Huang, X. C.; Stern, D.; Winkler, J.; Lockhart, D. J.; Morris, M. S.; Fodor, S. P. A. Science (Washington, D.C.) 1996, 274, 61014. (c) Pritchard, D. J.; Morgan, H.; Cooper, J. M. Angew. Chem. Int. Ed. Engl. 1995, 34, 91-3. (2) (a) Sundberg, S. A.; Barrett, R. W.; Pirrung, M.; Lu, A. L.; Kiangsoontra, B.; Holmes, C. P. J. Am. Chem. Soc. 1995, 117, 12050-2057. (b) Shiku, H.; Matsue, T.; Uchida, I. Anal. Chem. 1996, 68, 1276-278. (c) Shiku, H.; Hara, Y.; Matsue, T.; Uchida, I.; Yamauchi, T. J. Electroanal. Chem., in press. (3) (a) Hanazato, Y.; Nakano, M.; Maeda, M.; Shiono, S. Anal. Chim. Acta 1987, 193, 87-96. (b) Vopel, T.; Ladde, A.; Muller, H. Anal. Chim. Acta 1991, 251, 117-20. (c) Bhatia, S. K.; Hickman, J. J.; Ligler, F. S. J. Am. Chem. Soc. 1992, 114, 4432-433. (d) Nicolau, D. V.; Suzuki, H.; Mashiko, S.; Taguchi, T.; Yoshikawa, S. Biophys. J. 1999, 77, 1126-134. (4) Chan, K. C.; Kim, T.; Shoer, J. K.; Crooks, R. M. J. Am. Chem. Soc. 1995, 117, 5875-876. (5) Sundarababu, G.; Gao, H.; Sigrist, H. Photochem. Photobiol. 1995, 61, 54044. (6) Flounders, A. W.; Brandon, D. L.; Bates, A. H. Biosens. Bioelectron. 1997, 12, 447-56. (7) Dontha, N.; Nowall, W. B.; Kuhr, W. G. Anal. Chem. 1997, 69, 2619-625. 10.1021/ac000046a CCC: $19.00 Published on Web 07/06/2000

© 2000 American Chemical Society

ablation,11,12 by electrochemical means, e.g., through the electrochemical patterning of self-assembled monolayers (SAMs),13 using mechanical approaches, such as microcontact printing,14,15 and by scanning probe microscopy.16-31 Photolithography, which is the dominant patterning approach in fabricating solid-state devices, is usually limited due to the vacuum, organic solvents, and strong acids and bases that are employed and which are harmful to the biological substances. (8) Morgen, H.; Pritchard, D. J.; Cooper, J. M. Biosens. Bioelectron. 1995, 10, 841-46. (9) Hengsakul, M.; Cass, A. E. G. Bioconjugate Chem. 1996, 7, 249-54. (10) Delamarche, E.; Sundarababu, G.; Biebuyck, H.; Michel, B.; Gerber, C.; Sigrist, H.; Wolf, H.; Ringsdorf, H.; Xanthopoulos, N.; Mathieu, H. J. Langmuir 1996, 12, 1997-2006. (11) Vaidya, R.; Tender, L. M.; Bradley, G.; O’Brien, M. J.; Cone, M.; Lopez, G. P. Biotechnol. Prog. 1998, 14, 371-77. (12) (a) Schwarz, A.; Rossier, J. S.; Roulet, E.; Mermod, N.; Roberts, M. A.; Girault, H. H. Langmuir 1998, 14, 5526-531. (b) Rossier, J. S.; Schwarz, A.; Reymond, F.; Ferrigno, R.; Bianchi, F.; Girault, H. H. Electrophoresis 1999, 20, 727-31. (13) Tender, L. M.; Worley, R. L.; Fan, H.; Lopez, G. P. Langmuir 1996, 12, 5515-518. (14) Bernard, A.; Delamaarche, E.; Schmid, H.; Michel, B.; Bosshard, H. R.; Biebuyck, H. Langmuir 1998, 14, 2225-229. (15) James, C. D.; Davis, R. C.; Kam, L.; Craighead, H. G.; Isaacson, M.; Turner, J. N.; Shain, W. Langmuir 1998, 14, 741-44. (16) Gimzewski, J. K.; Jung, T. A.; Cuberes, M. T.; Schlittler, R. R. Surf. Sci. 1997, 386, 101-14. (17) Hong, S. H.; Zhu, J.; Mirkin, C. A. Science (Washington, D.C.) 1999, 286, 523-25. (18) (a) Sugimura, H.; Uchida, T.; Shimo, N.; Kitamura, N.; Masuhara, H., Ultramicroscopy 1992, 42-44, 468-74. (b) Sugimura, H.; Uchida, T.; Shimo, N.; Kitamura, N.; Shimo, N.; Masuhara, H. J. Electroanal. Chem. 1993, 361, 57-63. (19) Wittstock, G.; Hesse, R.; Schuhmann, W. Electroanalysis 1997, 9, 746-50. (20) Wittstock, G.; Schuhmann, W. Anal. Chem. 1997, 69, 5059-66. (21) Shiku, H.; Hara, Y.; Takeda, T.; Matsue, T.; Uchida, I. Solid Liquid Electrochem. Interfaces; ACS Symposium Series 656; Jerkiewicz, G., Soriaga, M. P., Uosaki, K., Wieckowski, A., Eds.; American Chemical Society: Washington, DC, 1997; p 202. (22) Shiku, H.; Takeda, T.; Yamada, H.; Matsue, T.; Uchida, I. Anal. Chem. 1995, 67, 312-17. (23) Shiku, H.; Uchida, I.; Matsue, T. Langmuir 1997, 13, 7239-244. (24) Nowall, W. B.; Wipf, D. O.; Kuhr, W. G. Anal. Chem. 1998, 70, 2601-606. (25) Mandler, D. In Scanning Electrochemical Microscopy; Bard, A. J., Mirkin, M. V., Eds.; Marcel Dekker: in press. (26) Wang, J.; Wu, L.-H.; Li, R. L. J. Electroanal. Chem. 1989, 272, 285-92. (27) Pierce, D. T.; Unwin, P. R.; Bard, A. J. Anal. Chem. 1992, 64, 1795-804. (28) Pierce, D. T.; Bard, A. J. Anal. Chem. 1993, 65, 3598-604. (29) Wittstock, G.; Yu, K.; Halsall, B.; Ridgway, T. H.; Heineman, W. R. Anal. Chem. 1995, 67, 3578-582. (30) Schiku, H.; Matsue, T.; Uchida, I. Anal. Chem. 1996, 68, 1276-278. (31) Kranz, C.; Wittstock, G.; Wohlschlager, H.; Schumann, W. Electrochim. Acta, 1997, 42, 3105-111.

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Substantial efforts in micropatterning surfaces with biological functionalities have involved self-assembled monolayers (SAMs).10,32-34 SAMs comprise arrays of molecular thickness to which biomolecules can easily be attached. Hence, many scientists envision that patterning SAMs could eventually pave the route for constructing arrays with two molecular dimensions. Consequently, methods aiming at either removing the SAMs locally or changing the hydrophobicity/hydrophilicity or charge of the monolayers have been reported and used for locally attaching biomolecules. To increase the lateral resolution of such local modifications, the scanning probe techniques have been invoked, which allow nanometer-scale surface manipulations. Among these techniques, the scanning electrochemical microscope (SECM) has been particularly promising for microfabrication18-25 and detection of immobilized biomolecules.26-31 Two different modes of operation, i.e., the direct18-20 and feedback modes,21-24 have been used to create and image microstructures of biologically active surfaces with the SECM. In the direct mode, the UME acts as the auxiliary electrode approaching a biased, conducting substrate. The first study, which utilized the direct mode, was reported by Masuhara et al.18 Fluorescent patterns of Rhodamine 6G in an ionically conducting polymer were formed as a result of the local decomposition of a quencher, methyl viologen, which was incorporated inside the polymer. More recent efforts to pattern surfaces with organic and biomolecules were carried out by Wittstock and Schuhmann.19,20 The localized desorption of alkanethiol monolayers was induced by the direct mode.19 This was exploited further20 for the formation of enzymatic patterns of glucose oxidase that was covalently attached to cystamine. The latter was adsorbed in the exposed gold patterns. The enzymatic activity was imaged using the generation-collection mode with an UME that acted as an amperometric probe and detected the oxidation of hydrogen peroxide. Exploitation of the SECM in the feedback mode for microfabrication of organic and biological structures has been reported only recently and basically lags behind the direct mode. The reason for this is that most organic and biomolecules do not exhibit simple-one-electron redox behavior, which prevents their use as mediators. The first study was reported by Matsue and co-workers22 and aimed at creating micropatterns of diaphorase on a glass substrate. The approach was based on the local electrogeneration of bromine that deactivated immobilized enzyme molecules forming nonreactive patterns. They also used the SECM as a means of characterizing enzyme and antigen/antibody patterns;22 however, patterning was carried out by micropipetting. Recently, Matsue presented a different approach,23 in which silane monolayers were modified locally by reactive species, i.e., hydroxyl radicals. The radicals were generated upon electrochemically reducing Fe3+ at the microelectrode in the presence of hydrogen (32) For example: (a) Kumar, A.; Biebuyck, H. A.; Whitesides, G. M. Langmuir 1994, 10, 1498. (b) Kumar, A.; Whitesides, G. M. Appl. Phys. Lett. 1993, 63, 2002-2004. (c) Kumar, A.; Abbott, N. L.; Kim, E.; Biebuyck, H. A.; Whitesides, G. M. Acc. Chem. Res. 1995, 28, 219-26. (33) For example: (a) Mooney, J. F.; Hunt, A. J.; Mclntosh, J. R.; Liberko, C. A.; Walba, D. M.; Rogers, C. T. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 122872291. (b) Herbert, C. B.; McLernon, T. L.; Hypolite, C. L.; Adams, D. N.; Pikus, L.; Huang, C. C.; Fields, G. B.; Letourneau, P. C.; Disfefano, M. D.; Hu, W. S. Chem. Biol. 1997, 4, 731-37. (34) Lopez, G. P.; Biebuyck, H. A.; Harter, R.; Kumar, A.; Whitesides, G. M. J. Am. Chem. Soc. 1993, 115, 10774-781.

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peroxide.23 The enzyme patterns were formed taking advantage of the differences in physical and chemical properties of the functional groups of the monolayers being exposed to the hydroxyl radicals. Another recent example of micropatterning surfaces with biological molecules by the feedback mode was demonstrated by Wipf24 and involved the local attachment or desorption of biotin onto a glassy carbon surface. Reviewing the literature reveals that, in most cases, the formation of patterns has been accomplished by locally changing the organic monolayers; in many cases, deactivated patterns have been formed as well. Here we describe a simple, yet versatile, approach for anchoring organic and biological substances with micrometer resolution. The essence of the approach involves the local deposition of gold particles by the SECM.35 The particles are subsequently used for attaching the molecules via standard surface chemistry. The concept is demonstrated by attaching a fluorescent dye, i.e., fluorescein, and the enzyme glucose oxidase. The activity of the latter was approved after attachment using the SECM in the generation-collection mode. EXPERIMENTAL SECTION All electrochemical measurements were performed with a BAS100B electrochemical analyzer. The basic apparatus used for the SECM experiments has been described previously.36 A bipotentiostat (EI-400 model, Ensman Instrumentation) was used to control the potential of both the microelectrode and the silicon substrate. Optical micrographs were obtained with an Axioskop 2 microscope (Zeiss). The fluorescence images were taken with the same microscope by collecting the fluorescence through a 590-nm cut off filter. All aqueous solutions were prepared from deionized water (Milli-Q, Millipore Corp.). Silicon wafers were a generous gift from Wacker Siltronic AG. The wafers were n-Si(111) (phosphorus doped) with resistivity of 1.0-10.0 ohm‚cm. A GaIn alloy (Alfa) was used for making an ohmic contact to the Si surface. The alloy was covered with an epoxy (Torr Seal, Variant). All other chemicals were purchased from Aldrich. Twenty-five-micrometer-diameter Au and Pt ultramicroelectrodes (UMEs) were made from the respective wires (Goodfellow, Cambridge, UK) that were heat-sealed in glass capillaries under vacuum as described previously.35 Smaller microelectrodes were prepared by etching the Pt wire prior to its sealing.22 Before each measurement, the electrodes were polished with emery paper (240 and 600 grits, Buehler) followed by diamond paste (from 6- to 1-µm) on a nylon polishing cloth (Buehler) and 0.05-µm alumina slurry on a microcloth polishing pad using a homemade motordriven polishing wheel. Finally, the UME was electrochemically cycled between -0.5 and 1.6 V vs Ag/AgCl at 100 mV‚s-1 in 0.1 M H2SO4 solution until a typical steady-state voltammogram of activated Pt or Au was obtained. An Ag/AgCl electrode in saturated KCl and a Pt wire were used as reference and counter electrodes, respectively. Therefore, all potentials are quoted vs this reference electrode. Prior to each experiment, the Si slide was dipped in a 49% HF solution for a few seconds to remove the oxide layer. Gold patterns were deposited upon leaving an Au UME (1.0 V) close above a (35) Meltzer, S.; Mandler, D. J. Electrochem. Soc. 1995, 142, L82-L84. (36) Shohat, I.; Mandler, D. J. Electrochem. Soc. 1994, 141, 995-99.

silicon surface (-0.1 V) in 0.01-0.05 M HCl solution for 1-5 min. Then the silicon wafer with the gold patterns was immersed in a 10 mM cystamine and 0.1 M acetate buffer (pH 5.5) solution for 30 min under ambient conditions. The fluorescent dye was attached by dipping the cystamine-modified surface into a 2 mM fluorescein isothiocyanate and 0.1 M phosphate buffer (pH 7.2) solution for 1 h. The surface was then washed with distilled water to remove any loosely bound species. Glucose oxidase (GOD) was covalently attached to the cystamine monolayer by immersing the cystamine-terminated surface into a 0.1 M phosphate buffer (pH 7.4) consisting of 5 × 10-6 M GOD and 2.5% glutardialdehyde for 4 h. The precise position of the Pt UME (after deposition of Au patterns) was determined by comparing the experimental and theoretical positive feedback currents detected over the gold patterns. The UME was biased at -0.1 V in a solution of 2 mM Fe(CN)63- and 0.1 M Na2SO4, and the substrate was biased at +0.5 V. Then, the Pt UME was withdrawn and the feedback current was recorded over an unbiased gold pattern. The position of the Pt UME was kept constant while modifying the gold patterns. The SECM image of the gold dots was obtained by scanning a Pt UME (2.8-µm-diameter, 0.5 V vs Ag/AgCl) laterally with a speed of 9.8 µm‚s-1 in a solution of 4 mM of Fe(CN)64and 0.1 M KCl. To verify the formation of the enzymatic pattern, the Pt UME (25-µm-diameter) was scanned across the enzyme pattern in an air-saturated solution that consisted also of 50 mM glucose and 0.1 M phosphate buffer (pH 7.4) with a speed of 5 µm‚s-1. The potential of the UME was set at -0.4 V in order to ensure the diffusion-controlled reduction of dissolved hydrogen peroxide. This measurement was repeated after 2300 units of catalase (soluble bovine liver catalase, Sigma) had been added into the solution. RESULTS AND DISCUSSION The covalent attachment of organic and biomolecules, e.g., enzymes, onto surfaces is well established and often takes advantage of the chemical activity of the amino functional group.37 Therefore, this step is usually preceded by the formation of an amino-terminated monolayer through the adsorption of cystamine or cysteamine on gold surfaces. We reported on the formation of micropatterns of gold,35 the so-called microwriting, by the controlled anodic dissolution of a gold microelectrode. The essence of the approach involves the continuous electrochemical dissolution of a gold microelectrode in the presence of complexation ions, such as chloride (Scheme 1). This approach offers two significant advantages in terms of local metal deposition. The resolution of the deposits is determined by the size of the conducting (metallic) part of the microelectrode, regardless of the insulating sheath. Second, the complexing ions, e.g., Cl-, which are regenerated upon metal deposition cause the anodic current (at the microelectrode) to increase as the distance to the surface decreases. In other words, a positive feedback current is observed while the microelectrode approaches a conducting surface, providing the means of controlling the microelectrode-surface distance. Figure 1A shows an optical (reflected light) micrograph of four gold dots, which were formed upon leaving a 25-µm-diameter gold (37) Methods in Enzymology; Mosbach, K., Colowick, S. P., Kaplan, N. O., Eds.; Academic Press: New York, 1976; Vol. 44.

Scheme 1: Schematic Representation of the Approach for Attaching Organic and Biological Molecules onto Surfaces Using the SECM

microelectrode biased at 1.0 V for 5 min above a n-Si wafer (-0.1 V) in 0.1 M HCl. The rate of gold dissolution and deposition is governed by the potential of the microelectrode, the nature of the halide ions in the electrolyte solution, and the potential of the substrate. The effect of these parameters has been studied by us in detail and will be reported elsewhere.38 In essence, we found that the dissolution of gold can be very fast, and therefore, the deposit depends primarily on the potential applied to the tip, the concentration of the complexing ions, and the pH of the solution. The deposition of gold proceeds through the formation of nanocrystals that grow and eventually merge into a continuous structure as a result of leaving the microelectrode constantly close above the substrate. The surface density of the nanoparticles can easily be controlled by the duration of the potential pulse that is applied to the tip. Figure 1B shows a three-dimensional image obtained by the SECM in the feedback mode as a result of scanning the gold dots previously formed. The solution consisted of 4 mM hexacyanoferrate(II) in 0.1 M KCl, which was electrochemically oxidized at the Pt tip (2.8-µm-diameter, 0.5 V) under diffusion control while the Si was not attached to an external power source. Evidently, the contrast obtained in Figure 1B is due to significant differences in conductivity between the gold dots and the silicon wafer. The latter is likely to be covered with a native oxide layer. It is worth mentioning that a positive feedback current is detected upon scanning above the dots with a much larger microelectrode, i.e., of the order of 25 µm in diameter. This clearly indicates that the dots must maintain an ohmic contact with the underlying silicon and that the oxide silicon layer does not completely block hexacyanoferrate(II) oxidation, which is the source of the feedback current in the case of an unbiased substrate.39 The dots could easily be dissolved upon applying a sufficiently positive potential (38) Ammann, E.; Mandler, D., in preparation.

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Figure 1. (A) Optical reflected micrograph of gold dots deposited by the SECM. The gold was deposited on a Si wafer (biased at -0.1 V) as a result of leaving a Au UME (25-µm-diameter, 1.0 V) above the surface for 5 min in 0.05 M HCl. (B) Current vs location image obtained by scanning a Pt UME (2.8-µm-diameter, 0.5 V) with a speed of 9.8 µm‚s-1 over gold dots (deposited on a Si wafer) in a solution of 4 mM Fe(CN)64- and 0.1 M KCl. (C) Optical fluorescence micrograph of the gold dots after immersing the surface into a solution of 10 mM cystamine (0.1 M acetate buffer, pH 5.5) for 30 min followed by FIT modification (2 mM FIT in 0.1 M phosphate buffer, pH 7.2) during 1 h. Images were acquired with a 590-nm cutoff filter.

to the silicon wafer in the presence of halide solutions. Clearly, the size of the dots obtained by the SECM is comparable to that obtained by optical microscopy. The enhancement of the current which is observed between the dots is due to some metal deposition that has taken place while moving the microelectrode from one location to the othersalthough the gold microelectrode was withdrawn before being brought to the next location. In the second step (Scheme 1), cystamine was adsorbed onto the gold patterns. It should be noted, at this stage, that the underlying substrate, i.e., silicon, was not incidentally chosen. While thiols are strongly adsorbed on gold as well as other metallic and inorganic surfaces, they do not chemisorb on Si. Hence, indium tin oxide could not be used as an alternative substrate as a few studies reported on the adsorption of thiols on this surface.40 The localization of the self-assembled monolayer on the surfaces was monitored by fluorescence microscopy after the (39) Bard, A. J.; Fan, F.-R. F.; Mirkin, M. V. In Electroanalytical Chemistry; Bard, A. J., Ed.; Marcel Dekker: New York, 1994; Vol. 18, 243-373.

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sample was tagged with fluoresceine isothiocyanate (FIT, Scheme 1). In other words, the cystamine-modified gold dots sample was dipped in a FIT solution for 1 h, following which it was washed with distilled water to remove any loosely bound species. Figure 1C shows a fluorescence optical micrograph of the modified sample. The fluorescence of the attached fluoresceine can clearly and only be seen on the gold micropatterns. The images were obtained by collecting the fluorescence through a filter, which cuts off the FIT absorption band (590 nm). Blank experiments showed that no fluorescence was detected with either bare silicon or gold samples that were treated with FIT in the absence of the cystamine monolayer. This clearly indicates that FIT reacts with the amino groups, hence, confirming the presence of an aminoterminated monolayer on the gold microstructures only. Interestingly, the size of the patterns obtained by fluorescence microscopy is somewhat larger than that measured by reflected light microscopy as well as that measured by the SECM. This suggests that some fluoresceine is attached to the periphery of the patterns, indicating the presence of very small gold particles that can only be detected by the fluorescence of the attached dye. The next step comprised the attachment of the enzyme instead of the fluorescent tag. Such cystamine monolayers have widely been used for the covalent attachment of different enzymes and, in particular, glucose oxidase (GOD).20 The immobilization of GOD onto the locally assembled cystamine monolayer was carried out in a buffer (pH 7.4) consisting of GOD and glutardialdehyde. The concentration of the enzyme, the pH of the solution, and the sequence of the addition of the various reagents during immobilization of the enzyme strongly affect the enzyme activity. It has been shown that best results in terms of GOD activity have been achieved by adding the enzyme and the cross-linking agent simultaneously.41 Unbound enzyme was rinsed off with water and phosphate buffer. One of the advantages of the SECM is its capability of monitoring local enzymatic activity.19-31 Specifically, the SECM was used in the generation-collection mode to verify the formation of an enzyme pattern following the approach of Schuhmann.20 Initially, the microelectrode was positioned above the gold patterns prior to their modification with the enzyme. This is easily accomplished through the positive feedback current observed over the gold micropatterns with hexacyanoferrate as a mediator (Figure 1B). Then, the gold patterns were modified following the above procedure, while the position of the tip was maintained constant. Finally, the Pt microelectrode was scanned across the enzyme pattern (with a speed of 5 µm‚s-1) in an air-saturated solution after 50 mM glucose (phosphate buffer, pH 7.4) had been added. The potential of the microelectrode was held at -0.4 V, which was sufficiently negative to reduce any hydrogen peroxide under diffusion-controlled conditions. Since glucose is not capable of reducing oxygen in the absence of GOD, no current is detected at the Pt tip in the bulk of the solution. At the same time, a distinct faradaic current can be detected while crossing the gold-modified pattern (Figure 2A). Evidently, this current must be due to the formation of H2O2 in an enzymatic-catalyzed reaction. In other (40) (a) Kondo, T.; Takechi, S. Y.; Uosaki, K. J. Electroanal. Chem. 1995, 381, 203-209. (b) Sato, Y.; Uosaki, K. Denki Kagaku oyobi Kogyo Butsuri Kagaku 1994, 62, 1269-275. (41) Bouin, J. C.; Atallah, M. T.; Hultin, H. O. Biotechnol. Bioeng. 1976, 18, 179-87.

Figure 2. (A) The steady-state current of a 25-µm-diameter Au microelectrode recorded in an air-saturated solution of 50 mM glucose in 0.1 M phosphate buffer (pH 7.4) while scanning across a gold dot, which was previously modified with GOD. The potential of the Au microelectrode was 0.4 V while the Si wafer was not biased. (B) Same as A, after the addition of catalase. (C) Blank experiment. Same as A, however, without glucose.

words, the Pt microelectrode is used as an amperometric sensor for the detection of H2O2 and, therefore, for imaging the enzyme activity. The main advantage of this approach, as suggested by Schuhmann,20 is the elimination of any contributions to the current at the tip due to surface conductivity. That is, imaging of the enzyme activity cannot be accomplished by using a simple oneelectron redox couple, such as hexacyanoferrate, since the increase of the current, in this case, would originate from the electronic conductivity of the gold patterns. On the other hand, it should be noted that the generation-collection mode results in a substantially larger activity area than the “true” area occupied by the bound enzyme, as is evident by the cross section of the enhanced current in Figure 2A. Obviously, the image obtained via the generation-collection mode is directly related to the diffusion layer of the collected species. This increase can be solved by either scanning closer to the surface or upon using the “chemical lens” mode reported by Heinze42,43 and later by Mao.44 Indeed, Figure 2B shows the current that was recorded while scanning the same spot (maintaining the tip at the same distance above the surface) after adding catalase to the solution.20 Two distinct differences can be observed: a decrease of the background current and a narrowing of the enhanced current above the spot. These effects are clearly due to the decomposition of the hydrogen peroxide by the catalase, as has been demonstrated (42) Heβ, C.; Borgwarth, K.; Ricken, C.; Ebling, D. G.; Heinze, J. Electrochim. Acta 1997, 42, 3065. (43) Borgwarth, K.; Heinze, J. J. Electrochem. Soc. 1999, 146, 3285-289. (44) Zu, Y. B.; Xie, L.; Mao, B. W.; Tian, Z. W. Electrochim. Acta 1998, 43, 1683690.

previously by Wittstock and Schuhmann20 and which exemplifies the “chemical lens” effect. Nonetheless, it should be noted that the resolution of the signal is not enhanced since the signal also decreases. Finally, in the absence of glucose (Figure 2C), the activity of the bound enzyme is not detectable. CONCLUSIONS A new and simple approach for the formation of stable micrometer-size organic and biological patterns using the scanning electrochemical microscope is demonstrated. This approach offers some distinct advantages. The high surface area of the gold particles allows the formation of high-density functionalized monolayers that can easily be detected by different microscopic methods. In addition, since much smaller microelectrodes can be constructed, the size of the patterns can be significantly decreased. Finally, and probably most importantly, the fact that the gold patterns are electrically connected with the underlying silicon can easily be applied for assembling organic or biological microelectrode arrays. Improving the local metal deposition using the SECM as well as applying it as a means of assembling biochemical arrays are the subjects of our current activity. ACKNOWLEDGMENT The Israeli Ministry of Science supported this work within the frame of cooperation with Japan (Grant 8931298). Received for review January 10, 2000. Accepted April 27, 2000. AC000046A

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