Fabrication of Phytic Acid Sensor Based on Mixed Phytase− Lipid

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Fabrication of Phytic Acid Sensor Based on Mixed Phytase-Lipid Langmuir-Blodgett Films Luciano Caseli,*,† Marli L. Moraes,† Valtencir Zucolotto,† Marystela Ferreira,‡ Thatyane M. Nobre,§ Maria Elisabete D. Zaniquelli,§ Ubirajara P. Rodrigues Filho,| and Osvaldo N. Oliveira, Jr.† Grupo de Polı´meros, Instituto Fı´sica de Sa˜ o Carlos, UniVersidade de Sa˜ o Paulo (IFSC-USP), Sa˜ o Carlos, SP, Brazil, Faculdade de Cieˆ ncia e Tecnologia, UniVersidade Estadual Paulista (FCT-UNESP), Presidente Prudente, SP, Brazil, Departamento de Quı´mica, Faculdade de Filosofia, Cieˆ ncias e Letras de Ribeira˜ o Preto, UniVersidade de Sa˜ o Paulo (FFCLRP-USP), Ribeira˜ o Preto, SP, Brazil, and Instituto de Quı´mica de Sa˜ o Carlos, UniVersidade de Sa˜ o Paulo (IFSC-USP), Sa˜ o Carlos, SP, Brazil ReceiVed June 22, 2006. In Final Form: August 1, 2006 This paper reports the surface activity of phytase at the air-water interface, its interaction with lipid monolayers, and the construction of a new phytic acid biosensor on the basis of the Langmuir-Blodgett (LB) technique. Phytase was inserted in the subphase solution of dipalmitoylphosphatidylglycerol (DPPG) Langmuir monolayers, and its incorporation to the air-water interface was monitored with surface pressure measurements. Phytase was able to incorporate into DPPG monolayers even at high surface pressures, ca. 30 mN/m, under controlled ionic strength, pH, and temperature. Mixed Langmuir monolayers of phytase and DPPG were characterized by surface pressure-area and surface potential-area isotherms, and the presence of the enzyme provided an expansion in the monolayers (when compared to the pure lipid at the interface). The enzyme incorporation also led to significant changes in the equilibrium surface compressibility (in-plane elasticity), especially in liquid-expanded and liquid-condensed regions. The dynamic surface elasticity for phytase-containing interfaces was investigated using harmonic oscillation and axisymmetric drop shape analysis. The insertion of the enzyme at DPPG monolayers caused an increase in the dynamic surface elasticity at 30 mN m-1, indicating a strong interaction between the enzyme and lipid molecules at a high-surface packing. Langmuir-Blodgett (LB) films containing 35 layers of mixed phytase-DPPG were characterized by ultravioletvisible and fluorescence spectroscopy and crystal quartz microbalance nanogravimetry. The ability in detecting phytic acid was studied with voltammetric measurements.

Introduction Phytase is relevant for the poultry and food industry, for it is able to catalyze the hydrolysis of phytic acid (myo-inositol hexaphosphate), producing inorganic phosphate. Phytic acid is the common storage form of phosphorus that accounts for about 70% of the phosphate content in legume seeds, cereal grains, and beans;1,2 it is antinutritional, attaching to minerals, proteins, and starch, decreasing their absorption. Phytic acid is also antioxidant and anticarcinogenic, being important for the environmental phosphorus pollution. Sensors for phytic acid have been developed over the years,3-8 but less attention was given to biosensors based on phytase.9 For instance, Mak et al.10 developed an * Author to whom correspondence should be addressed. Tel/fax: + 55 16 3371 5365; e-mail: [email protected]. † Instituto Fı´sica de Sa ˜ o Carlos, Universidade de Sa˜o Paulo. ‡ Universidade Estadual Paulista. § Cie ˆ ncias e Letras de Ribeira˜o Preto, Universidade de Sa˜o Paulo. | Instituto de Quı´mica de Sa ˜ o Carlos, Universidade de Sa˜o Paulo. (1) Cosgrove, D. J. Their Chemistry, Biochemistry, and Phsiology; Elsevier: Amsterdam, 1980. (2) Raboy, N. R. Trends Plant Sci. 2001, 6 (10), 458-462. (3) Keeme, P. A.; Lommen, A.; De Jonge, L. H.; Van der Klis, J. D.; Jongbloded, A. W.; Mroz, Z.; Breynem, A. C. J. Agric. Food Chem. 1999, 47 (12), 51165121. (4) Graf, E.; Dintzis, F. R. Anal. Biochem. 1982, 119, 413-417. (5) Vaintraub, I. A.; Lapteva, N. A. Anal. Biochem. 1988, 175 (1), 227-230. (6) Talamond, P.; Doulbeau, S.; Rochette, I.; Guyot, J. P. J. Chromatogr., A 1988, 871 (1-2), 7-12. (7) Phillippy, B. Q.; Bland, J. M.; Evens, T. J. J. Agric. Food Chem. 2003, 51 (12), 350-353. (8) Wyss, M.; Brugger, R.; Kronenberger, A.; Re´my, R.; Fimbel, R.; Oesterhelt, G.; Lehmann, M.; Loong, A. P. G. M. Appl. EnViron. Microbiol. 1999, 65 (2), 367-373. (9) March, J. G.; Villacampa, A. I.; Grases, F. Anal. Chim. Acta 1995, 300, 269-72.

electrode based on polycarbamoyulsulfonate (PSS) hydrogel with co-immobilization of phytase and pyruvate oxidase coating platinum to detect phytic acid. Also, Pandey et al.11 suggested that the immobilization of phytase on polymers and solid supports could increase its thermal and kinetic stability. New methods to develop biosensors in organized systems at the nanometric scale have been developed in the past years, making use of liposomes,12 layer-by-layer,13 and LangmuirBlodgett (LB) films.14,15 The LB method, in particular, is advantageous for its intrinsic control of film thickness and molecular architecture, in addition to the suitability for designing artificial systems with biological functions, applicable in immunosensors,16-18 enzyme sensors,19-21 biomolecular microphotodiodes,22,23 and biocatalysis membranes.24-26 With the LB method, one may seek for appropriate lipids that interact (10) Mak, W. C.; Ng, Y. M.; Chiyui, C.; Kwong, W. K.; Renneberg, R. Biosens. Biolectron. 2004, 19, 1029-1035. (11) Pandey, A.; Szakacs, G.; Soccol, C. R.; Rodriguez-Leon, J. A. Soccol, V. T. Biores. Technol. 2001, 77, 203-214. (12) Liang J. F.; Li, Y. T. J. Pharm. Sci. 2000, 89 (8), 979-990. (13) Campas, M.; O’Sullivan, C. Anal. Lett. 2003, 36 (12), 2551-2569. (14) Troitsky, V. I.; Berzina T. S.; Pastorino, L.; Bernasconi E.; Nicolin, C. Nanotechnology 2003, 14, 597-602. (15) Girard-Egrot, A. P.; Godoy, S.; Blum, L. C. AdV. Colloid Interface Sci. 2005, 116, 205-225. (16) Tieke, B. AdV. Mater. 1991, 3 (11), 532-. (17) Sanches-Gonzalez, S.; Ruiz-Garcia, J.; Galvez-Ruiz, M. J. J. Colloid Interface Sci. 2003, 267 (2), 286-293. (18) Barraud, A.; Perrtot, H.; Billard, V.; Martelet, C.; Therasse, J. Biosens. Biolectron. 1993, 8 (1), 39-48. (19) Choi, J. W.; Bae, J. Y.; Min, J. H.; Cho, K. S.; Lee, W. H. Sens. Mater. 1996, 8 (8), 493-504. (20) Girard-Egrot, A. P.; Morelis, R. M.; Coultet, P. R. Thin Solid Films 1997, 292 (1-2), 282-289. (21) Tsuji, H.; Mitsubayashi, K. Electroanalysis 1997, 9 (2), 161-164.

10.1021/la061799g CCC: $33.50 © 2006 American Chemical Society Published on Web 09/02/2006

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strongly with the biomolecules, which has led to a host of studies of mixed lipid-protein monolayers at the air-water interface.27-29 Moreover, the molecular architecture inherent in Langmuir monolayers or LB films may be exploited in controlling the enzyme activity, which has been shown to depend on the supramolecular structure of protein-lipid interfaces.28,30 In this study, we investigate the adsorption of phytase at airwater interfaces, with emphasis on its interaction with a charged phospholipid, dipalmitoyl phosphatidyl glycerol (DPPG). We show that the strong adsorption allows one to transfer phytaseDPPG mixed monolayers onto solid substrates, which can then be used in biosensors for phytic acid detection using cyclic voltammograms. Materials and Methods DPPG (Sigma) and phytase from Aspergillus ficuum (EC: 3.1.3.8, 1.1 units/mg, Sigma), NaCl (Merck, 99%), NiCl2‚6H2O (Reagen, 97%), acetic acid (Sigma), phytic acid (Sigma), and sodium acetate (Merck, 98%) were used as received. Water was supplied by a Milli-Q system (resistivity ) 18.2 MΩcm). The Langmuir monolayers were prepared on a Nima Langmuir trough (model 601M) or KSV (model 5000), housed in a class 10 000 clean room, equipped with a Wilhelmy plate made of a filter paper and a Kelvin probe to measure surface potential. Langmuir monolayers of DPPG were obtained by spreading a chloroform (Sigma, HPLC grade) solution of DPPG on the aqueous subphase, made of 0.01 mol L-1 acetate buffer (pH 5.5), 0.1 mol L-1 NaCl, and 1.0 × 10-3 mol L-1 NiCl2. Mixed monolayers of phytase and DPPG were prepared by injecting a protein aqueous solution in the subphase, just below the interface, after the DPPG spreading. Various proportions of phytase/DPPG were tested, using different injection volumes of a 2.5 mg mL-1 phytase solution (prepared in 0.01 mol/L acetate buffer, pH 5.5). Surface pressurearea (π-A) and surface potential-area (∆V-A) isotherms were obtained with a monolayer compression rate of 10 cm2‚min-1, with the subphase at either 20 °C or 25 °C. For control, the air-water interface on phytase solution, without DPPG, was compressed and variation of surface pressure and surface potential was measured. Curves for the adsorption kinetics were obtained by injecting the phytase solution under the interface and by following the increase in surface pressure with time, both with a DPPG monolayer at the interface or for the pure water subphase. Several initial surface pressures were tested, producing several lipid packing conditions, which were obtained by spreading distinct volumes of DPPG solution. Dynamic elasticity measurements of DPPG, phytase, and mixed DPPG-phytase monolayers were carried out using the harmonic oscillation and axisymmetric drop analysis method. The latter was carried out using an OCA-20 from Dataphysics Instruments GmvH, Germany, with oscillating drop accessory ODG-20. A drop of liquid was formed from a syringe into a thermostated optical glass cuvette containing water in the bottom to avoid drop evaporation. The drop oscillation began after the surface tension reached a constant value. A piezo actuator device, connected to a function generator and located above the needle, produced a sinusoidal movement of the drop in a given frequency. The images were recorded with a video camera (22) Choi, J. W.; Jung, G. Y.; Oh, S. Y.; Lee, W. H.; Lee, W. H.; Shin. D. M. Thin Solid Films 1996, 285, 876-878. (23) Choi, J. W.; Nam, Y. S.; Cho, K. S.; Lee, W. H.; Park, S. Y.; Fujihira, M. J. Ind. Eng. Chem. 2003, 9 (1), 31-36. (24) Reichert, W. M.; Bruckner, C. J.; Joseph, J. Thin Solid Films 1987, 152, 345-376. (25) Caseli, L.; Zaniquelli, M. E. D.; Furriel, R. P. M.; Leone, F. A.; Colloids Surf., B 2002, 25 (2), 119-128. (26) Caseli, L.; Furriel, R. P. M.; de Andrade, J. F.; Leone, F. A.; Zaniquelli, M. E. D. J. Colloid Interface Sci. 2004, 275 (1), 123-130. (27) Wiedmann, T. S.; Jordan, K. R. Langmuir 1991, 7, 318-322. (28) Maggio, B.; Rosseti, B.; Borioli, G. A.; Fanani, M. L. Braz. J. Med. Biol. Res. 2005, 38, 1735-1748. (29) Vollhart, D.; Fainerman, V. B. AdV. Colloid Interface Sci. 2000, 86 (12), 103-151. (30) Caseli, L.; Oliveira, R. G.; Masui, D. C.; Furriel, R. P. M.; Leone, F. A.; Maggio B.; Zaniquelli, M. E. D. Langmuir 2005, 21 (9), 4090-4095.

Caseli et al. with a minimum of 200 frames per second. At the end of the experiment, the software retrieved the images and calculated the change in area and corresponding changes is surface tension for each cycle. Using Fourier transform analysis, the dilational elasticity and phase angle were determined. For these experiments, aliquots of phytase (acetate buffer pH ) 5.5; Na+ 0.1 mol L-1 and Ni2+ 10-3 mol L-1) solutions in three concentrations (0.5, 16.7, and 167 mg L-1) were used to form a drop in a surface area of ca. 30 mm2. Periodic oscillations with amplitude of 0.1 mm in the frequency range between 0.7 and 1.5 Hz were imposed to the drop at a temperature of 25 ( 0.1 °C. For the dynamic elasticity measurements with the monolayer, a solution of 10-4 mol L-1 of DPPG was gently touched with a Hamilton syringe on the surface of the drop, which was formed with the buffer solution, with or without phytase. Two surface pressures were investigated, namely, 15 and 30 mN m-1. The transfer of the mixed DPPG/phytase monolayers onto different solid supports (see below) was done at a surface pressure of 30 mN m-1. The DPPG monolayer formed on a Na+ 0.1 mol L-1, Ni2+ 1.0 × 10-3 mol L-1, 0.1 mol L-1 acetate buffer was compressed until reaching the surface pressure of 20 mN m-1, after which it was left to stabilize for ca. 10 min. An aliquot of 100 µL of 2.5 mg mL-1 phytase solution, prepared in 0.1 mol L-1 acetate buffer, pH 5.5, was then injected into the subphase, a few millimeters below the airwater interface. For the subphase volume of 500 mL, the final enzyme concentration was 0.50 mg L-1. With the injection of the phytase solution, the surface pressure increased to 23-24 mN m-1, and the monolayer was again compressed to 30 mN m-1. After 20 min at this pressure to check monolayer stability, the solid support, which was already immersed in the subphase, was taken off from the subphase. Several immersions and withdrawals were performed to produce Y-type, 35-layer LB films from mixed DPPG-phytase. The dipping speed was 2.0 mm‚min-1, and the transfer ratio was between 0.8 and 1.0. The formation of LB films was confirmed with UV-vis (Hitachi U 2001) and fluorescence (Shimadzu RF-5301PC) spectroscopy, in addition to nanogravimetry analysis with a quartz crystal microbalance (QCM, Stanford Research Systems Inc). The substrates used were quartz plates and optical glass for UV-vis and fluorescence spectroscopy measurements, respectively, while for QCM nanogravimetry we employed AT-cut quartz crystal coated with Au, Stanford Research Systems INC, fundamental frequency of ca. 5 MHz. Cyclic voltammograms in the range between -500 mV and 500 mV were obtained with LB films deposited onto ITO using a potentiostat Autolab PGSTAT30 and a three-electrode, 15-mL electrochemical cell. The reference electrode was Hg/Hg2SO4/K2SO4(sat.) (MSE); a 1.5 cm2 platinum foil was used as an auxiliary electrode, and the working electrode was a 35-layer LB film from mixed DPPG-phytase onto ITO. The experiments were conducted in acetate buffer solution (at pH 5.5) at room temperature (24-26 °C). Cyclic voltammograms were obtained in the presence of phytic acid at concentrations varying from 6 to 30 mmol L-1 and at a sweeping ratio of 25 mV s-1. After the measurements, the films tested were exhaustively washed with the electrolytic solution and the reproducibility was investigated. The LB films were stable in all experiments.

Results and Discussion Adsorption of Phytase at the Air-Liquid Interface. Figure 1 (a and b) shows the kinetics of adsorption for phytase on the air-water interface. As the surface pressure increases immediately after enzyme injection in the subphase, the induction time is practically zero (Figure 1a). In fact, the ionic strength of 0.1 mol L-1 was used to facilitate enzyme adsorption,31 as we tried ionic strengths of 0.01, 0.05, and 0.1 mol L-1, and only the latter resulted in a good enzyme adsorption at the air-water interface. At a bare interface (without lipids), the surface pressure reached a stable value in less than 10 min after the phytase injection, with (31) Britt, D. W.; Mobius, D.; Hlady V. Phys. Chem. Chem. Phys. 2000, 2 (20), 4594-4599.

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Figure 1. Adsorption kinetics for phytase (100 µL of 2.5 mg‚mL-1) inserted in acetate buffer (0.01 mol‚L-1), Na+ 0.1 mol‚L-1, Ni2+ 10-3 mol‚L-1, pH 5.5, corrected with acetic acid, total volume of 500 mL. Final phytase concentration in the subphase is 0.50 mg‚L-1. For adsorption at DPPG monolayer, the initial surface pressure is 0 mN‚m-1 in a phospholipid molecular area of 105 Å2. (a) Curve relating the increase of surface pressure with time. (b) Corresponding kinetic plots according to eq 1. Temperature ) 20 °C.

a change in pressure (∆π ) πf - πi) in which πi is the initial surface pressure and πf is the final surface pressure of approximately 11 mN m-1. When a DPPG monolayer was present at the interface, but with a low surface density corresponding to a null surface pressure, equilibrium was attained in less than 6 min with ∆π ) 4.5 mN m-1. The area per lipid molecule used was 105 Å2, at which the surface pressure starts to increase upon further compression. Therefore, the surface activity of the enzyme decreases in the presence of the phospholipids, which has also been observed for other globular,32 nonglobular,33 and anchored proteins.34 The kinetics data for phytase adsorption were treated with the approach suggested by Magett-Dana35 who applied a first-order equation36 to analyze adsorption phenomena of proteins on liquid interfaces.

ln[(πf - πt)/(πf - πi)] ) -kt

(1)

where πt is the surface pressure at a time t and k is the rate (32) Li, J. R.; Rosilo, V.; Boissonnade, M. M.; Baskin, A. Colloids Surf., B 2003, 29 (1) 13-20. (33) Del Boca, M.; Caputo, B. L.; Maggio, B.; Borioli, G. A. J. Colloid Interface Sci. 2005, 287, 80-84.

constant. According to Figure 1b, two rate constants are identified, and the change in slope coincides with the time where the surface pressure begins to stabilize, that is, 8.3 and 6.4 min for phytase adsorbing on a bare interface and on a DPPG monolayer, respectively. These rate constants are higher than those reported for other proteins,35 usually in the range of 10-2-10-3 min-1 indicating a strong surface activity for phytase under the conditions employed. The isoelectric point for extracellular phytase from Aspergillus ficuum, which is a glycoprotein with molecular weight of 85100-KDa,37,38 is ca. 4.5,38 and therefore the enzyme at pH 5.5 used in this work is negatively charged. Its activity at 25 °C should be preserved, because in thermal inactivation studies it has been shown that this enzyme is capable of retaining activity even after treatment at 60 °C.39 Apparently, the presence of a lipid monolayer provides a faster adsorption of the enzyme to the interface, reflecting the high adsorption rate in the first step (0.473 min-1), and therefore a favorable interaction of the enzyme with the lipid. Probably, this interaction is electrostatically stabilized by Ni2+ ions, considering that both lipid and protein are negatively charged. A zwitterionic phospholipid, such as dipalmitoyl phosphatidyl choline (DPPC), was not used because multilayer LB films with more than two layers could not be transferred with mixed monolayers formed by phytase and such lipids. The change in surface pressure because of penetration of enzyme in DPPG monolayers is lower than for the enzyme adsorbed on a bare air/water interface. This indicates that the stabilization is attained because of interaction between the enzyme and the lipid, which can provide several conformations, geometrical shapes, amphiphilicity, and thermodynamic properties.28 Therefore, the adsorption of proteins to the interface constitutes a major perturbing factor for the Langmuir monolayer both in terms of surface topography and dynamics, which will reflect in the equilibrium surface pressure. Obviously, for a protein adsorbed alone at the interface, there are no lipid-polypeptide interactions and the conformation properties assumed for the adsorbed protein are different from those observed when interacting with lipids. The suggested mechanism for protein adsorption comprises two steps. Usually, the first step follows a previous phase of diffusion to which an “induction time” may be associated, where no change in surface pressure is detected. Assuming a zero induction time because of the rapid diffusion of phytase to the air-liquid interface, the first step corresponds to continuous penetration of the enzyme in the lipid monolayer. The second step, after the slope change, involves a slow increase in surface pressure with a plateau corresponding to the equilibrium. In fact, complete surface pressure stabilization can take long time, owing to desorption and readsorption and continuous conformation changes of the protein adsorbed. The higher rate constant obtained for phytase adsorbing at a bare interface (2.017 min-1) can be ascribed to several factors, including desorption and readsorption processes and mainly surface-induced denaturation. Figure 2 shows the increase in surface pressure (∆π) because of the protein adsorption on a DPPG monolayer and reveals that ∆π decreases with initial surface pressure of the experiment (πi). A high-surface packing hampers the protein penetration, con(34) Caseli, L.; Zaniquelli, M. E. D.; Furriel, R. P. M.; Leone, F. A. Colloids Surf., B 2003, 30 (4), 273-282. (35) Maget-Dana, R. Biochim. Biophys. Acta 1999, 1462, 109-140. (36) Graham, D. E.; Philips, M. C. J. Colloid Interface Sci. 1979, 70, 403414. (37) Liu, B.-L.; Rafiq, A.; Tzeng, Y.-M.; Rob, A. Enzyme Microb. Technol. 1998, 22, 415-424. (38) Ullah, A. H.; Gibson, D. M. Prep. Biochem. 1987, 17 (1), 63-91. (39) Howson, S. J.; Davis, R. P. Enzyme Microb. Technol. 1983, 5, 377-382.

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Figure 2. Increase of surface pressure caused by insertion of phytase solution (100 µL of 2,5 mg‚mL-1) in a subphase with acetate buffer (0.01 mol‚L-1), Na+ 0.1 mol‚L-1, Ni2+ 10-3 mol‚L-1, pH 5.5, corrected with acetic acid, total volume of 500 mL, for DPPG monolayers at several initial surface pressures. Final phytase concentration is 0.50 mg‚L-1. The extrapolation of linear fit to nil surface pressure indicates the exclusion surface pressure.

Caseli et al.

Figure 4. Surface pressure-area isotherms for DPPG monolayers on subphase of pure water (pH 5.5) and buffer (acetate 0.01 mol‚L-1, Na+ 0.1 mol‚L-1, Ni2+ 10-3 mol‚L-1, pH 5.5, corrected with acetic acid). Total volume of trough: 500 mL. The temperatures are indicated in the inset. The area per molecule was considered as all enzyme molecules was adsorbed at the air-water interface.

sistent with the literature.35 ∆π decreases linearly with πi, and the extrapolation to a null ∆π defines the exclusion surface pressure35,40,41 at which it is no longer possible for the protein to penetrate the lipid monolayer. At 20 °C, the exclusion surface pressure is 27.5 mN m-1, increasing to 40.0 mN m-1 at 25 °C. Thus, the thermal motion of DPPG chains favors protein incorporation, which may occur even at surface pressures higher than that corresponding to the lateral pressure for bilayers42 such as liposomes or cell membranes, that is, 30-35 mN m-1 at room temperature. Figure 3 shows the surface pressure-area isotherm when phytase is spread on the interface and a compression experiment is performed. On pure water, the surface pressure increases up to 10 mN m-1, while a pressure of 27 mN m-1 is reached on

the buffer. Hysteresis appears in the compression-decompression cycles, probably because of irreversible aggregation of enzyme molecules. Subsequent compressions lead to identical isotherms, with the extrapolated area (taken in the second compression as the area at which the steepest part of the curve is extrapolated to zero pressure) about 930 Å2, reasonable for a protein with a molecular weight of 85-100-KDa.37,38 In calculating the area per molecule, we assumed all phytase molecules to be at the interface. Since phytase is a soluble surfactant, it is distributed between the interface and across the bulk of the subphase, though at a high ionic strength a large partition at the interface is expected. Therefore, the area values should be understood as an extrapolation. The surface pressure-area (π-A) isotherms for DPPG monolayers shown in Figure 4 are similar to those from the literature.43 For pure water, there is a liquid-expanded to liquidcondensed transition in the range from 95 to 60 Å2 per molecule. The transition (assumed here arbitrarily at a molecular area of 65 Å2, taken for comparison) occurs at 2.2 mN m-1 and 12.9 mN m-1 for 20 and 25 °C, respectively, in agreement with Grigoriev et al.44 The minimum area changed slightly from 53.9 Å2 to 55.9Å2 when the temperature increased from 20 to 25 °C. When the buffer (acetate/acetic acid 0.1 mol L-1, NaCl 0.1 mol L-1, and Ni2+ 10-3 mol L-1) was employed, the transition occurred at higher pressures, namely, 6.4 and 19.2 mN m-1 for 20 and 25 °C, respectively. When phytase was incorporated in the DPPG monolayer, the surface pressure-area (π-A) isotherm was shifted, as shown in Figure 5. In this case, the enzyme was incorporated by injection into the subphase with the DPPG monolayer at an initial pressure of 0 mN m-1 and was allowed to reach equilibrium according to the adsorption kinetics in Figure 1 before compression started. The shift in the isotherm increased with increasing amounts of enzyme injected into the subphase (see Figure 6). Phytase incorporation also caused changes in the monolayer compressibility. Indeed, the equilibrium surface compressibility modulus,

(40) Brockman, H. Curr. Opin. Struct. Biol. 1999, 9, 438-443. (41) Ronzon, F.; Desbat, B.; Chauvet, J. P.; Roux, B. Colloids Surf., B 2002, 23 (4), 365-373. (42) Marsh, M. Biochim. Biophys. Acta 1996, 1286, 183-223.

(43) Xiao-Feng, X.; Fu, W.; Mengsu, Y.; Sen-Fang, S. Colloids Surf., B 2004, 39, 105-112. (44) Grigoriev, D.; Miller, R.; Wu¨stneck, R.; Wu¨stneck, N.; Pison, U.; Mo¨hwald, H. J. Phys. Chem. B 2003, 107, 14283-14288.

Figure 3. Surface pressure-area isotherms, showing compression and decompression cycles, for phytase monolayers spread on subphase of buffer (acetate 0.01 mol‚L-1, Na+ 0.1 mol‚L-1, Ni2+ 10-3 mol‚L-1, pH 5.5, corrected with acetic acid). Total volume of trough: 500 mL. Temperature: 25 °C. Insert: Compression isotherm on pure water.

Fabrication of Phytic Acid Sensor

Figure 5. Surface pressure-area isotherms for mixed DPPG monolayers on buffer subphase (acetate 0.01 mol‚L-1, Na+ 0.1 mol‚L-1, Ni2+ 10-3 mol‚L-1, pH 5.5, corrected with acetic acid). Total volume of trough: 500 mL. Temperature: 25 °C. Phytase concentrations are indicated in the inset.

Figure 6. Variation of surface pressure for DPPG monolayers (squares: at a fixed area of 90 Å2), lipid molecular area (triangles: at a fixed surface pressure of 30 mN‚m-1), and surface compressional modulus (circles: at a fixed surface pressure of 30 mN‚m-1) as a function of phytase concentration.

taken as -A(∂π/∂A)45 and also known as in-plane elasticity,46 decreased with phytase incorporation, as shown in Figure 6. This result is typical of proteins incorporated into lipid monolayers.30,47 Figure 7 shows surface potential-area isotherms that are practically the same for DPPG on pure water or on the buffer, in agreement with the literature.48 The initial potential was negative because of the partial ionization of DPPG polar heads, and the potential started to increase at a critical area between 90 and 95 Å2 until reaching ca. 200 mV for the monolayer in the condensed phase. When phytase was inserted, the initial surface potential increased to zero, because of either an enzyme dipole moment or a rearrangement of DPPG molecules induced by adsorption of phytase. The inset in Figure 7 shows that phytase on its own leads to a relatively large surface potential (220 mV), which indicates that their dipole moments must contribute (45) Davies, J. T.; Rideal, E. K. Interfacial Phenomen, 2nd ed.; Academic Press: New York, 1963; Chapter 5, p 265. (46) Smaby, J. M.; Kulkarni, V. S.; Momsen, M.; Brown, R. E. Biophys. J. 1996, 70 (2), 868-877. (47) Yin, F.; Shin, H.-K.; Kwon, Y.-S. Biosens. Bioelectron. 2005, 21, 21-29. (48) Borissevitch, G. P.; Tabak, M.; Oliveira, O. N., Jr. Biochim. Biophys. Acta 1996, 1278, 12-18.

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Figure 7. Surface potential-area isotherms for DPPG on buffer pH 5.5, without (circles) and with phytase 0.5 µg‚mL-1 (squares). The inset shows the surface potential-area isotherms for pure enzyme at the interface.

significantly. However, under maximum compression the surface potential was only 20-30 mV higher than for pure DPPG, that is, either enzyme molecules must have been expelled from the air-water interface, similarly to other systems involving mixtures of protein and lipids,49 or the charged polypeptide has a strong influence on the surface potential amplitude because of the electrical double layer formed at the interface. Rheological Properties of Phytase-Containing Interfaces. When enzyme molecules adsorb on monolayers, the rheological properties of the latter are affected, making it difficult to investigate the direct influence of the enzyme. A possible alternative is the use of dynamic measurements, such as the harmonic oscillation and axisymmetric drop analysis method, already proven suitable to other monolayer studies.50 Of particular relevance is to understand how phytase may affect the transfer of the monolayer onto solid substrates. For DPPG monolayers, the equilibrium properties such as surface pressure, area per molecule, and surface compressibility vary with phytase concentration up to 0.5 µg‚mL-1, above which no further changes occur and the amount of phytase adsorbed no longer increases. Figure 8 shows the dynamic surface elasticity for solutions of phytase in three concentrations, where the lowest one is the same used in the Langmuir trough, 0.5 µg‚mL-1. The elasticity increased with phytase concentration, and with surface pressure, indicating a fast phytase adsorption-desorption and confirming the high enzyme affinity for the interface. The phase angles (φ) are always non-negligible, usually lower than 10°, and therefore the imaginary contribution to elasticity must be considered, especially for low concentrations such as 0.5 µg‚mL-1. The imaginary part of the dynamic surface elasticity is related to the surface dilatational viscosity, which depends on the phase angle, while the real part is associated with the pure elastic component.51 One concludes that viscous effects contribute to the surface elasticity as phytase was adsorbed. Table 1 shows the elasticity for DPPG monolayers at 15 and 30 mN m-1. The first pressure was chosen because it is just above the phase transition, while the second is close to the deposition value for LB films employed here. (49) Xicohtencatl-Cortes, J.; Mas-Oliva, J.; Castillo, R. J. Phys. Chem. B 2004, 108, 7307-7315. (50) Cornec, M.; Narsimham, G. Langmuir 2000, 16, 1216-1225. (51) Benjamins, J.; de Feijter, J. A.; Evans, M. T. A.; Graham, E. E.; Phillips, M. J. C. Faraday Discuss. 1975, 59, 218-229.

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Figure 8. Surface elasticity for a drop (about 50 µL) of phytase solution. Oscilation frequency: 1.025 Hz, amplitude: 0.1 mm (relative area variation of 5.5%). Table 1. Dynamic Surface Elasticity for DPPG (Spread on a Drop of Buffer pH 5.5 Solution of about 50 µL, with or without Phytase)a DPPG surface pressure (mN‚m-1) elasticity (mN‚m-1) f

15 77.4 4.3°

30 123.8 5.2°

DPPG + phytase (0.5 µg‚mL-1) 15 28.1 3.2°

30 246.5 1.6°

a φ represents the phase angle. Oscilation frequency: 1.025 Hz, amplitude: 0.1 mm.

The phase angles were low, indicating a small contribution of viscous effects. The elasticity for pure or mixed DPPG was higher at 30 mN m-1, as expected since a lower compressibility is obtained in equilibrium (in Langmuir monolayers, as depicted in surface pressure-area isotherms). The presence of phytase caused the dynamic elasticity to decrease considerably at 15 mN m-1, in which the elasticity for the DPPG/phytase monolayer was only one-third of that for pure DPPG. The latter suggested that phytase penetrated into the monolayer, affecting the DPPG packing and causing the monolayer to be more fluid. On the other hand, the elasticity for the mixed DPPG/phytase monolayer is twice the value for the pure DPPG monolayer at 30 mN m-1, which indicates that the film becomes less fluid upon phytase adsorption on the DPPG monolayer. This imparts a higher ability to restore a surface pressure condition when submitted to a deformation. This result may be explained by an increase of DPPG molecular packing in the presence of phytase. The process of enzyme adsorption is faster than the frequency imposed to the oscillating drop. Hence, it is likely that the relaxation process due to area variations should be related to protein molecular rearrangements or protein accommodation at the interface rather than to diffusion or adsorption process In contrast to the increase in surface elasticity in the oscillating drop caused by phytase, its presence led to a decrease in compressional modulus for a Langmuir monolayer (see Figure 6). Since surface elasticity in the drop is a nonequilibrium phenomenon, dynamic processes involving phytase penetration in the first steps of adsorption may be undetected in surface pressure-area curves. Moreover, for a monolayer in a Langmuir trough, the adsorption of phytase occurs at a low state of packing (0 mN m-1), and the surface pressure of 30 mN m-1 is attained by compression. On the other hand, in the oscillating drop, DPPG is spread in a phytase-containing interface, after the formation of a phytase solution drop. Expanding and compressing the drop will form dynamically new interfaces, and processes of adsorption/

Figure 9. Nanogravimetry for LB films of phytase and DPPG transferred from a mixed enzyme-lipid monolayer at a surface pressure of 30 mN‚m-1 at 25 °C, indicating the oscillation frequency variation (∆f) and the gain of mass.

desorption of phytase may occur. The high dynamic surface elasticity for mixed phytase-DPPG monolayers may be explained by the rapid adsorption of phytase to the interface, at a high state of packing, leading to a pistonlike effect.52 During the expansion/ compression process of the drop, with DPPG monolayers on the air-water interface, the enzyme penetrates rapidly, thus compressing the monolayer and leading to larger changes in surface pressure. As elasticity is defined as dγ/d ln A, a sudden compression of DPPG monolayer because of fast penetration of the protein yields higher surface elasticity. As the compressional modulus is smaller for equilibrium measurements (surface pressure-area isotherms), it is possible that after phytase penetration a rearrangement of protein molecules at the monolayer occurs because of relaxation. This confirms the high affinity of phytase for DPPG interfaces, even at high surface pressures, which is promising for transfer of mixed DPPGphytase LB films to be employed as sensors, as we show below. LB Films of Phytase and DPPG. The mixed phytase (0.5 µg‚mL-1)/DPPG monolayer could be compressed to surface pressures of 30 mN m-1 and transferred onto solid supports, namely, quartz crystal, quartz plate, glass, or ITO. Y-type LB films with 35 layers were deposited with transfer ratios of 0.98 ( 0.06 in the upstrokes and 0.8 ( 0.2 in the downstrokes. Using a quartz crystal microbalance, the average gain of mass (including lipid and phytase) was calculated with the Sauerbrey equation53 to be 69 ( 6 ng in the upstrokes and 40 ( 10 ng in the downstrokes. These values are consistent with the higher transfer ratios for the upstrokes. Furthermore, the uniform deposition was confirmed by an almost linear decrease in frequency with the number of deposited layers in Figure 9. It is important to mention, nevertheless, that the direct correlation between frequency change and mass is a rough approximation since the lipid-protein layers exhibit viscous effects (they are not perfectly elastic). Figure 10 shows the UV-vis spectra for the LB films, featuring an absorption peak at 275 nm, corresponding to a π-π* transition in the polypeptide. Since the films were very thin, at least 11 layers had to be deposited to obtain measurable signals. The absorbance increased linearly with the number of layers with a change in optical density (∆o) of 0.025 per layer. The LB films also exhibited fluorescence emission at 330 nm (with excitation at 295 nm) owing to excitation of tryptophan groups of phytase, as shown in Figure 11. The fluorescence signal increased with (52) Caseli, L.; Masui, D. C.; Furriel, R. P. M.; Leone, F. A.; Zaniquelli, M. E. D. Colloids Surf., B 2005, 46 (4), 248-254. (53) Sauerbrey, G. Z. Phys. 1959, 155, 206-222.

Fabrication of Phytic Acid Sensor

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Figure 10. UV-vis spectra for LB films of phytase and DPPG transferred from a mixed enzyme-lipid monolayer at a surface pressure of 30 mN‚m-1 at 25 °C. The signal was multiplied by 5 in absorbance scale. The inset shows the linear evolution of absorbance at 275 nm with number of layers.

Figure 12. Cyclic voltammograms (A) for a 35-layer mixed phytase-DPPG LB film in acetate buffer solution in the presence of phytic acid at concentrations of 6, 30, and 50 mmol‚L-1. Figure B shows the electrochemical response to pure DPPG LB films in acetate buffer solution (__) or in phytic acid solution 50 mmol‚L-1 (---). Figure C shows the electrochemical response to bare ITO to buffer (__) and in phytic acid solution 50 mmol‚L-1 (---). Scan rate: 25 mVs-1.

the number of deposited layers, again confirming the transfer of phytase to the LB films. As has been extensively discussed in the literature,54 immobilized enzymes may be used in biosensors, provided that their activity can be preserved. For phytase, in particular, we may envisage biosensors with molecular recognition capability toward phytic acid. This was confirmed using cyclic voltammetry, with ITO electrodes coated with DPPG/phytase LB films. Figure 12A shows voltammograms obtained with a scan rate of 25 mV/s in an acetate buffer solution in the range between -500 mV and 500 mV. It is readily seen that the LB films exhibit a different electrochemical response, compared to bare ITO or for pure DPPG LB films, both in the acetate buffer solution or in a phytic acid solution 5 × 10-2 mol L-1 (Figure 12B).The bare ITO substrate has no electrochemical response in the potential window used here. The oxidation/reduction peak appears at ca. 180-150 mV and -150-260 mV, respectively, being particularly well defined for 50 mmol L-1. The oxidation/reduction peaks were shifted depending on the phytic acid concentration. For instance, the reduction peak shifted from -290 mV to -220 mV when the concentration was varied from 30 to 50 mmol L-1. The shifts are due to the formation of O2 (which is denoted by the increase

in current for a potential above 550 mV) with the increase of the phytic acid concentration. The cathodic processes appearing in the voltammogram are due to reduction of both the O2 generated and the phytic acid. With low phytic acid concentration, the anodic and cathodic peaks are not well defined, but a shoulder due to the acid oxidation appears for a concentration of 30 mmol L-1, which is accompanied by a large oxidation current above 450 mV because of O2 generation. For 50 mmol L-1 of phytic acid, the anodic peaks are well defined while the two cathodic processes appear as a peak and shoulder. Detection of phytic acid occurs with the enzyme catalyzing the reaction with phytic acid, thus generating inositol and phosphate ions. In subsidiary experiments, we observed that both inositol and phosphate affect the electrochemical response of the LB films in the presence of phytic acid, but further studies are necessary to elucidate the role of each of these two chemical groups. Taking the oxidation peak as reference, the sensitivity of the LB immobilized phytase sensor toward phytic acid is 25.2 nA‚cm-2 mol-1‚L. This sensitivity is sufficient to detect phytic acid in seed of grains, legumes, and nuts.55 Mak et al.10 have found a linear response from 0.2 to a limit of 2 mmol L-1, with a current ranging from 5 to 52 nA. In our work, the linear response extended to a wider range, from 6 to 50 mmol L-1. We noted no changes in performance of the sensor even after 1 month of use, which means that phytase preserved its activity and the LB films were stable. Also, good reproducibility of the cyclic voltammetry results presented here was obtained by repeating

(54) Schwinte´, P.; Voegel, J.-C.; Picart, C.; Schaaf, Y.; Szalontai, B. J. Phys. Chem. B 2001, 105, 11906-11916.

51.

Figure 11. Fluorescence spectra for LB films of phytase and DPPG transferred from a mixed enzyme-lipid monolayer at a surface pressure of 30 mN‚m-1 at 25 °C. Excitation at 280 nm.

(55) Reddy, N. R. Food Phytate; CRC Press: Boca Raton, FL, 2002; pp 25-

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the measurements three times with three distinct LB films containing phytase. Other strategies have been used for sensing phytic acid, such as nanofilms with TiO2 nanoparticle phytates formed on gold electrode surfaces by directed assembly methods56,57 and use of enzymes immobilized in hydrogel.10 Using the LB technique, on the other hand, may be particularly interesting because the enzyme was immobilized in a highly organized system. Here, we have shown the viability of transferring LB films containing phytase, whose activity was preserved for at least 1 month, thus allowing detection of phytic acid. As far as the sensing ability is concerned though, this work was aimed as a proof of concept with no attempts to optimize the biosensor performance. Optimization

of the sensitivity and investigation of selectivity of phytasecontaining biosensors are now underway, in which amperometric measurements rather than cyclic voltammetry are being used.

(56) McKenzie, K. J.; Marken, F.; Gao, X.; Tsang, S. C.; Tam, K. Y. Eletrochem. Commun. 2003, 5, 286-291. (57) Paddon, C. A.; Marken, F. Electrochem. Commun. 2004, 6, 1249-1253.

Acknowledgment. The authors are grateful to FAPESP, CNPq, and IMMP/MCT (Brazil) for the financial assistance.

Conclusions We demonstrated that phytase can adsorb onto Langmuir films of DPPG, forming stable mixed monolayers. Dynamic and equilibrium properties of mixed films were markedly changed with penetration of the enzyme, indicating a significant affinity of phytase with DPPG, even at high states of packing. The nanostructured and nano-organized system with phytase incorporated was able to provide a good phytic acid sensor at room temperature.

LA061799G