Chitosan-Mediated and Spatially Selective Electrodeposition of

ACS Applied Materials & Interfaces 0 (proofing),. Abstract | Full Text ... and Gregory F. Payne. Langmuir 2007 23 (1), 286-291 ... Electroanalysis 201...
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Chitosan-Mediated and Spatially Selective Electrodeposition of Nanoscale Particles Li-Qun Wu,† Kyuyong Lee,‡ Xiang Wang,§ Douglas S. English,§ Wolfgang Losert,‡ and Gregory F. Payne*,†,| Center for Biosystems Research, University of Maryland Biotechnology Institute, 5115 Plant Sciences Building, College Park, Maryland 20742, Department of Physics, Institute for Physical Sciences and Technology, and Institute for Research in Electronics and Applied Physics, and Department of Chemistry and Biochemistry, University of Maryland, College Park, Maryland 20742, and Department of Chemical and Biochemical Engineering, University of Maryland, Baltimore County, 1000 Hilltop Circle, Baltimore, Maryland 21250 Received October 19, 2004. In Final Form: December 21, 2004 Nanoscale particles offer a variety of interesting properties, and there is growing interest in their assembly into higher ordered structures. We report that the pH-responsive aminopolysaccharide chitosan can mediate the electrodeposition of model nanoparticles. Chitosan is known to electrodeposit at the cathode surface in response to a high localized pH. To demonstrate that chitosan can mediate nanoparticle deposition, we suspended fluorescently labeled latex nanoparticles (100 nm diameter spheres) in a chitosan solution (1%) and performed electrodeposition (0.05 mA/cm2 for several minutes). Results demonstrate that chitosan is required for nanoparticle electrodeposition; chitosan confers spatial selectivity to electrodeposition; and nanoparticles distribute throughout the electrodeposited chitosan film. Additionally, we observed that the deposited films reversibly swell upon rehydration. This work indicates that chitosan provides a simple means to assemble nanoparticles at addressable locations and provides further evidence that stimuliresponsive biological materials may facilitate fabrication at the microscale.

Introduction Particles that span size ranges of 1-100 nm have attracted considerable recent attention for a variety of reasons. Metal and semiconductor nanocrystals have tunable properties (e.g., optical, electronic, and magnetic) that depend on particle size, interparticle spacing, and higher order structure.1-4 Various biological and biomimetic systems self-assemble at the nanoscale (e.g., vesicles,5,6 liposomes,7 polymersomes,8 and virus particles9) enabling them to serve as nano-compartments10,11 or templates for further assembly.12-14 Nanoscale polymeric * To whom correspondence should be addressed. E-mail: [email protected]. † University of Maryland Biotechnology Institute. ‡ Department of Physics, Institute for Physical Sciences and Technology, and Institute for Research in Electronics and Applied Physics, University of Maryland. § Department of Chemistry and Biochemistry, University of Maryland. | University of Maryland, Baltimore County. (1) Alivisatos, A. P.; Barbara, P. F.; Castleman, A. W.; Chang, J.; Dixon, D. A.; Klein, M. L.; McLendon, G. L.; Miller, J. S.; Ratner, M. A.; Rossky, P. J.; Stupp, S. I.; Thompson, M. E. Adv. Mater. 1998, 10, 1297-1336. (2) Schmid, G.; Chi, L. F. Adv. Mater. 1998, 10, 515-526. (3) Murray, C. B.; Kagan, C. R.; Bawendi, M. G. Annu. Rev. Mater. Sci. 2000, 30, 545-610. (4) Pileni, M. P. J. Phys. Chem. B 2001, 105, 3358-3371. (5) Michel, M.; Vautier, D.; Voegel, J. C.; Schaaf, P.; Ball, V. Langmuir 2004, 20, 4835-4839. (6) Katagiri, K.; Hamasaki, R.; Ariga, K.; Kikuchi, J. J. Am. Chem. Soc. 2002, 124, 7892-7893. (7) Katagiri, K.; Hamasaki, R.; Ariga, K.; Kikuchi, J. Langmuir 2002, 18, 6709-6711. (8) Napoli, A.; Boerakker, M. J.; Tirelli, N.; Nolte, R. J. M.; Sommerdijk, N. A. J. M.; Hubbell, J. A. Langmuir 2004, 20, 34873491. (9) Strable, E.; Johnson, J. E.; Finn, M. G. Nano Lett. 2004, 4, 13851389. (10) Douglas, T.; Young, M. Nature 1998, 393, 152-155. (11) Jang, H.; Pell, L. E.; Korgel, B. A.; English, D. S. J. Photochem. Photobiol., A 2003, 158, 111-117.

systems (e.g., dendrimers15 and nanofibers16,17) have been combined with other components to create materials with multiple properties (e.g., biocatalytic18 or biocatalytic plus magnetic19,20). Often, applications of nanoparticles require their assembly into higher ordered structures, and several approaches have been used to assemble nanoparticles in two and three dimensions. In some cases, nanoparticle organization has been guided by interparticle interactions that occur during evaporation,21,22 precipitation,23 or film casting.24 In other cases, nanoparticles have been assembled using polymermediated layer-by-layer methods.25-27 In still other cases, (12) Lee, S. W.; Mao, C.; Flynn, C. E.; Belcher, A. M. Science 2002, 296, 892-895. (13) Mao, C.; Solis, D. J.; Reiss, B. D.; Kottmann, S. T.; Sweeney, R. Y.; Hayhurst, A.; Georgiou, G.; Iverson, B.; Belcher, A. M. Science 2004, 303, 213-217. (14) Mao, C.; Flynn, C. E.; Hayhurst, A.; Sweeney, R.; Qi, J.; Georgiou, G.; Iverson, B.; Belcher, A. M. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 6946-6951. (15) Crooks, R. M.; Zhao, M. Q.; Sun, L.; Chechik, V.; Yeung, L. K. Acc. Chem. Res. 2001, 34, 181-190. (16) Jia, H. F.; Zhu, G. Y.; Vugrinovich, B.; Kataphinan, W.; Reneker, D. H.; Wang, P. Biotechnol. Prog. 2002, 18, 1027-1032. (17) Jin, H. J.; Fridrikh, S. V.; Rutledge, G. C.; Kaplan, D. L. Biomacromolecules 2002, 3, 1233-1239. (18) Jia, H. F.; Zhu, G. Y.; Wang, P. Biotechnol. Bioeng. 2003, 84, 406-414. (19) Chen, J. P.; Su, D. R. Biotechnol. Prog. 2001, 17, 369-375. (20) Caruso, F.; Schuler, C. Langmuir 2000, 16, 9595-9603. (21) Tripp, S. L.; Pusztay, S. V.; Ribbe, A. E.; Wei, A. J. Am. Chem. Soc. 2002, 124, 7914-7915. (22) Sun, S. H.; Murray, C. B.; Weller, D.; Folks, L.; Moser, A. Science 2000, 287, 1989-1992. (23) Kolny, J.; Kornowski, A.; Weller, H. Nano Lett. 2002, 2, 361364. (24) Kariuki, N. N.; Han, L.; Ly, N. K.; Patterson, M. J.; Maye, M. M.; Liu, G. J.; Zhong, C. J. Langmuir 2002, 18, 8255-8259. (25) Dutta, A. K.; Ho, T. T.; Zhang, L. Q.; Stroeve, P. Chem. Mater. 2000, 12, 1042-1048. (26) Sun, S. H.; Anders, S.; Hamann, H. F.; Thiele, J. U.; Baglin, J. E. E.; Thomson, T.; Fullerton, E. E.; Murray, C. B.; Terris, B. D. J. Am. Chem. Soc. 2002, 124, 2884-2885.

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nanoparticle assembly has employed molecular recognition.28-30 For instance, several groups have modified nanoparticles with oligonucleotides for the DNA-directed assembly of gold nanoparticles,31-35 semiconductor quantum dots,36 and virus particles.9 It may also be desirable to localize nanoparticle assembly to specific, addressable regions. Soft lithography has often been used to pattern surfaces to have laterally varying surface functionalities. Such patterned surfaces have been exploited to direct the assembly of polyelectrolyte microcapsules37 and to spatially control layer-bylayer assembly38 for the incorporation of nanocrystals.39 We are examining an alternative, voltage-dependent approach to spatially assemble nanoparticles onto addressable surfaces. Specifically, we mediate nanoparticle electrodeposition using the pH-responsive aminopolysaccharide chitosan. Chitosan is a cationic polyelectrolyte at low pH (pH < 6), but becomes less charged and insoluble as the pH approaches or exceeds its pKa (chitosan’s apparent pKa ≈ 6.3).40-44 As illustrated in Scheme 1, chitosan can be electrodeposited at the cathode because proton consumption at this electrode leads to a localized region of high pH that can exceed chitosan’s solubility limit.45,46 Previous studies have shown that chitosan is electrodeposited onto patterned gold cathodes with high spatial selectivity47 and that deposition can be controlled by adjusting current density and bulk solution pH.46 Adhesion between the deposited chitosan film and the gold surface is stable and reversible, and the adhesion mechanism appears to involve physical interactions. After rinsing, the film is retained in air and liquid environments without the need for an applied voltage. However, the deposited films can be removed by treatment with mild acid to resolubilize chitosan, and thick chitosan films (≈40 (27) Sun, S. H.; Anders, S.; Thomson, T.; Baglin, J. E. E.; Toney, M. F.; Hamann, H. F.; Murray, C. B.; Terris, B. D. J. Phys. Chem. B 2003, 107, 5419-5425. (28) Boal, A. K.; Frankamp, B. L.; Uzun, O.; Tuominen, M. T.; Rotello, V. M. Chem. Mater. 2004, 16, 3252-3256. (29) Frankamp, B. L.; Uzun, O.; Ilhan, F.; Boal, A. K.; Rotello, V. M. J. Am. Chem. Soc. 2002, 124, 892-893. (30) Liu, J.; Mendoza, S.; Roman, E.; Lynn, M. J.; Xu, R. L.; Kaifer, A. E. J. Am. Chem. Soc. 1999, 121, 4304-4305. (31) Elghanian, R.; Storhoff, J. J.; Mucic, R. C.; Letsinger, R. L.; Mirkin, C. A. Science 1997, 277, 1078-1081. (32) Storhoff, J. J.; Mirkin, C. A. Chem. Rev. 1999, 99, 1849-1862. (33) Mirkin, C. A.; Letsinger, R. L.; Mucic, R. C.; Storhoff, J. J. Nature 1996, 382, 607-609. (34) Alivisatos, A. P.; Johnsson, K. P.; Peng, X. G.; Wilson, T. E.; Loweth, C. J.; Bruchez, M. P.; Schultz, P. G. Nature 1996, 382, 609611. (35) Loweth, C. J.; Caldwell, W. B.; Peng, X. G.; Alivisatos, A. P.; Schultz, P. G. Angew. Chem., Int. Ed. 1999, 38, 1808-1812. (36) Mitchell, G. P.; Mirkin, C. A.; Letsinger, R. L. J. Am. Chem. Soc. 1999, 121, 8122-8123. (37) Nolte, M.; Fery, A. Langmuir 2004, 20, 2995-2998. (38) Clark, S. L.; Hammond, P. T. Adv. Mater. 1998, 10, 1515+. (39) Chen, C. C.; Yet, C. P.; Wang, H. N.; Chao, C. Y. Langmuir 1999, 15, 6845-6850. (40) Rinaudo, M.; Pavlov, G.; Desbrieres, J. Polymer 1999, 40, 70297032. (41) Sorlier, P.; Denuziere, A.; Viton, C.; Domard, A. Biomacromolecules 2001, 2, 765-772. (42) Varum, K. M.; Ottoy, M. H.; Smidsrod, O. Carbohydr. Polym. 1994, 25, 65-70. (43) Strand, S. P.; Tommeraas, K.; Varum, K. M.; Ostgaard, K. Biomacromolecules 2001, 2, 1310-1314. (44) Anthonsen, M. W.; Smidsrod, O. Carbohydr. Polym. 1995, 26, 303-305. (45) Wu, L. Q.; Gadre, A. P.; Yi, H. M.; Kastantin, M. J.; Rubloff, G. W.; Bentley, W. E.; Payne, G. F.; Ghodssi, R. Langmuir 2002, 18, 86208625. (46) Fernandes, R.; Wu, L. Q.; Chen, T. H.; Yi, H. M.; Rubloff, G. W.; Ghodssi, R.; Bentley, W. E.; Payne, G. F. Langmuir 2003, 19, 40584062. (47) Wu, L.-Q.; Yi, H.; Li, S.; Rubloff, G. W.; Bentley, W. E.; Ghodssi, R.; Payne, G. F. Langmuir 2003, 19, 519-524.

Wu et al. Scheme 1. Illustration of Chitosan-Mediated Electrodepositiona

a Polarization of the gold cathode leads to proton consumption and a localized pH gradient. Chitosan chains in the bulk solution (pH < 6.0) remain soluble (indicated by thin lines with associated cationic sites). Chitosan chains near the cathode experience a higher pH and become insoluble (indicated by thick lines), forming a film. Nanoparticles suspended in the chitosan solution become entrapped when chitosan is electrodeposited.

mm) can be peeled from the surface if they are soaked overnight in strong base (1 M NaOH). Because of chitosan’s nucleophilic properties, the deposited chitosan films can be readily functionalized with proteins48,49 and nucleic acids.50,51 We should note that others have used polymers to assist the deposition of metal oxide films52,53 and to combine electrochemical and electrophoretic deposition to generate composite organic-inorganic films.54-57 Also, others have micropatterned electrodes and applied localized fields to direct layer-by-layer assembly to generate multilayers containing nanocrystals58 or enzymes.59 Here, we report that chitosan can mediate the electrodeposition of 100 nm fluorescently labeled latex spheres (i.e., model nanoparticles) onto the patterned cathodes. As illustrated in Scheme 1, we suspended these fluorescently labeled nanoparticles in the chitosan solution and immersed the cathodes into this suspension to initiate deposition. Our results indicate that the suspended nanoparticles are trapped throughout the electrodeposited chitosan film and that deposition can be spatially controlled. Materials and Methods Chitosan from crab shells (15% deacetylation and a molecular weight of 200 000 as reported by the supplier) and glass slides coated with indium tin oxide (ITO) were purchased from Sigma(48) Chen, T.; Small, D. A.; Wu, L.-Q.; Rubloff, G. W.; Ghodssi, R.; Vazquez-Duhalt, R.; Bentley, W. E.; Payne, G. F. Langmuir 2003, 19, 9382-9386. (49) Kastantin, M. J.; Li, S.; Gadre, A. P.; Wu, L. Q.; Bentley, W. E.; Payne, G. F.; Rubloff, G. W.; Ghodssi, R. Sens. Mater. 2003, 15, 295311. (50) Yi, H. M.; Wu, L. Q.; Sumner, J. J.; Gillespie, J. B.; Payne, G. F.; Bentley, W. E. Biotechnol. Bioeng. 2003, 83, 646-652. (51) Yi, H. M.; Wu, L. Q.; Ghodssi, R.; Rubloff, G. W.; Payne, G. F.; Bentley, W. E. Anal. Chem. 2004, 76, 365-372. (52) Takenaka, S.; Kozuka, H. Appl. Phys. Lett. 2001, 79, 34853487. (53) Jia, Q. X.; Mccleskey, T. M.; Burrell, A. K.; Lin, Y.; Collis, G. E.; Wang, H.; Li, A. D. Q.; Foltyn, S. R. Nat. Mater. 2004, 3, 529-532. (54) Zhitomirsky, I.; Petric, A. Mater. Sci. Eng., B 2000, 78, 125130. (55) Zhitomirsky, I.; Niewczas, M.; Petric, A. Mater. Lett. 2003, 57, 1045-1050. (56) Zhitomirsky, I. Surf. Eng. 2004, 20, 43-47. (57) Pang, X.; Zhitomirsky, I. Langmuir 2004, 20, 2921-2927. (58) Gao, M. Y.; Sun, J. Q.; Dulkeith, E.; Gaponik, N.; Lemmer, U.; Feldmann, J. Langmuir 2002, 18, 4098-4102. (59) Shi, L. X.; Lu, Y. X.; Sun, J.; Zhang, J.; Sun, C. Q.; Liu, J. Q.; Shen, J. C. Biomacromolecules 2003, 4, 1161-1167.

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Figure 1. Chitosan-mediated electrodeposition of fluorescently labeled nanoparticles onto an ITO-coated glass slide. (a) Fluorescence photomicrograph of nanoparticles electrodeposited from suspension (0.003% nanoparticles, 1% chitosan, pH ) 5) for 3 min. (b) Fluorescence photomicrograph of nanoparticles electrodeposited from a concentrated suspension (0.03% nanoparticles, 1% chitosan, pH ) 5) for 10 min. (c) Magnified view of the sample from part b. Aldrich Chemicals. Fluorescently labeled latex nanoparticles (FluoSpheres, 100 nm, excitation maximum 540 nm and emission maximum 560 nm) and 5- (and 6-)carboxyfluorescein succinimidyl ester (NHS-fluorescein, excitation maximum 495 nm and emission maximum 519 nm) were purchased from Molecular Probes. A chitosan solution was prepared by adding chitosan flakes to water and incrementally adding small amounts of HCl to the solution to maintain the pH near 3. After filtering undissolved material, the pH of the chitosan solution was adjusted to 5.0 using NaOH (1 M). Fluorescently labeled chitosan was prepared by reacting chitosan with NHS-fluorescein.47 Gold patterned silicon wafers were prepared using standard photolithographic methods.45,47 As described in the text, the cathode for electrodeposition was either an ITO-coated glass slide or a gold patterned silicon wafer, and the anode was either an ITO-coated glass slide or an unpatterned gold-coated silicon wafer. For electrodeposition, these electrodes were immersed in a suspension of fluorescently labeled latex nanoparticles in chitosan (pH ) 5.0, 1%, w/w, polymer). In some experiments, fluorescein-labeled chitosan was used in place of chitosan. Deposition was performed by connecting both the cathode and the anode to a direct current power supply (model 6614C, Agilent Technologies) and applying a voltage to achieve a constant current density of 0.05 mA/cm2. The deposition time and the concentration of nanoparticles were varied in individual experiments (see text). After deposition, the electrodes were disconnected from the power supply, removed from the suspension, rinsed with distilled water, and dried at room temperature. The gold patterned wafers were examined by a fluorescence stereomicroscope (MZFLIII, Leica) using two fluorescence filter sets (FluoIII). The first filter set was used to detect the fluorescently labeled nanoparticles using an excitation wavelength of 560 nm (bandwidth of 40 nm) and a long-pass filter at 610 nm. The second filter set was used to detect fluoresceinlabeled chitosan using an excitation wavelength of 480 nm (bandwidth of 40 nm) and a long-pass filter at 510 nm. Photomicrographs were prepared from the fluorescence microscope using a digital camera (Spot 32, Diagnostic Instruments). In some cases, the spatial resolution of electrodeposition was examined by analyzing the fluorescence intensity profile using standard imaging software (Scion Corp.). Deposits on the ITO-coated glass slides and the gold patterned silicon wafers were examined using a laser scanning confocal microscope (Leica TCS SP2) equipped with a 100× oil immersion objective lens. To image the fluorescently labeled nanoparticles we used an excitation wavelength of 488 nm and collected the emitted light in the range of 560-580 nm. The images were optically sectioned in the direction normal to the substrate surface (i.e., the z direction) using z-step sizes of 122.1 nm. We observed that rehydration of the deposited films resulted in considerable swelling. To study film rehydration, we first electrodeposited nanoparticle-containing chitosan films onto gold patterned surfaces and then air-dried the films at room temperature overnight. To rehydrate the dried films, we added deionized water to the sample, waited for 1 min, and placed a coverslip on top of the sample. Silicon oil was then applied to the boundary between the coverslip and the wafer to limit evaporation

during imaging. Subsequent film drying was performed using a vacuum oven (45 °C) for approximately 90 min.

Results and Discussion Role of Chitosan in Particle Electrodeposition. In our initial study to demonstrate particle electrodeposition, we suspended fluorescently labeled nanoparticles (100 nm diameter) into a chitosan solution and electrodeposited this suspension onto an electrode surface. Both the cathode and the anode in these experiments were ITOcoated glass slides that were immersed in the suspension and polarized at a constant current density (0.05 mA/ cm2). After deposition, the electrodes were disconnected from the power supply, removed from the solution, rinsed with distilled water, and examined using a confocal microscope. When the cathode was examined, a significant number of labeled nanoparticles were observed in the field of view as shown by the fluorescence photomicrograph of Figure 1a. Figure 1b shows that more nanoparticles are deposited at the cathode when the deposition time is increased (10 vs 3 min) and the nanoparticle concentration is increased (0.03 vs 0.003%). Figure 1c shows a magnified fluorescence image of the sample from Figure 1b and demonstrates that the nanoparticles in the deposited film are well-dispersed laterally. When the anode was examined, no fluorescence could be observed indicating that deposition is limited to the cathode. Thus, Figure 1 provides initial evidence for chitosan-mediated particle electrodeposition. To assess the spatial selectivity of nanoparticle electrodeposition we fabricated the silicon wafer of Figure 2a to have micropattterned gold lines, and we used this patterned wafer as our cathode. In our experiment, both the patterned cathode and the unpatterned anode (i.e., an unpatterned gold-coated wafer) were immersed in the chitosan solution that contained fluorescently labeled nanoparticles. After polarizing the electrodes (0.05 mA/ cm2) for 3 min, the wafers were disconnected from the power supply and removed from the suspension, and the cathode was rinsed with distilled water and examined. The fluorescence photomicrograph in Figure 2b shows that the nanoparticles are only deposited onto the conducting gold surfaces that served as the cathode. The spatial resolution of nanoparticle electrodeposition on the cathodes was assessed by image analysis of the fluorescence in the region enclosed by the dashed lines in Figure 2b. The fluorescence intensity profile in Figure 2c shows that the chitosan-mediated deposition of the fluorescently labeled nanoparticles is achieved with high spatial resolution. As a control, we electrodeposited chitosan in the absence of fluorescently labeled nanoparticles and observed no fluorescence in the image (not shown),

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Figure 2. Spatially selective electrodeposition of fluorescently labeled nanoparticles onto a patterned cathode. (a) Reflection photomicrograph of the gold-patterned silicon wafer before deposition. (b) Fluorescence photomicrograph of the fluorescently labeled nanoparticles that were electrodeposited onto the patterned gold cathodes. Electrodeposition was performed from a suspension (0.003% nanoparticles, 1% chitosan, pH ) 5) for 3 min, and the fluorescence was examined with the filter set with an excitation wavelength of 560 nm (bandwidth 40 nm). (c) Fluorescence intensity profile for the image in part b that is enclosed by the dashed lines.

indicating that neither chitosan nor the patterned gold contributed fluorescence in Figure 2b. We performed a series of controls to demonstrate that chitosan is necessary for deposition and that chitosan confers spatial selectivity to deposition. In the first control, deposition was attempted from a suspension of fluorescently labeled nanoparticles in the absence of chitosan. Reflection (Figure 3a) and fluorescence (Figure 3b) photomicrographs indicate that no deposition occurred for this “chitosan-less” control; thus, chitosan is necessary for deposition. In a second set of controls, we performed deposition experiments with a fluorescein-labeled chitosan that appears green using our filter set. Deposition of this fluorescein-labeled chitosan onto the patterned cathode occurs with high spatial selectivity when either (i) the

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labeled chitosan is deposited by itself (Figure 3c)47 or (ii) the labeled chitosan is deposited in the presence of the fluorescently labeled nanoparticles (Figure 3d). Thus, chitosan confers spatial selectivity to deposition. Spatial Distribution of Particles in the Electrodeposited Chitosan Film. While wide field images showed the nanoparticles to be well-distributed laterally, confocal imaging was required to determine if the particles were distributed throughout the chitosan film in the z direction (or if the particles were preferentially adsorbed at an interfacesthe inner gold-chitosan interface or the outer surface of the chitosan film). For this study, we electrodeposited a suspension of fluorescently labeled nanoparticles in chitosan (0.03% nanoparticles in 1% chitosan solution) onto a 20 µm patterned gold line. As illustrated by the schematic in Figure 4a, when the deposit is air-dried at room temperature the film becomes very thin (on the order of 1 µm as measured by profilometry). A fluorescence projection image of a dried film is shown in Figure 4b, and from this image, the film was estimated to be 2.5 µm thick. Because the dried films are so thin, we are unable to perform accurate optical cross-sectioning in the z direction, and, thus, it is not possible to determine if the nanoparticles are located throughout these dried films. As illustrated in Figure 4a, hydration leads to swelling of the chitosan film. Figure 4c shows a fluorescence projection image of a hydrated nanoparticle-containing film. The patterned gold line is also apparent in this cross section. Swelling of the hydrated chitosan film makes it possible to perform optical cross sectioning, and Figure 4d shows the fluorescence intensity profile in the z direction. This profile indicates that the fluorescent nanoparticles are distributed throughout the film. We did not attempt to quantify the nanoparticle distribution throughout the film (i.e., to determine if the nanoparticles are distributed homogeneously) because of difficulties in accounting for the strong reflection from the gold surface and potential quenching effects. The intensity profile in Figure 4d indicates that the fluorescence extends 11.5 µm from the gold surface. We believe the edge of the fluorescence corresponds to the edge of the chitosan film, although this could not be independently confirmed. More importantly, Figure 4d shows that the nanoparticles are distributed throughout the chitosan film and that the nanoparticles are not confined to an interface. The hydrated nanoparticle-containing film was examined using the confocal microscope. Figure 5a shows the overall three-dimensional projection image for this film while Figure 5b shows a magnified projection image of a central region of the film. This latter image indicates that

Figure 3. Controls that indicate that chitosan is required for electrodeposition and chitosan confers spatial-selectivity to electrodeposition. (a) Reflection and (b) fluorescence photomicrographs for the patterned cathodes after experiments to electrodeposit fluorescently labeled nanoparticles in the absence of chitosan. (c) Fluorescence photomicrograph after the electrodeposition of fluorescein-labeled chitosan in the absence of nanoparticles. (d) Fluorescence photomicrograph after the electrodeposition of fluoresceinlabeled chitosan in the presence of the fluorescently labeled nanoparticles. The fluorescein-labeled chitosan was examined using the filter set with an excitation wavelength of 480 nm (bandwidth 40 nm).

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Figure 6. Reversibility of film hydration. (a) Nanoparticle fluorescence intensity profiles for a film sequentially hydrated four times. (b) Thickness of the nanoparticle-containing films during four sequential drying and rehydration steps.

Figure 4. Fluorescently labeled nanoparticles deposit throughout the film in the z direction. (a) Schematic showing the cross section of dried and hydrated films that had been electrodeposited onto a patterned gold cathode (in experiments the gold line was 20 µm wide). Fluorescence projection image for the labeled nanoparticles in (b) dried and (c) hydrated films. (d) Fluorescence intensity profile for the labeled nanoparticles in the hydrated film. Electrodeposition was performed for 10 min from a suspension (0.03% nanoparticles in 1% chitosan).

not disrupt the nanoparticle distribution within the chitosan film or lead to ejection of the nanoparticles. In these experiments, we performed a series of drying and rehydration steps and examined the films using the confocal microscope. Figure 6a shows the nanoparticle fluorescence profiles for a film that had been sequentially rehydrated four times. Although the film’s profile after the initial hydration step appears somewhat different from the profiles observed after subsequent hydration steps, we believe this difference is due to difficulties in performing this experiment. To limit water evaporation during imaging we added oil to the interface between the wafer and the coverslip, and we believe that contact between the film and the oil reduced to some extent the film’s ability to swell in subsequent hydration steps. The similarity of the shapes of the profiles in Figure 6a provides evidence that the fluorescent beads remain distributed throughout the film. We did not attempt to quantify particle retention by integrating the fluorescence intensity profiles because we were unable to repeatedly image the same region of the film during this 10 day experiment. Nevertheless, we believe the majority of the particles remain entrapped within the polymeric network during repeated rehydrations because we did not visually observe fluorescence in wetted regions away from the films. Figure 6b compares the thicknesses of the nanoparticlecontaining films during the four sequential drying and rehydration steps. As shown, rehydration results in about a fourfold increase in film thickness and the observed changes in the film thickness were repeatable. The fact that this film did not delaminate during the 10 days of this sequential drying and rehydration experiment provides further evidence for the stable adhesion between the chitosan film and the substrate surface. Conclusions

Figure 5. Three-dimensional projection images for the hydrated nanoparticle-containing film. (a) Overall three-dimensional projection image. (b) Magnified three-dimensional projection image.

individual nanoparticles and aggregates are distributed throughout the hydrated chitosan film. Thus, Figures 4 and 5 show that particles are distributed throughout the electrodeposited film. Reversibility of Film Hydration. The observation that electrodeposited chitosan films swell substantially upon rehydration may have important implications as stimuli-responsive and addressable sensors.60-63 We performed initial studies to ensure that film hydration does

We demonstrate that nanoparticles can be electrodeposited onto cathode surfaces using the pH-responsive aminopolysaccharide chitosan. When micropatterned gold cathodes were used, we observed that nanoparticle deposition can be spatially controlled with a lateral resolution of 20 µm (this is the resolution limit for the microfabrication method we used to pattern gold onto the silicon wafer). Additionally, our results indicate that the nanoparticles are distributed throughout the chitosan film (60) Lavigne, J. J.; Anslyn, E. V. Angew. Chem., Int. Ed. 2001, 40, 3119-3130. (61) Gunter, R. L.; Delinger, W. G.; Manygoats, K.; Kooser, A.; Porter, T. L. Sens. Actuators, A 2003, 107, 219-224. (62) Kooser, A.; Gunter, R. L.; Delinger, W. D.; Porter, T. L.; Eastman, M. P. Sens. Actuators, B 2004, 99, 474-479. (63) Lei, M.; Gu, Y.; Baldi, A.; Siegel, R. A.; Ziaie, B. Langmuir 2004, 20, 8947-8951.

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(and not localized at an interface). Finally, the chitosan films were observed to reversibly swell upon rehydration. These results indicate that chitosan-mediated electrodeposition provides a simple means to assemble nanoparticles at addressable locations. We expect chitosan-mediated nanoparticle deposition to be broadly applicable, within the constraints that (i) the nanoparticles can be suspended in the chitosan solution and (ii) the nanoparticle size exceeds the mesh size of the deposited chitosan. The former constraint is common to many applications of nanoparticles, and surface modifications are often necessary to enhance particle solubility and diminish aggregation. The latter constraint may also be unimportant because the mesh sizes of polymer networks are on the order of 10100 nm64-66 and particles below this size range (e.g., proteins) could be covalently attached to chitosan prior to deposition.48,67 Potentially, this work could provide a generic means to assemble nanoscale particles into functional devices. A variety of nanoscale particles (e.g., quantum dots and proteins) offer distinct properties that could be enlisted to detect and transduce a host of signals (e.g., optical, electrical, and biological). Additional nanoparticles (e.g., (64) Lowman, A. M.; Peppas, N. A. Macromolecules 1997, 30, 49594965. (65) Morse, D. C. Macromolecules 1998, 31, 7044-7067. (66) Berger, J.; Reist, M.; Mayer, J. M.; Felt, O.; Peppas, N. A.; Gurny, R. Eur. J. Pharm. Biopharm. 2004, 57, 19-34. (67) Vazquez-Duhalt, R.; Tinoco, R.; D’Antonio, P.; Topoleski, L. D. T.; Payne, G. F. Bioconjugate Chem. 2001, 12, 301-306.

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vesicles and virus particles) provide compartments that allow individualized components (e.g., therapeutics, reagents, or catalysts) to be controllably delivered and released. However, exploiting the potential of these nanomaterials will, in many cases, require their controllable assembly into higher ordered structures. The ability of chitosan to entrap these particles within a polymeric mesh that forms in response to localized electrical stimuli provides a particularly simple means to direct nanoparticle assembly at specific addresses. Further, the ability of the chitosan matrix to reversibly swell in response to other stimuli (i.e., hydration) may provide a mechanism for “reporting” localized environmental conditions (e.g., by monitoring signals that vary with the spacing between nanoparticles). More broadly, this work provides another example of how stimuli-responsive biopolymers can be used for microfabrication.68 Note Added in Proof. After submitting this manuscript, a report appeared on the use of chitosan to electrodeposit MnO2 nanoparticles.69 Acknowledgment. Financial support for this work was provided by the United States National Science Foundation (Grant BES-0114790). W.L. acknowledges equipment funding from the Office of Naval Research. LA047420C (68) Wu, L. Q.; Payne, G. F. Trends Biotechnol. 2004, 22, 593-599. (69) Xu, J.; Luo, X.; Du, Y.; Chen, H. Electrochem. Commun. 2004, 6, 1169-1173.