Thin Films of Polyelectrolyte-Encapsulated Catalase Microcrystals for

May 10, 2003 - Thin Films of Polyelectrolyte-Encapsulated Catalase Microcrystals for Biosensing ..... part of its Biofuture research program and the A...
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Anal. Chem. 2003, 75, 3031-3037

Thin Films of Polyelectrolyte-Encapsulated Catalase Microcrystals for Biosensing Aimin Yu†,§ and Frank Caruso*,‡

Max Planck Institute of Colloids and Interfaces, D-14424 Potsdam, Germany, and Department of Chemical and Biomolecular Engineering, The University of Melbourne, Victoria 3010, Australia

Polyelectrolyte (PE)-encapsulated catalase microcrystals were assembled onto gold electrodes by their sequential deposition with oppositely charged PEs, utilizing electrostatic interactions to form enzyme thin films for biosensing. The PE coating around the microcrystals provided a regular surface charge, thus facilitating the stepwise film growth, and it effectively prevented catalase leakage from the assembled films. The encapsulated catalase was shown to retain both its biological and its electrochemical activity. Direct electron transfer between catalase molecules and the gold electrode was achieved without the aid of any electron mediator. In pH 5.0 phosphate buffer solution, the apparent formal potential (E°′) of catalase was -0.131 V (vs Ag/AgCl). As a H2O2 biosensor, films consisting of one layer of the encapsulated catalase displayed considerably higher (∼5-fold) and more stable electrocatalytic responses to the reduction of H2O2 than did corresponding films made of one layer of nonencapsulated catalase or solubilized catalase. An increase in either the number of “precursor” PE layers between the gold electrodes and the catalase microcrystal layers in the film or the number of PE layers encapsulating the catalase microcrystals was found to decrease the electrocatalytic activity of the electrode. At low precursor PE layer numbers (∼2) and PE encapsulating layers (∼4), the current response was proportional to the H2O2 concentration in the range 3.0 × 10-6 to 1.0 × 10-2 M. The overall electroactivity of the multilayer film increased for the first two layers of encapsulated catalase, after which a plateau was observed. This was attributed to the increasing difficulty of electron transfer and substrate diffusion limitations. The current approach of using immobilized PE-encapsulated enzyme microcrystals for biosensing provides a versatile method to prepare high enzyme content films with high and tailored enzyme activities. Electron transfer reactions between redox proteins are essential to metabolic process in living organisms. This has motivated electrochemical studies on the direct electron transfer * To whom correspondence should be addressed. Fax: +61 3 8344 4153. E-mail: [email protected]. † Max Planck Institute of Colloids and Interfaces. ‡ The University of Melbourne. § Permanent address: Department of Chemistry, Nanjing University, 210093, P. R. China. 10.1021/ac0340049 CCC: $25.00 Published on Web 05/10/2003

© 2003 American Chemical Society

between enzymes and electrodes, largely because enzymes can be activated and regenerated by the exchange of electrons between electrodes and redox proteins.1,2 Furthermore, enzymecoated electrodes provide the basis for constructing biosensors, biomedical devices, and enzymatic bioreactors that have wide application in biotechnology.3 However, various factors can prohibit direct electron transfer between electrodes and proteins, including the deep burial of the electroactive cofactors in the protein structure, adsorption denaturation of the protein at the electrode surface, and unfavorable orientation of the protein structure.4 Much effort has been focused on facilitating electron transfer between proteins and electrodes, including using electrontransfer mediators,5 and modifying electrode surfaces to provide an electron-transfer compatible interface.6 Another viable method is to cast proteins, such as myoglobin or cytochrome P450, into insoluble surfactant films on pyrolytic graphite electrodes, which has shown to achieve reversible voltammetry of the proteins.2 However, the main drawback of frequently employed methods used to immobilize proteins on electrode surfaces (e.g., covalent binding, entrapment in matrixes) is that it is difficult to control the architecture of the films, and often, these methods produce irregular films with a low density of protein, thus limiting their wide application. The layer-by-layer (LbL) assembly method, introduced by Decher and Hong in 1991,7 has emerged as a promising and versatile approach to fabricate functional molecular assemblies with well-defined architectures and with nanoscale-level control over the film thickness. The basis of this method is the stepwise electrostatic assembly of oppositely charged species. Various multilayered systems composed of inorganic nanoparticles,8 dye (1) (a) Armstrong, F. A.; Hill, H. A. O.; Walton, N. J. Acc. Chem. Res. 1988, 21, 407. (b) Armstrong, F. A.; Herring, H. A.; Hirst, J. Chem. Soc. Rev. 1997, 26, 169. (2) Rusling, J. F. Acc. Chem. Res. 1998, 31, 363. (3) For examples, see: (a) Wilson, G. In Biosensors; Turner, A., Karube, I., Wilson, G., Eds; Oxford University Press: New York, 1987. (b) Kauffmann, J. M.; Guibault, G. G. Bioanalytical Applications of Enzymes, Methods Biochem. Anal.; Wiley: New York, 1992; Vol. 36, pp 63-113. (4) Armstrong, F. A. Bioinorganic Chemistry: Structure and Bonding 72; Springer-Verlag: Berlin, 1990; pp 137-219. (5) (a) Song, S.; Dong, S. Bioelectrochem. Bioenerg. 1988, 19, 337. (b) Ye, J.; Baldwin, R. P. Anal. Chem. 1988, 60, 2263. (c) Ye, B.; Zhou, X. Electroanalysis 1996, 8, 1165. (d) Tatsuma, T.; Okawa, Y.; Watanabe, T. Anal. Chem. 1989, 61, 2352. (6) (a) Tarlov, M. J.; Bowden, E. F. J. Am. Chem. Soc. 1991, 113, 1847. (b) Reeves, J. H. Anal. Chem. 1993, 65, 683. (c) Glenn, J. D. H.; Bowden, E. F. Chem. Lett. 1996, 399. (7) (a) Decher, G.; Hong, J. D. Ber. Bunsen-Ges. Phys. Chem. 1991, 95, 1430. (b) For a review, see: Decher, G. Science. 1997, 277, 1232.

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molecules,9 and dendrimers10 alternating with polyelectrolytes (PE) have been prepared by this technique. Water-soluble proteins, which are charged at various pHs, such as streptavidin,11 immunoglobulin G (IgG),12 albumin,13 bacteriorhodopsin,14 glucose oxidase15 myoglobin, and hemoglobin,15b,16 have been alternately assembled with oppositely charged polyelectrolytes to form multilayer films. The LbL method has also been applied to prepare biocatalytic core-shell particles with tailored enzymatic activities, depending on the number of enzyme layers deposited onto the core particles.17 Recent studies have investigated the electrochemical characteristics of LbL assembled films. Willner and coworkers examined the electron transfer characteristics of multilayer PE films and demonstrated that the layered charged polymer assembly exhibits a neutralized porous structure that allows redox transformations at the electrode surface.18 Bowden and co-workers used positively charged poly(lysine) to assemble films with negatively charged cytochrome b5 and showed that direct electron transfer of the protein could be achieved.19 A detailed review is provided by Rusling on the assembly and properties of electroactive and enzyme-active protein-PE films prepared by the LbL method.20 Recently, we presented a novel and general strategy for using biocrystals as templates for the controlled encapsulation of biomolecules by the LbL deposition of oppositely charged PEs.21,22 Advantages of using this approach are that it provides a high enzyme loading in encapsulated form, the bioactivity of the encapsulated enzyme is preserved, and the semipermeable PE coating prevents the solubilized enzyme from leakage while simultaneously permitting the diffusion of small (substrate) molecules for enzyme reaction.21 Additionally, the PE layers encapsulating the enzyme effectively increase the surface charge (8) (a) Feldheim, D. L.; Grabar, K. C.; Natan, M. J.; Mallouk, T. E. J. Am. Chem. Soc. 1996, 118, 7640. (b) Kotov, N. A.; Dekany, I.; Fendler, J. H. J. Phys. Chem. 1995, 99, 13065. (c) Musick, M. D.; Keating, C. D.; Keefe, M. H.; Natan, M. J. Chem. Mater. 1997, 9, 1499. (9) (a) Araki, K.; Wagner, M. J.; Wrighton, M. S. Langmuir 1996, 12, 5393. (b) Ariga, K.; Lvov, Y.; Kunitake, T. J. Am. Chem. Soc. 1997, 119, 2224. (c) Tedeschi, C.; Caruso, F.; Mo¨hwald, H.; Kirstein, S. J. Am. Chem. Soc. 2000, 122, 5841. (10) Khopade A. J.; Caruso, F. Langmuir 2002, 18, 7669. (11) Cassier, T.; Lowack, K.; Decher, G. Supramol. Sci. 1998, 5, 309. (12) (a) Caruso, F.; Niikura, K.; Furlong, D. N.; Okahata, Y. Langmuir 1997, 13, 3427. (b) Caruso, F.; Furlong, N.; Ariga, K.; Ichinose, I.; Kunitake T. Langmuir 1998, 14, 4559. (13) Houska, M.; Brynda, E. J. Colloid Interface Sci. 1997, 188, 243. (14) He, J.-A.; Samuelson, L.; Li, L.; Kumar, J.; Tripathy, S. K. J. Phys. Chem. B 1998, 102, 7067. (15) (a) Hodak, J.; Etchenique, R.; Calvo, E.; Singhal, K.; Bartlett, P. Langmuir 1997, 13, 2708. (b) Lvov, Y.; Ariga, K.; Ichinose, I.; Kunitake, T. J. Am. Chem. Soc. 1995, 117, 6117. (16) (a) Lvov, Y.; Ariga, K.; Kunitake, T. Chem. Lett. 1994, 2323. (b) Lvov, Y.; Lu, Z.; Schenkman, J.; Zu, X.; Rusling, J. F. J. Am. Chem. Soc. 1998, 120, 4073. (17) (a) Caruso, F.; Fiedler, H.; Haage, K. Colloids Surf. A 2000, 169, 287. (b) Caruso, F.; Schu ¨ ler, C. Langmuir 2000, 16, 9595. (c) Schu ¨ ler, C.; Caruso, F. Macromol. Rapid Commun. 2000, 21, 750. (d) Lvov, Y.; Caruso, F. Anal. Chem. 2001, 73, 4212. (e) For a review, see: Caruso, F. Chem.-Eur. J. 2000, 413. (18) Pardo-Yissar, V.; Katz, E.; Lioubashevski, O.; Willner, I. Langmuir 2001, 17, 1110. (19) Glenn, J. D. H.; Bowden, E. F. Chem. Lett. 1996, 399. (20) Rusling, J. F. In Protein Architecture: Interfacing Molecular Assemblies and Immobilization Biotechnology; Lvov, Y., Mo ¨hwald, H., Eds; Marcel Dekker: New York, 2000. (21) Caruso, F.; Trau, D.; Mo¨hwald, H.; Renneberg, R. Langmuir 2000, 16, 1485. (22) Jin, W.; Shi, X.-Y.; Caruso, F. J. Am. Chem. Soc. 2001, 123, 8121.

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density of the enzyme microcrystals,22 making them suitably charged components for the construction of biofunctional thin films. We also showed that the LbL deposition of PE-encapsulated microcrystals and oppositely charged PE allows the preparation of multilayered films with tailored bioactivity (depending on the number of microcrystal layers deposited).22 In a related study, it was demonstrated that PE-encapsulated organic microcrystals can be utilized to create highly amplified biochemical assays.23 In the present study, we extend our work in utilizing PEencapsulated enzyme microcrystal building blocks to prepare a high-activity electrochemical biosensor. We investigate the direct electron-transfer behavior of PE-encapsulated catalase and its electrocatalytic response to H2O2. The influence of both the number of PE layers between the electrode and the microcrystal layers and the effect of the number of PE layers used for encapsulation of the enzyme on the electrocatalytic properties of the films are investigated. Since the thickness of the PE layers can be tuned to within 1-2 nm, these films permit the study of nanoscale thickness and composition changes on the electrocatalytic responses. The effect of the number of enzyme multilayers deposited (that is, enzyme content) on the electrochemical behavior, as well as the temperature stability of the films prepared, are also examined. To our knowledge, this is the first study on the electrochemical behavior of PE multilayer-encapsulated enzymes. EXPERIMENTAL SECTION Chemicals. Catalase crystals (C-100) were obtained from Sigma. Poly(allylamine hydrochloride) (PAH, Mw 70 000), poly(sodium 4-styrene-sulfonate) (PSS, Mw 70 000), sodium chloride (NaCl), hydrogen peroxide (H2O2), and 3-mercaptopropionic acid (MPA) were obtained from Aldrich. Phosphate buffer solutions (PBS) of different pH were prepared from 0.1 M H3PO4, NaH2PO4, Na2HPO4, and NaOH. All chemicals were used as received, except for PSS, which was dialyzed against Milli-Q water (Mw cutoff 14 000) and lyophilized before use. The water used in all of the experiments was prepared in a three-stage Millipore Milli-Q Plus 185 purification system and had a resistivity higher than 18.2 MΩ cm. Catalase Encapsulation. The catalase crystals (positively charged at pH 5.0) were washed three times with a chilled (4 °C) solution of 1 M potassium acetate buffer at pH 5.0 with intermittent centrifugation steps (500g for 4 min at 4 °C) to remove the supernatant. The polymer layers were then assembled onto the catalase crystals by the sequential deposition of PSS and PAH. The first layer was deposited by adding a 0.5 mL aliquot of a 5 mg mL-1 aqueous PSS solution (containing 1 M potassium acetate buffer, pH 5.0, 4 °C) to 0.2 mL of the microcrystal suspension, occasionally shaking the suspension, and allowing 25 min for PSS adsorption. Excess PSS was removed by three repeated centrifugation/chilled buffer wash/redispersion cycles. PAH and additional PSS/PAH layers were deposited using the same procedure and conditions (as described) until the desired number of PE multilayers was achieved. Microelectrophoresis. The PE encapsulation of the catalase microcrystals was followed qualitatively by measuring the micro(23) Yang, W.; Trau, D.; Lehmann, M.; Caruso, F.; Yu, N. T.; Renneberg, R. Anal. Chem. 2002, 74, 5480.

electrophoretic mobility of the coated crystals using a Malvern Zetasizer 4 by taking the average of three measurements at the stationary level. The mobilities (µ) were converted to a ζ-potential using the Smoluchowski relation ζ ) µη/, where η and  are the viscosity and permittivity of the solution, respectively. All measurements were performed in air-equilibrated pure water (pH ∼5.8) without added electrolyte. Quartz Crystal Microbalance (QCM) Measurements. ATcut quartz crystals with a fundamental resonance frequency (Fo) of ∼9 MHz were supplied by Kyushu Dentsu Co. (Omura-City, Nagasaki, Japan). The 4.5-mm-diameter crystals were supplied with 100-nm-thick gold-coated electrodes. Before use, the connecting wires of the QCM electrodes were insulated with siloxane polymer. The crystals were then cleaned by using piranha solution (one part 30% H2O2 to three parts 98% H2SO4). (Caution: Piranha solution is a strong oxidizing agent which causes severe burns in contact with skin and reacts violently with organic materials.) Piranha solution was carefully spread on the gold surface of each QCM for 1 min, after which the electrode surfaces were thoroughly rinsed with Milli-Q water. This process was repeated an additional two times. The QCM measurement system has been described in detail elsewhere.24 After adsorption of the PEencapsulated enzyme microcrystals or PEs from solution, the electrodes were washed thoroughly with pure water and nitrogendried, and the QCM frequency change in air (Fair) was measured. The ∆Fair (frequency difference before and after adsorption) was used to determine the mass adsorbed after each immersion step according to the Sauerbrey equation:25

∆Fair ) -(1.83 × 104)∆mA where mA (g m-2) is the mass change per quartz crystal unit area. Preparation of PE-Encapsulated Catalase Microcrystal Films. Before coating, the electrodes were first immersed into a 1 mM ethanolic MPA solution for 24 h to generate a negatively charged surface. (An initial monolayer of MPA on Au has also been found to facilitate the direct electron transfer of proteins.26) They were then modified with a “precursor” film of PAH/PSS, which was formed by the alternate deposition of PAH and PSS from a 3 mg mL-1 aqueous solution (containing 0.5 M NaCl) of the corresponding PE. The precursor layers provide a uniform charge and a smooth surface, and they are important to obtain stably adsorbed enzyme layers.17d,21 The substrates were then immersed in a suspension of the PE-coated catalase crystals with an outermost layer of PAH (∼5 mg mL-1 in 1 M potassium acetate buffer, pH 5.0, 4 °C) for 60 min. Multilayer catalase films were constructed on the QCM electrodes by alternately adsorbing PSS layers (5 mg mL-1 in 1 M potassium acetate buffer, pH 5.0, 4 °C, 15 min) and PE-coated catalase microcrystal layers, with intermediate water washing (at 4 °C) and nitrogen drying. Finally, two additional PE layers were deposited on top of the outer microcrystal layer to prevent desorption of the crystals with time. Films (24) (a) Caruso, F.; Rodda, E.; Furlong, D. N. J. Colloid Interface Sci. 1996, 178, 104. (b) Caruso, F.; Rinia, H.; Furlong, D. N. Langmuir 1996, 12, 2145. (25) Sauerbrey, G. Z. Phys. 1959, 155, 206. (26) (a) Hill, H. A. O.; Hunt, N. I. Methods Enzymol. 1993, 227, 501. (b) Tarlov, M. J. Bowden, E. F. J. Am. Chem. Soc. 1991, 113, 1847. (c) Glenn, J. D. H.; Bowden, E. F. Chem. Lett. 1996, 399.

Figure 1. ζ-Potential of PSS/PAH-encapsulated catalase crystals as a function of the number of PE encapsulating layers. Odd layers correspond to PSS, and even layers, to PAH. The encapsulated crystals were redispersed in air-equilibrated pure water prior to measurement of the microelectrophoretic mobilities.

from nonencapsulated catalase crystals were prepared under the same conditions and in the same way as those of PE-encapsulated catalase. Films from solubilized catalase were obtained by adsorbing solubilized catalase (∼5 mg mL-1 in pH 7.0 PBS, 4 °C) for 60 min on the precursor film PAH/PSS/PAH. Electrochemical Measurements. Electrochemical measurements were performed with a three-electrode system consisting of a platinum wire as the auxiliary electrode; a Ag/AgCl (3 M KCl) as reference electrode, against which all potentials are quoted; and the QCM electrode as the working electrode. The electrodes were connected to an Autolab PGSTAT30 Electrochemical Instrument (Netherlands). Amperometric measurements were carried out in a stirred system by applying a potential step of -300 mV to the working electrode. Aliquots of a H2O2 standard solution were successively added to the solution. A single point was recorded after a steady-state current had been achieved. The response time is the time for obtaining the steady-state current after adding the H2O2 solution. All solutions were deoxygenated by bubbling high purity nitrogen for 15 min, and a nitrogen atmosphere was kept over the solutions during the measurements. RESULTS AND DISCUSSION PE-Encapsulation of Catalase Microcrystals. We chose catalase crystals as the templates because at pH ∼5-6, catalase exists as a crystalline suspension in water.27 In pH 5.0 acetate buffer, catalase is positively charged with a ζ-potential of 20 mV (catalase isoelectric point, pI ) 5.8). This positive charge on the surface of the crystals makes them suitably charged templates for the deposition of PE multilayers on their surface, beginning with a negatively charged PE. Figure 1 shows the change in the ζ-potential of catalase crystals when sequentially coated with PSS and PAH by the LbL method. After coating with PE layers, the ζ-potential of catalase changed to about -40 mV (outermost PSS layer) and +30 mV (outermost PAH layer). The reversal of the surface charge with increasing layer number demonstrates PE multilayer growth on the catalase crystal templates. (27) Biochemica Information; Boehringer: Mannheim, Germany, 1987; pp 1516.

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Figure 2. UV-vis spectra of (a) solubilized catalase in water, (b) (PSS/PAH)2-encapsulated catalase crystals in water, and (c) the supernatant after centrifugation of the four-layer PE-encapsulated catalase dispersion after exposure to pH 7.0 PBS for 3 days.

PE multilayers are known to be readily permeable to small mobile molecules (e.g., ions and low-molecular-weight dyes).28,29 Proteins such as catalase (diameter ∼9 nm), however, are sufficiently large; thus, after encapsulation, the PE multilayers can prevent the enzyme from leakage, even when it is solubilized in pH 6 solutions. This was proved by adding 4-layer PE-encapsulated catalase microcrystals to a pH 7.0 PBS solution for 3 days, sedimenting the encapsulated catalase crystals by centrifugation, and finally, measuring the amount of catalase in the supernatant solutions. Figure 2 shows the UV-vis absorption spectra of solubilized catalase in water (spectrum a) and 4-layer PE-encapsulated catalase crystals in water (spectrum b). The absorption peak at 406 nm corresponds to the Soret absorption band of the iron (III) heme structure in the catalase molecule.30Although catalase can be solubilized at pH 7.0,27 there is no distinct UV-vis absorption peak near 400 nm when the supernatant is measured after solubilization of the PE-encapsulated catalase (spectrum c), indicating no significant enzyme leakage. Formation of PE-Encapsulated Catalase Microcrystal Multilayer Films on QCM Electrodes. The adsorbed mass of catalase on the QCM electrodes can be measured via the QCM frequency shift according to the Sauerbrey relation.25 For the deposition of one layer of nonencapsulated catalase (Eo) and solubilized catalase (Es) onto a PAH/PSS precursor film, QCM frequency changes (∆F) of -410 ((150) Hz (or 356 ng) and -185 ((20) Hz (or 162 ng), respectively, were observed. These values are in close agreement with those obtained for various protein films prepared by a range of methods.31,32 However, after PE encapsulation, the mass loading of the deposited catalase microcrystals significantly increased. For example, when assembling (28) Caruso, F.; Donath, E.; Mo ¨wald, H. J. Phys. Chem. B 1998, 102, 2011. (29) Caruso, F.; Lichtenfeld, H.; Donath, E.; Mo¨hwald, H. Macromolecules 1999, 32, 2317. (30) Theorell, H.; Ehrenberg, A. Acta Chem. Scand. 1951, 5, 823. (b) George, P.; Hanania, G. Biochem. J. 1953, 55, 236. (c) Nassar, A.-E. F.; Willis, W. S.; Rusling, J. F. Anal. Chem. 1995, 67, 2386. (31) Caruso, F. In Protein Architecture: Interfacing Molecular Assemblies and Immobilization Biotechnology; Lvov, Y., Mo ¨hwald, H., Eds; Marcel Dekker: New York, 2000. (32) Proteins at Interfaces: Fundamentals and Applications; Horbett, T. A., Brash, J. L., Eds.; ACS Symposium Series 602; American Chemical Society: Washington, DC, 1995.

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Figure 3. Cyclic voltammograms of electrodes modified with (a) PAH/PSS/Eo/PSS/PAH and (b) PAH/PSS/EL4/PSS/PAH in pH 5.0 PBS at 50 mVs-1.

four-layer PE-encapsulated catalase microcrystals onto PAH/PSScoated QCM electrodes (PAH/PSS/EL4; EL4 is defined as fourlayer PE-encapsulated catalase), the average frequency decrease is ∼3584 ((1012) Hz, corresponding to an enzyme loading of ∼100 mg m-2. This enzyme coverage is ∼9 times higher than the value measured for the nonencapsulated catalase crystals, and ∼20 times higher than for the case of solubilized catalase. The difference in the nonencapsulated and PE-encapsulated enzyme values is mainly attributed to the enhanced surface charge of the microcrystals as a result of PE coating. Multilayer catalase microcrystal films were prepared by sequentially adsorbing PSS and PE-coated catalase microcrystals. The amount of the deposited PE-encapsulated enzyme linearly increased with the number of enzyme microcrystal adsorption steps, according to QCM experiments.22 Thus, the LbL method provides the possibility to build high enzyme content films with tailored enzyme loadings, which impacts on the film bioactivity (see below). Direct Electron Transfer of PE-Encapsulated Catalase and Its Electrochemical Response to H2O2. Catalase is known as an efficient H2O2 catalyst. The heme group in its molecular structure is the electroactive center of the enzyme.33 However, compared with other heme-containing proteins, such as cytochrome c, myoglobin, and horseradish peroxidase, there are few reports on its electrochemical behavior.34 In this work, we first studied the electrochemical behavior of nonencapsulated catalase at PE-modified electrodes using cyclic voltammetry. Figure 3 (curve a) shows the cyclic voltammograms (CV) of a PAH/PSS/ Eo/PSS/PAH-modified electrode in pH 5.0 PBS at 50 mV s-1. The nonencapsulated catalase displays a pair of redox peaks with an oxidation peak potential (Epa) at -0.051 V and a reduction peak potential (Epc) at -0.172 V. This is a typical quasi-reversible redox reaction resulting from the redox process of Fe2+/Fe3+ couple in the heme prosthetic structure.2,34 The apparent formal potential (E°′, estimated as the midpoint potentials of redox peaks) is calculated to be -0.111 V, which for comparison is slightly more (33) Voet, D.; Voet, J. F. Biochemistry, 2nd ed.; John Wiley & Sons: New York, 1995. (34) (a) Zhang, Z.; Chouchane, S.; Magliozzo, R. S.; Rusling, J. F. Chem. Commun. 2001, 177. (b) Chen, X.; Xie, H.; Kong, J. Biosens. Bioelectron. 2001, 16, 115. (c) Zhang, Z.; Chouchane, S.; Magliozzo, R. S.; Rusling, J. F. Anal. Chem. 2002, 74, 163. (d) Lai, M.; Bergel, A. Bioelectrochem. 2002, 55, 157.

Figure 4. Cyclic voltammograms of (a) PAH/PSS/EL4/PSS/PAHand (b) PAH/PSS/Eo/PAH/PSS-modified electrodes in pH 7.0 PBS containing 2 mM H2O2.

negative than the reported value for catalase entrapped in a didodecyldimethylammonium bromide crystal film (-0.073 V vs Ag/AgCl (3 M KCl), pH 5.0 buffer).34b Figure 3 (curve b) shows the CV of a PAH/PSS/EL4/PSS/PAH-modified electrode in pH 5.0 PBS. After being encapsulated in four PE layers, the redox peaks of catalase are still clearly observed. Compared with the CV of nonencapsulated catalase, the apparent formal potential is more negative (E°′ ) -0.131 V), and the peak currents increase considerably. This suggests that four layers of PE do not significantly impair electron transfer; direct electron transfer between four-layer PE-encapsulated catalase and the electrode can still be obtained. The biological activity of the encapsulated catalase was then examined by monitoring its catalytic activity toward H2O2. Figure 4 (curve a) shows the catalytic response of an EL4-modified electrode to 2 mM H2O2 in pH 7.0 PBS. It can be seen that the electrocatalytic current rises from ∼0 V and reaches a stable response at ∼-0.3 V. The cathodic peak current increased considerably, and the anodic peak current decreased, which is characteristic of an electrocatalytic reduction process. However, the overall mechanism of the electrocatalytic process is not clearly established. The accepted theory for the electrocatalytic process of peroxidase can be described as follows:34c,35

CatalaseFeIII + H2O2 f [CatalaseFeIVdO]• + H2O

(1)

[CatalaseFeIVdO]• + H2O2 f CatalaseFeIII + O2 + H2O (2) This scheme suggests that the iron heme enzyme (CatalaseFeIII) first forms reactive oxidants with H2O2 with a cation radical on the heme porphyrin ([CatalaseFeIVdO]•, compound I). Compound I then accepts an electron from H2O2 to form a nonradical compound ([CatalaseFeIVdO], compound II), resulting in oxidation of H2O2. Compound II can further accept a second electron to regenerate the iron heme enzyme. The catalytic H2O2 current is attributed to the reduction of compound I. In addition, it should be noted that the electrodes prepared from nonencapsulated catalase and solubilized catalase also display catalytic activity to H2O2, but the responses are much smaller and (35) Peroxidase in Chemistry and Biology; Everse, J., Everse, K. E., Grisham, M. B., Eds.; CRC Press: Boca Raton, FL, 1991; Vol. II.

Figure 5. Effect of the layer number of PE precursor (on the electrodes) on the (a) current response and (b) response time to H2O2. The film structure on the modified electrodes was (PAH/PSS)n/EL4/ PSS/PAH.

less stable than that observed for PE-encapsulated catalase. Figure 4 (curve b) shows the response of nonencapsulated catalase (PAH/PSS/Eo/PSS/PAH) in the solution containing the same concentration of H2O2. At -0.35 V, the catalytic current response (Icat ) IH2O2 - Ibackground) is 14.5 µA, which is ∼4.5 times lower than that observed for the PE-encapsulated catalase (65.3 µA). The higher response of the PE-encapsulated catalase may be due to the larger amount of enzyme immobilized on the electrode. However, the relative increase in the response is not proportional to the amount of enzyme deposited (∼9 times more enzyme is deposited when it is PE-encapsulated). This indicates that not all of the catalase molecules are in an electrocatalytic active state after PE encapsulation and immobilization on the electrode. One reason for this is that after PE encapsulation, the electron-transfer distance between the catalase molecules and the electrode surface increases. According to the electron transfer theory, 36 the electron tunneling rate constant (k′) is strongly dependent on the distance between the donor and acceptor (d, in this case is the separation between the electrode and the immobilized catalase),

k′ ) ko exp[-β(d - do)]

(3)

where ko is the rate constant at contact distance do, and β is the decay constant. The presence of the PE layers between the electrode and immobilized enzyme increases the distance between the enzyme and the electrode surface, which decreases the electrochemical activity of the whole electrode. This was verified by observing a decrease in the electrocatalytic activity of the electrode with increasing PE layer number between the enzyme and the electrode (see below). Influence of Precursor Film Thickness and Number of Catalase Multilayers on the Electrocatalytic Activity of the Electrode. Figure 5 shows the effect of the number of PE precursor layers (i.e., layers between the electrode and enzyme layers) on the electrocatalytic response of catalase to H2O2. It can be seen that adding one bilayer of precursor (n ) 1) does not decrease the current very much. However, with increasing (36) (a) Marcus, R. A.; Sutin, N. Biochim. Biophys. Acta 1985, 811, 265. (b) Closs, G.; Miller, J. Science 1988, 240, 440. (c) Wuttke, D.; Bjeerrum, M.; Winkler, J.; Gray, H. Science 1992, 256, 1007.

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precursor PE layer number, the current response to peroxide decreases significantly, and the electrode response time increases. Similar phenomena were reported by Bruening and co-workers when they examined the permeability of multilayered PAH/PSS films using Fe(CN)63- as an electrochemical probe.37 In that study,37 the current response of Fe(CN)63- decreased with increasing PE layer number on the electrode as a result of the charge-transfer resistance increase in the film, especially when the third PE bilayer was introduced. Keeping the number of precursor layers (e.g., PAH/PSS) constant but increasing the number of PE layers used to encapsulate the catalase crystals (e.g., EL6, EL8) resulted in the same trend (data not shown); that is, the current response decreased upon increasing the number of PE layers used to encapsulate the catalase microcrystals. These results suggest that under the conditions of having a stable biosensor composed of PE-encapsulated catalase, fewer layers of PE are needed in order to obtain a sensitive and fast response. We also investigated the electrocatalytic activity of PEencapsulated enzyme multilayer films. Although the amount of enzyme increases linearly with the enzyme layer (see earlier), the total electrode response for two layers of catalase microcrystals deposited onto the electrodes increased by only a further 30%, after which a plateau in the response was observed. In addition, the current response time to H2O2 increased, and the catalytic potential shifted to negative values, which indicate that the catalytic reaction became more difficult (i.e., less efficient). This is mainly attributed to the increasing difficulty for the substrate (H2O2) and product (O2) to diffuse through the film upon increasing the film thickness. Previous studies have also suggested that saturation of the enzymatic activity of enzyme/PE multilayer films is due to the substrate diffusion limitation and reduced penetration into the film.38 Another contributing factor is that the electrochemical activity of the whole electrode is not expected to increase linearly with the number of catalase layers. According to the electron transfer theory, with increasing distance from the electrode, the second layer of enzyme may not have its whole electrochemical activity. Our data is in excellent agreement with a recent study on the LbL assembly of myoglobin (Mb) and PSS, wherein Rusling and co-workers showed that a second Mb layer provides only a 30-40% electroactivity increase, with no further increase occurring upon deposition of additional Mb layers.16b Catalase Multilayer Films as H2O2 Biosensors. We then applied the PE-encapsulated catalase-modified electrodes to measure the concentration of H2O2. This was undertaken in a stirred system by applying a potential step of -0.30 V to the working electrode. Figure 6 shows the chronoamperometric response of a EL4-modified electrode when adding aliquots of 0.2 mM H2O2 in pH 7.0 PBS. It can be seen that the response is very quick, with an average response time of ∼2.62 s. The current response obtained (using the same electrode) is also reproducible, and the relative standard deviation for eight successive measurements is 8.2%. Figure 7 shows the linear dependence between the catalytic peak current and the concentration of H2O2. A linear range from 3.0 × 10-6 to 1.0 × 10-2 M H2O2 was observed. The linear (37) Harris, J. J.; Bruening, M. L. Langmuir 2000, 16, 2006. (38) Lvov, Y. In Protein Architecture: Interfacing Molecular Assemblies and Immobilization Biotechnology; Lvov, Y., Mo ¨hwald, H., Eds.; Marcel Dekker: New York, 2000.

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Figure 6. Chronoamperometric currents at -0.30 V as a function of time as a result of adding aliquots of 0.2 mM H2O2 in pH 7.0 PBS. Working electrode: PAH/PSS/EL4/PSS/PAH.

Figure 7. Linear calibration curve for determining H2O2. Working electrode: PAH/PSS/EL4/PSS/PAH.

Figure 8. Stability of the current response to H2O2 at (a) PAH/PSS/ PAH/Es-, (b) PAH/PSS/Eo-, and (c) PAH/PSS/EL4-modified electrodes in pH 7.0 PBS.

regression equation is log I (µA) ) 4.662 + 1.013 log C (M), with a correlation coefficient of 0.999. At a higher concentration of H2O2 (3.0 × 10-2 M), the response suddenly sharply increased and then reached a plateau (>7.0 × 10-2 M), which may be due to the denaturation of catalase in high concentrations of H2O2.33 Enzyme Film Stability. Figure 8 compares the current response stability of solubilized catalase (a), nonencapsulated catalase (b), and four-layer PE-encapsulated catalase (c) in pH 7.0 PBS to H2O2. The nonencapsulated catalase has the lowest stability, losing nearly all of its catalytic activity after 2 h as a result

of solubilization and desorption in pH 7.0 solution. In comparison, the solubilized catalase has the lowest response, but higher stability, with 30% of its original activity remaining after 120 min. The PE-encapsulated catalase has the highest stability. After 2 h and more than 10 measurements, the PE-encapsulated catalase retained 85% of its original activity. This decrease of the activity is probably due to the loss of loosely attached catalase crystals from the electrode surface. Further experiments showed that the PE-encapsulated catalase is also more stable than nonencapsulated catalase at high temperatures and when exposed to high-pH solution. For example, at 85 °C, the nonencapsulated catalase lost all of its activity, while the PE-encapsulated enzyme retained 30% of its original activity. In pH 9.0 solution, the nonencapsulated catalase showed 40% of its original activity, compared with 60% for the PE-encapsulated catalase. These results clearly demonstrate that a thin polymer coating of four layers (thickness of ∼5∼8 nm) does not only significantly increase the catalase loading amount onto surfaces (e.g., electrodes), but it also improves the stability of the enzyme at elevated temperature and in high-pH solutions. CONCLUSIONS We have reported the utilization of PE-encapsulated microcrystals as building blocks for the construction of biocatalytic films with tailored activity and high stability. The strategy employed provides a new route to prepare films of immobilized enzymes that may have potential applications in biotechnology. The significance of our approach is (i) A high density of enzyme in relatively thin films (few micrometers) is obtained, which leads to a sensitive biosensor with low diffusion barriers; (ii) The PE layers encapsulating the catalase can prevent enzyme molecules

directly contacting the electrode surface, thereby keeping their biospecific and electrochemical activities and allowing them to undergo facile electron-transfer reactions; (iii) Direct electron transfer between catalase and the electrode can be achieved without the aid of an electron mediator; (iv) The PE coatings prevent the leakage of the enzyme, thus increasing its stability, even at high temperature and in high pH solutions; and (v) The thickness and composition of the PE coatings can be controlled by varying the number and type of PE layers deposited, thus providing a simple way to control the permeability of the coating. Methods to improve the electron transfer between electrodes and PE-enzyme systems, for example, by infiltrating conductive gold nanoparticles and the use of enzyme microcrystal templates to construct microreactor systems for sequential biocatalytic reactions are currently in progress in our group. ACKNOWLEDGMENT We thank C. Pilz for assistance with ζ-potential measurements. T. Cassagneau is acknowledged for helpful discussions. H. Mo¨hwald is thanked for supporting this work within the MPIInterface section. This work was funded by the German Federal Ministry of Education, Science, Research and Technology (BMBF) as part of its Biofuture research program and the Australian Research Council under the Federation Fellowship and Discovery Project schemes.

Received for review January 8, 2003. Accepted March 26, 2003. AC0340049

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