Spongiform Immobilization Architecture of Ionotropy Polymer Hydrogel

Jun 11, 2004 - Department of Chemistry, Hong Kong Baptist University, Kowloon Tong, .... immobilization architecture is that the apparent enzyme activ...
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Anal. Chem. 2004, 76, 4279-4285

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Spongiform Immobilization Architecture of Ionotropy Polymer Hydrogel Coentrapping Alcohol Oxidase and Horseradish Peroxidase with Octadecylsilica for Optical Biosensing Alcohol in Organic Solvent Xiao Jun Wu and Martin M. F. Choi*

Department of Chemistry, Hong Kong Baptist University, Kowloon Tong, Hong Kong SAR, P.R. China

An organic-phase optical alcohol biosensor consisting of alcohol oxidase and horseradish peroxidase coimmobilized in a spongiform hydrogel matrix of hydroxethyl carboxymethyl cellulose, an adduct of 3-methoxy-4-ethoxy benzaldehyde, 4-tert-butylpyridinium acetohydrazone, silica gel particles, and octadecylsilica particles in conjunction with an optical oxygen transducer has been successfully fabricated. The novel enzyme entrapment structure was mainly characterized with desirable solvent permeability, high efficiency of mass transfer for reactants, and good accessibility and stability of the immobilized enzymes. The biosensor could work in water-miscible solvent such as a solvent mixture of acetonitrile and phosphate aqueous buffer, as well as hydrophobic organic solvent such as n-hexane. The biosensor had the highest sensitivity to methanol in both solvent systems. Under the stop-flow mode, the biosensor had the analytical working ranges from 80 µM to 90 mM methanol in n-hexane and 0.10 to 90 mM methanol in acetonitrile/buffer. When the biosensor functioned in n-hexane, it could take benzaldehyde as an alcohol substrate and was free from any pH disturbance. In the presence of coimmobilized horseradish peroxidase, the operational life of the biosensor was 60 assays and the shelf life was longer than two weeks. The biosensor has been satisfactorily applied to the determination of methanol in commercial gasolinemethanol blend samples. Massive entrapment enzyme within hydrophilic polymer is a common technique to construct the biphase biosensor operating in organic solvents.1-7 A distinct advantage of this technique is that large amounts of hydrophilic polymer are capable of being * Corresponding author. Fax: +852 3411 7348. E-mail: [email protected]. (1) Saini, S.; Hall, G. F.; Downs, M. E. A.; Turner, A. P. F. Anal. Chim. Acta 1991, 249, 1-15. (2) Iwuoha, E. I.; Smyth, M. R.; Lyons, M. E. G. Biosens. Bioelectron. 1997, 12, 53-75. (3) Wang, J. Talanta 1993, 40, 1905-1909. (4) Dong, S.; Guo, Y. Anal. Chem. 1994, 66, 3895-3899. (5) Brink, L. E. S.; Tramper, J. Biotechnol. Bioeng. 1985, 27, 1258-1269. 10.1021/ac049799d CCC: $27.50 Published on Web 06/11/2004

© 2004 American Chemical Society

massively coimmobilized with substances such as enzyme(s), cofactor, reaction mediator, solid support(s), conducting powder, and even aqueous buffer within the immobilization phase so that the enzyme can be immobilized in any desired combination to suit specific applications. Hydrogel techniques can create a stable enzyme immobilization platform both chemically and physically due to the swelling function of the filled water and the shrinkage inclination of the network.4,8 The membrane of the hydrogel can provide a desirable water-rich buffering environment for the entrapped enzyme as compared to other immobilization techniques, which in turn leads to better hydration of the enzyme. Unfortunately, the biocatalytic reactions with the hydrogelentrapped enzyme are generally rather sluggish due mainly to the innately low efficiency of the mass transfer of the substrate and product within the hydrogel network,1,5 which usually leads to an undesirable low overall reaction rate. Therefore, one of the biggest challenges in constructing enzyme entrapment within the massive hydrogel polymer matrixes performing in organic solvents is to make the substrate, product, and solvent have good transfer rate across the phase boundary and within the immobilization matrixes2,9 in order to achieve high sensitivity and stable biosensing response to the analyte. In this article, we report a novel enzyme entrapment structure consisting of a spongiform of the immobilized enzymes, formed by kneading hydrogel membrane entrapped with enzymes and other components with tiny highly hydrophobic particles. This enzyme entrapment technique possesses at least three unique characteristics over the traditional hydrogel techniques. First, the hydrogel membrane becomes thinner and increases the interfacial contact area. Second, the spongiform immobilization matrixes of the dual hydrophilic and hydrophobic structure have excellent solvent permeability for both hydrophilic and hydrophobic organic solvents. Third, the partition limit of the reactants is significantly minimized across the larger boundary areas and the longitudinal (6) Bickerstaff, G. F., Ed. Immobilization of Enzymes and Cells; Humana Press: Totowa, NJ, 1997. (7) Diaz-Garcia, M. E.; Valencia-Gonzalez, M. J. Talanta 1995, 42, 1763-1773. (8) Wu, X.; Choi, M. M. F. Anal. Chem. 2003, 75, 4019-4027. (9) Kawakami, K.; Furukawa, S. Appl. Biochem. Biotechnol. 1997, 67, 23-31.

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diffusional path is notably shortened in the thinner hydrogel layer. As a result, the major advantage of the proposed enzyme immobilization architecture is that the apparent enzyme activity is much higher than that of traditional hydrogel techniques since it has enormous large surface area-to-volume ratio. Hydroxyethyl carboxymethyl cellulose (HECMC) has been reported to be very useful in constructing organic-phase biosensors.4,8 In our preliminary studies, we observed that HECMC in aqueous solution easily interacted with quaternary ammonium cation, for example, an adduct of 3-methoxy-4-ethoxybenzaldehyde and 4-tert-butylpyridinium acetohydrazone (PAB), to form an insoluble ionotropy polymer hydrogel (HECMC-PAB).10 This polymer is thermodynamically stable, and its hydrogel is less swollen and is more plastic and elastic. Octadecylsilica (ODS) is a typical reversed-phase silica gel and possesses good mechanical properties, high hydrophobicity, and chemical inertness. Hence, we propose that HECMC-PAB and ODS would be the ideal materials for constructing the spongiform for enzyme immobilization. Alcohol biosensors based on alcohol oxidase (AOx) have been well studied to respond to short aliphatic alcohols in aqueous medium or gas phase since 1975.11 To fabricate a biosensor, AOx is generally immobilized on a support surface12-16 or entrapped within immobilization matrix17-22 with or without horseradish peroxidase (HRP). These biosensors mainly utilize an electrochemical transducer13,14,16-19,22-24 or optical transducer12,15,17 for signal transducing. Those alcohol biosensors utilizing hydrogen peroxide electrodes have the advantages of high sensitivity, wide linear range, and rather rapid steady response time (less than 2 min).21,23 But the disadvantages are that they have limited available potential windows and suffer from more electrochemical interference from the sample constituents. Biosensor utilizing oxygen electrode has the advantage of ease of operation and less electrochemical interference;14,19 however, the detection limit, is not as low as that of the hydrogen peroxide electrode due to the high background signal. Alcohol biosensors based on optical transducers detecting oxygen have the advantages of simple sensor structure and potential of remote assay when incorporating with an optical fiber. They are not easily poisoned and have no electrochemical interference,12,17 but they suffer from ambient light (10) Hoffmann, H.; Kastner, U.; Donges, R.; Ehrler, R. Polym. Gels Networks 1996, 4, 509-526. (11) Nanjo, M.; Guilbault, G. G. Anal. Chim. Acta 1975, 75, 169-180. (12) Lau, R. C. W.; Choi, M. M. F.; Lu, J. Talanta 1999, 48, 321-331. (13) Hall, C. E.; Datta, D.; Hall, E. A. H. Anal. Chim. Acta 1996, 323, 87-96. (14) Guilbault, G. G.; Danieisson, B.; Mandenius, C. F.; Moshbach, K. Anal. Chem. 1983, 55, 1582-1585. (15) Xiao, D.; Choi, M. M. F. Anal. Chem. 2002, 74, 863-870. (16) Varadi, M.; Adanyi, N. Analyst 1994, 119, 1843-1847. (17) Wolfbeis, O. S.; Posch, H. E. Fresenius Z. Anal. Chem. 1988, 332, 255257. (18) Patel, N. G.; Meier, S.; Cammann, K.; Chemnitius, G. C. Sens. Actuators, B 2001, 75, 101-110. (19) Mitsubayashi, K.; Yokoyama, K.; Takeuchi, T.; Karube, I. Anal. Chem. 1994, 66, 3297-3302. (20) Hikuma, M.; Matsuoka, H.; Takeda, M.; Tonooka, Y. Anal. Lett. 1993, 26, 209-221. (21) Vijayakumar, A. R.; Csoregi, E.; Heller, A.; Gorton, L. Anal. Chim. Acta 1996, 327, 223-234. (22) Gulce, H.; Glulce, A.; Kavanoz, M.; Coskun, H.; Yildiz, A. Biosens. Bioelectron. 2002, 17, 517-521. (23) Boujtita, M.; Hart, J. P.; Pittson, R. Biosens. Bioelectron. 2000, 15, 257263. (24) Liu, J.; Wang, J. Biotechnol. Appl. Biochem. 1999, 30, 177-183.

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interference compared with that of electrochemical alcohol biosensors. To date, most of the optical AOx-based biosensors are confined to functioning only in aqueous solutions. To our knowledge, no AOx-based biosensors working in organic solvents have been reported. The major difficulty in developing an organicphase alcohol biosensor is that the biocatalytic activity of AOx in nonaqueous solvents is rather low due to the dehydration effect on AOx in organic solvents.25,26 Although the stability of the enzyme can be much enhanced after the enzyme has been treated with stabilizers27,28 and well protected with a large immobilization matrix, a large mass-transfer resistance will be built up within the immobilization matrix. Furthermore, though immobilized AOx in n-hexane could show activity in the range from 40% to almost 100%,29 the alcohol bioreactors had mostly been set up with waterimmiscible organic solvent in the present of HRP and only applied in classical enzymology or bioconversion for a long-time reaction,9,29,30 rather than in analytical biosensing. Analysis of alcohols, especially methanol, in organic matrix has been considered as an important task in industry,1,2 and traditionally various instrumental techniques such as gas chromatography (GC),31,32 liquid chromatography,33,34 refractometry,35 and nuclear magnetic resonance spectrometry have been used.36 These techniques possess all of the necessary analytical requirements for sensitive, quantitative, and rapid assay of alcohols. However, their specificities are not very satisfactory especially when there is a wide variation in sample matrix since no specific enzymatic reaction is involved in the detection scheme. As such, the development of simple approaches for biocatalytic assay of alcohols in organic solvents is still challenging in response to the wide possible variation of sample matrix. Optical oxygen transducers based on quenching of fluorescence offers an advantage over other optical transducers due to their low background and simple operation.8 Herein, an organicphase optical alcohol biosensor is reported by coupling an optical oxygen transducer with the spongiform of immobilized AOx and HRP in silica gel/HECMC-PAB/ODS. To our knowledge, this is the first report on an organic-phase optical alcohol biosensor that could work in a solvent mixture of acetonitrile and phosphate aqueous buffer (as the water-miscible organic solvent) as well as n-hexane (as the water-immiscible organic solvent). This work mainly focuses on the studies of the analytical properties of our organic-phase alcohol biosensors in terms of selectivity, sensitivity, response time, ability to be immune to interferences, and lifetime. The mechanisms of the mass transfer governing the response are (25) Zaks, A.; Klibanov, A. M. J. Biol. Chem. 1988, 263, 8017-8021. (26) Klibanov, A. M. Trends Biotechnol. 1997, 15, 97-101. (27) Gibson, T. D.; Higgings, I. J.; Woodward, J. R. Analyst 1992, 117, 12931297. (28) Gibson, T. D.; Hulbert, J. N.; Pierce, B.; Webster, J. I. The Stabilization of Analytical Enzyme Using Polyelectrolytes and Sugar Derivatives. In Stability and Stabilization of Enzymes; van den Tweel, W. J. J., Harder, A., Buitelaar, R. M., Eds.; Elsevier Science B.V.: Amsterdam, 1993; pp 337-346. (29) Borzeix, F.; Monot, F.; Vandecasteele, J. Enzyme Microb. Technol. 1995, 17, 615-622. (30) Duff, S. J. B.; Murray, W. D. Biotechnol. Bioeng. 1989, 34, 153-159. (31) Agarwal, V. K. Analyst 1988, 113, 907-909. (32) Tackett, S. I. Analyst 1987, 112, 339-340. (33) Zinbo, M. Anal. Chem. 1984, 56, 244-247. (34) Pauls, R. E.; McCoy, R. W. J. Chromatogr. Sci. 1981, 19, 558-561. (35) William, S., Ed. Official Methods of Analysis of AOAC, 14th ed.; Association of Official Analytical Chemists: Washington, DC, 1984; pp 186-187. (36) Renzoni, G. E.; Shankland, E. G.; Gaines, J. A.; Callis, J. B. Anal. Chem. 1985, 57, 2864-2867.

also discussed. The primary difference between our biosensors and those reported in the literature or commercial sensors is that our biosensors are able to stably respond to short-chain aliphatic alcohols in organic phase. Another advantage of our alcohol biosensors over existing ones is that substrate and product inhibition is effectively reduced since they have good solubilities in hydrophobic organic solvents. In addition, the pH effect of the solvent on our biosensors can also be neglected. In real applications, our optical alcohol biosensors can simply and reliably determine the methanol contents in some commercial gasolinemethanol blend samples and are even able to respond to aromatic alcohol. EXPERIMENTAL SECTION Reagents. Alcohol oxidase (EC 1.1.3.13. from Hansenula species) with a specific activity of 22 units/mg of protein and horseradish peroxidase (EC 1.11.1.7.) with a specific activity of 290 units/mg of solid were purchased from Sigma (St. Louis, MO). Carboxymethyl cellulose sodium salt (medium viscosity) and 3-methoxy-4-ethoxybenzaldehyde were obtained from Fluka (Buchs, Switzerland). Silica gel particles (60 Å, 50 µm) were from Matrex (Leipzig, Germany). Octadecylsilica (32-63 µm, 100 Å) was obtained from Acros Organics (Geel, Belgium). Teflon tape was a product of Shuang Xing Teflon Factory (Zhuhai, China). Hydroxyethyl carboxymethyl cellulose sodium salt was synthesized from carboxymethyl cellulose sodium salt according to a modified method.37 Tris(4,7-diphenyl-1,10-phenanthroline)ruthenium(II) didodecyl sulfate dye ion pair was synthesized and oxygen-sensitive silica gel particles were prepared in our laboratory previously.38 An adduct of 3-methoxy-4-ethoxybenzaldehyde and 4-tert-butylpyridinium acetohydazone was prepared as described in the literature.39 All reagents were of analytical reagent grade or above and were used without further purification. All aqueous solutions were prepared with deionized water. Acetonitrile and n-hexane were of HPLC-grade solvents (Aldrich, Milwaukee, WI) and used without further purification. The acetonitrile/ buffer used in this work referred to the solvent mixture of pure acetonitrile and 0.050 M, pH 7.5 aqueous phosphate buffer (9:1 v/v). Air cylinder gases were purchased from Chun Wang Industrial Gases (Shenzhen, China). Enzymes Immobilization in Spongiform of Silica Gel/ HECMC-PAB/ODS Matrix. A 5% stock solution of HECMC was prepared by dissolving 5.0 g of HECMC in 100 mL of water, and its pH value was adjusted to 7.5 by a 0.050 M phosphate buffer. Five milligrams of PAB, 2 mg of HRP and 15 mg of AOx were successively dissolved in 0.75 mL of 0.050 M phosphate buffer (pH 7.5). The resulting solution was mixed with 65 mg of silica gel particles and left for 20 min. The mixture was then blended with 0.6 mL of the 5.0% HECMC solution, and the hydrogel matrix of the immobilized enzyme in silica gel/HECMC-PAB was formed. The hydrogel matrix was spread onto a glass plate under ambient conditions within 2 h to form a soft membrane. Last, the membrane was kneaded with 65 mg of ODS particles to engender a spongiform comixture. Assembly of Biosensor. The typical flow-through cell was machined from stainless steel and had a chamber volume of 0.38 (37) Li, H.; Huang, C. Y.; Yang, Z. L. Polym. Mater. Sci. Eng. 1998, 14, 34-37. (38) Wu, X.; Choi, M. M. F.; Xiao, D. Analyst 2000, 125, 157-162. (39) Chan, W. H.; Wu, X. Analyst 1998, 123, 2851-2856.

Figure 1. Schematic cross section of the flow-through cell constructed with coimmobilized AOx and HRP in the spongiform of silica gel/HECMC-PAB/ODS, and an optical oxygen transducer: (1) stainless steel cell body, (2) spongiform matrixes of immobilized enzymes, (3) Teflon membranes, (4) optical oxygen-sensitive particles, (5) transparent window, (6) sample inlet, (7) sample outlet, (8) excitation light beam, and (9) emission light beam.

mL (20 mm i.d. × 1.2 mm). Ten milligrams of oxygen-sensitive particles was packed against the window of the flow-through cell8 and covered with a piece of Teflon membrane (80 µm). The matrix of the immobilized enzymes was put into the flow-through cell and covered with another piece of Teflon membrane (80 µm) to form an alcohol biosensor. The volume of the immobilized enzyme matrix in the flow-though cell was 0.28 mL (20 mm i.d. × 0.90 mm). A schematic cross section of the flow-through cell is shown in Figure 1. When the biosensor was not in use, it was stored at 4 °C. Apparatus. The alcohol biosensor was positioned in a PerkinElmer LS-50B spectrofluorometer (Buckinghamshire, U.K.). Fluorescence intensity of the biosensor was controlled and measured by FL WinLab software (Perkin-Elmer). The emission intensity at 602 nm was collected at an excitation wavelength of 440 nm under batch conditions at 20 ( 1 °C and at a pressure of 101.3 kPa. All fluorescence measurements were made with 3-nm bandwidths for both the emission and excitation monochromators. Before sample analysis, all test solutions were passed through by a stream of compressed air at flow rate of 30 mL/min for 8 min and then left for 12 min. A homemade liquid flow controller8 was then used to deliver 5-mL test solutions through the alcohol biosensor. All biosensing processes were carried out in the stopflow mode. The response of the biosensor was brought back to the baseline by flushing with few milliliters of pure solvent. RESULTS AND DISCUSSION Optical Response of Alcohol Biosensor. The response mechanism of the biosensing process is based on the biocatalytic oxidation of alcohols by an oxygen molecule in the presence of immobilized AOx and HRP. The enzymatic reactions of the immobilized enzyme take place as follows:29,30 AOx

RCH2OH + O2 98 RCHO + H2O2 HRP

H2O2 98 H2O + 1/2O2

(1) (2)

At the first step, which is associated with AOx catalytic action, Analytical Chemistry, Vol. 76, No. 15, August 1, 2004

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Figure 2. Response curves of the alcohol biosensor on exposure to various concentrations of methanol in n-hexane: (1) 0.0, (2) 6.0, (3) 30.0, (4) 60.0, and (5) 90.0 mM methanol and (6) a signal of 90 mM methanol from baseline to saturation.

Figure 3. Response curves of the alcohol biosensor subjected to various concentrations of methanol in the acetonitrile/buffer: (1) 0.0, (2) 30.0, and (3) 60.0 mM methanol and (4) a signal of 60.0 mM methanol from baseline to saturation.

one alcohol molecule is oxidized by one oxygen molecule, releasing one hydrogen peroxide molecule and one aldehyde molecule. At the second step, which is related to the HRP catalytic action, hydrogen peroxide is further degraded to one molecule of water and equivalent half molecule of oxygen. Accordingly, the enzymatic reactions induced the depletion of the oxygen level surrounding the enzymes so that an oxygen gradient was created between and within two segments and, finally, induced the response of the oxygen transducer. The initial dissolved oxygen concentrations of the solvent, standard solutions, and samples might be slightly different. Thus, they were adjusted to the same level by passing through a stream of air. Figures 2 and 3 display the typical response curves of the alcohol biosensor in the stopflow mode on exposure to various concentrations of methanol in n-hexane and the acetonitrile/buffer solvents, respectively. In both biosensing systems, an increase in the fluorescence intensity was observed in the presence of methanol as oxygen was consumed in the enzymatic reactions. The higher the methanol concentration, the faster the increase and the larger the fluorescence intensity recorded. A 95% steady signal could only be obtained in 13 min for 90 mM methanol in n-hexane or 19 min for 60 mM methanol in acetonitrile/buffer, respectively (Figures 2 and 3). Hence, the response time for obtaining a steady signal will be too long. As displayed in Figures 2 and 3, the longer the contact time of the biosensor with the sample, the larger the fluorescence 4282

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Figure 4. Calibration curves at various concentrations of methanol in (1) n-hexane and (2) acetonitrile/buffer. Each data is an average of five tests.

intensity, the higher the sensitivity of the biosensor, and the better the signal-to-noise ratio achieved.15 But when the contact time was too long, exceptionally long response time would also result. To optimize these factors, the duration of the biosensing measurement was taken as 10 min. The dynamic working range under the selected experimental conditions was extended to nearly 2 orders when compared to the method of measuring the steady signal. It was also noted that the biosensor exhibited slight hysteresis for relatively higher concentrations of methanolic n-hexane solutions when comparing the signal heights of 60 or 90 mM methanolic solutions (Figure 2). This problem, in fact, can be minimized by exposing the biosensor to a running pure solvent for 10 min. The overall process correlates to the concentration of methanol in the organic solvent, which can be quantified by intensity quenching measurements and described arbitrarily as the following equation:8,17

Rs ) (Itest)/(Ibaseline)

(3)

where Rs is defined as the relative signal change of the fluorescence intensity of the biosensor, which is a function of the methanol concentration; the terms Ibaseline and Itest represent the detected fluorescence signals of the biosensor when exposed to the organic solvent and to methanol in the organic solvent in a defined time duration, respectively. Calibration curves of the biosensor operating under the stop-flow mode in both solvent systems are displayed in Figure 4. The maximum signal ratios of saturation to baseline are 3.8-fold for the 90.0 mM methanol in n-hexane and 2.8-fold for the 90.0 mM methanol in acetonitrile/ buffer solvent mixture. The analytical working ranges are as follows: 80 µM-60.0 mM with detection limit of 80 µM methanol (S/N ) 3) in n-hexane (Rs ) 0.0381[methanol] + 1.0603; r2 ) 0.9975); 0.10-90 mM with detection limit of 0.10 mM methanol (S/N ) 3) in the acetonitrile/buffer solvent mixture (Rs ) 0.0205[methanol] + 1.0394; r2 ) 0.9933). Therefore, the proposed biosensor could provide an efficacious analysis of methanol in both n-hexane and acetonitrile/buffer solvents. In principle, this biosensor can also determine other low primary alcohols as well. Configuration of Enzyme Immobilization. The configuration of the enzyme immobilization was critical to the performance of the alcohol biosensor. It was noticed that the solution of the

enzymes and other ingredients including PAB was easily distributed on the large surface of silica gel particles. Once AOx and HRP had been deposited, they were firmly retained on the surface of the silica gel particles accompanied with some residue water. When the HECMC solution reached the surface of the silica gel, the hydrogel of HECMC-PAB/water system was in situ formed40 due to the charge neutralization mechanism.10 We found that this ionotropy polymer hydrogel of the immobilization matrixes was stable with some types of organic solvent systems. Pure acetonitrile or the acetonitrile/buffer alcoholic solutions were able to diffuse through this hydrogel membrane, but alcoholic n-hexane solution was almost not able to diffuse into the hydrogel membrane. This observation suggested that the acetonitrile/buffer (as a typical water-miscible organic solvent) together with alcohols could penetrate into the polymer hydrogel membrane.4 But n-hexane (as a typical water-immiscible organic solvent) generally worked as a solvent phase in which the alcohols had to undergo a phase-transfer process from the organic solvent into the polymer hydrogel membrane and then slowly diffuse within the water-filled polymer hydrogel phase.9,30 As expected, we found that the biosensing system composed of such bulk thick immobilized enzyme membrane and any of the organic solvents could not effectively respond to alcohols. This was because the thick, as well as the small surface-to-volume ratio, the hydrogel membrane seriously hindered the transport of the substrate. When the membrane was kneaded and inlaid with some tiny hydrophobic ODS particles to form matrixes of silica gel/HECMC-PAB/ODS, the whole system became spongiform, where the hydrogel polymer membrane became thinner. The special feature of this spongiform is the dual hydrophilic and hydrophobic properties on the interface between the polymer film and the ODS particles.8 We found that pure acetonitrile, the acetonitrile/buffer mixture, and n-hexane solvents could directly penetrate through the spongiform (e.g., 4 mm thick) of the immobilized enzyme within 12 s. As both organic solvents could efficiently diffuse through the Teflon membrane in less than 6 s, it was employed to steadily hold the spongiform of the immobilized enzymes in the flow cell.19 Thus, the enzymatic oxidation reaction was strictly restricted within the immobilization matrixes. The stable and reproducible response of the biosensing system was attributed to the Teflon membrane since the immobilized enzyme phase was no longer flushed by the moving organic solution. In addition, the spongiform could also firmly affix to the optical oxygen transducer. We found that the biosensor without HRP could repetitively respond to 45 mM methanol in n-hexane for ∼25 times and the response would degrade significantly afterward, while another biosensor with the coimmobilized HRP could achieve reproducible responses to 90 mM methanol in n-hexane for ∼60 times. This result evidently implied that hydrogen peroxide could inhibit AOx,30 and its inhibition effect was effectively minimized by reaction 2 when HRP was coimmobilized in the matrixes. However, the fluorescence signal of this biosensor was 22% lower than that without using HRP because of the simultaneous production of oxygen by the reaction 2. Selectivity. Methanol, ethanol, and n-butanol in n-hexane or in the solvent mixture of acetonitrile/buffer were separately tested by the proposed biosensor. The results of the relative sensitivity (40) Hussain, S.; Keary, C.; Craig, D. Q. M. Polymer 2002, 43, 5623-5628.

Table 1. Relative Sensitivity of the Biosensor toward Various Alcohols in n-Hexane and the Acetonitrile/ Buffer Solvents relative signal change (Rs) in n-hexanea compound methanol ethanol 1-butanol benzyl alcohol a

relative signal change (Rs) in acetonitrile/buffera

30 mM 60 mM 90 mM 30 mM 2.26 1.86 1.19 1.16

3.31 2.43 1.26 1.24

3.82 2.95 1.37 1.35

1.76 1.44 1.13 1.00

60 mM

90 mM

2.29 1.91 1.20 1.00

2.83 2.14 1.32 1.00

Each datum is an average of three tests.

of the biosensor toward three alcohols are depicted in Table 1. The biosensor shows the highest sensitivity to methanol in both solvent systems, while the rates of oxidation decrease with the increase in the chain length of the aliphatic alcohols. This trend is the same as AOx in aqueous solutions.12 The results demonstrated that regardless of the ways of substrate transfer into the polymer film, e.g., by diffusing from the acetonitrile/buffer or partitioning from n-hexane, the enzyme in the proposed immobilization structure exhibited good accessibility for alcohols in both water-miscible and water-immiscible organic solvents. Moreover, the polymeric hydrogel provided AOx with sufficient conformational flexibility needed for catalysis in both organic solvents. It was also noted that the biosensor showed markedly lower sensitivity to alcohols in the acetonitrile/buffer than that in n-hexane as shown in Table 1. Moreover, the response signals of the proposed biosensor toward methanol in pure acetonitrile were completely suppressed after several tests. These observations suggested that acetonitrile in the immobilization polymer film would cause an undesirable low rate of overall biocatalytic oxidation for the immobilized-AOx.25 On the other hand, it was noted that the response of the biosensor was not much affected by the water content following pretreatment with commercially available 4-Å dry molecular sieves pellets and by water-saturated n-hexane. This fact, together with the biosensor working in n-hexane being free from pH disturbances as discussed in the following section, strongly suggested that the hydrogel indeed provided better protection for the entrapped AOx compared to that in acetonitrile/buffer. As such, the biocatalytic activity of immobilized AOx is retained and consequently the higher sensitivity of the biosensor is achieved.29 Benzyl alcohol was also further investigated. To our surprise, the biosensor produced some responses to benzyl alcohol in n-hexane as shown in Table 1 though the sensitivity of the biosensor was fairly low. Heretofore, this is the first report on the immobilized AOx to treat benzyl alcohol as the substrate in organic-phase analytical applications. This result revealed that the retention of activity and stability of AOx in the polymer hydrogel was high enough for the biocatalytic oxidation of benzyl alcohol in n-hexane solvent. It showed that the benzyl alcohol molecules could partition into the polymer film fast enough to couple the biocatalytic oxidation reaction, though not as good as the aliphatic alcohols. In addition, it also strongly suggested that the transfer of the enzymatic product benzaldehyde out of the polymer film into the n-hexane phase was favorable and rapid so that the Analytical Chemistry, Vol. 76, No. 15, August 1, 2004

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Figure 5. Effect of pH on the response of alcohol biosensor upon exposure to 50 mM methanol in acetonitrile/buffer.

inhibition effect of benzaldehyde did not pose a problem.9,29,30 On the other hand, the proposed biosensor could not produce any response signal even to 90 mM benzyl alcohol in the acetonitrile/ buffer for more than 0.5 h though benzyl alcohol together with the solvent easily penetrated into the hydrogel membrane. It implied that the biocatalytic activity of the immobilized AOx appeared to be too low to perform biosensing of benzyl alcohol.25,26 Therefore, the selectivity and sensitivity of our biosensor mainly depended on the substrate, as well as the organic solvent system chosen. Effect of pH. The pH effect on the biosensor was first investigated in the acetonitrile/buffer system. Nine separate 50 mM methanolic acetonitrile/buffer solutions were prepared in acetonitrile/buffer that had pH values adjusted from 5.5 to 9.5. Figure 5 shows the response of the biosensor changed markedly over the pH ranges of 5.5-6.5 and 8.5-9.5. The highest and broader range of the response toward methanol was in the pH 6.5-8.5. These results were consistent with other aqueous alcohol biosensors based on immobilized AOx.12,18 Hence, this again suggested that the acetonitrile/buffer solutions could significantly affect the enzymatic activity or reaction due to the diffusion of the organic solvent into the microenvironment surrounding the enzymes. Further experiments were performed by separately exposing the biosensor to five 50 mM methanolic n-hexane solutions. Each of these methanolic n-hexane solutions had been previously equilibrated with a 0.2 M phosphate buffer with pH ranging from 4.6 to 10.2. The results showed that the relative signal changes did not exhibit any significant change in the response of the biosensor. This observation demonstrated that the immobilization matrixes contained sufficient amounts of aqueous phosphate buffer and possessed a good pH buffering capacity so that the relatively small portion of aqueous buffer solution in n-hexane could not induce any significantly pH change on the immobilization matrixes.8 Hence, our organic-phase alcohol biosensor can be free from proton disturbance when it works with hydrophobic organic solvents. In addition, the results also indirectly proved our proposed mechanisms of the mass transfer of alcohols in the two types of organic solvents. Repeatability. Reproducibility and repeatability are important factors for considering the success of a biosensing process. Figures 2 and 3 separately display the signal changes of the biosensor when it is exposed to step concentration changes from 0.0 to 90.0 mM and then back to 0.0 mM of methanol in n-hexane and from 0.0 to 60.0 mM and then back to 0.0 mM of methanol in 4284

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the acetonitrile/buffer, respectively. This demonstrates that the biosensor has a highly reproducible response to methanol in both organic solvent systems. To further investigate the repeatability, the biosensor was alternatively exposed to the pure n-hexane and 60.0 mM methanolic n-hexane solution nine times. The biosensor exhibited a fairly desirable analytical feature of repeatability (n ) 9, RSD ) 2.03%). Lifetime. The lifetime of the biosensor means both storage stability and operation stability. It was noted that the lifetime of the biosensor operating in n-hexane is notably longer than that in acetonitrile/buffer. The operation stability was systematically examined and consisted of 60 assays of 90.0 mM methanol in n-hexane over a period of 4 days. The responses were reasonably consistent for more than 45 assays. After 60 assays, the response signal to 90.0 mM methanol decreased to 80% of its original value. This relatively long lifetime of our biosensor is attributed to the better protection of the immobilized AOx, as well as the coentrapment of HRP as described in the above sections. For storage stability, it could reproduce its signal as 55% magnitude of its first response to 90.0 mM methanol in n-hexane after it had been sealed under moist conditions and stored at 4 °C for longer than two weeks. The relatively stable and repeatable responses of the biosensor indicated that leaching of the enzymes from the immobilization matrixes was not a problem for the operational lifetime. On the other hand, the enzymatic activity of the coentrapped HRP was 40% of its initial value as checked by the literature method28 even after 2-month storage. Thus, the lifetime of the biosensor must be mainly determined by the stability of the AOx rather the HRP. Interference. The proposed alcohol biosensor was separately investigated with some potential interferences such as formaldehyde (30 mM), formate (15 mM), benzaldehyde (40 mM), L-ascorbic acid (25 mM), and glucose (25 mM) in acetonitrile/ buffer or formaldehyde (30 mM), formate (15 mM), and benzaldehyde (40 or 100 mM) in n-hexane. AOx is able to oxidize formaldehyde to formate.30 However, our biosensor showed its 97-100% responses toward 30.0 mM methanol in both solvent systems in the presence of formaldehyde compared to that of methanolic solutions in the absence of formaldehyde. The response of the biosensor toward methanol was also not affected by the presence of formate. These facts show that the oxidization of formaldehyde has not effectively taken place within the duration of the response time (10 min). Thus, the stoichiometry of methanol oxidation under the catalysis of the immobilized AOx and the operational condition is indirectly confirmed with which methanol is consumed and formaldehyde is produced. We found that the presence of benzaldehyde in the acetonitrile/buffer could produce an inhibiting effect on the response of the biosensor. For example, the response signals toward 30.0 mM methanolic acetonitrile/buffer solution with 40 mM benzaldehyde were not reproducibile and disappeared after several tests. Again, this suggested that the acetonitrile/buffer could easily carry the solute into the polymer hydrogel film and directly diffuse to the immobilized enzymes. The biosensor was further investigated by exposing it to 30 mM methanolic n-hexane solution containing 40 and 100 mM benzaldehyde, respectively. Fortunately, benzaldehyde in n-hexane did not inhibit the immobilized enzymes. Hence, it strongly suggested that the concentration of benzalde-

Table 2. Results of Methanol Assay on the Gasoline-Methanol Blend Samples value from the biosensor method

AOAC value

sample

methanol contenta (% v/v)

RSD (%)

methanol added (% v/v)

recoveryb (%)

RSD (%)

methanol contenta (% v/v)

RSD (%)

M15

14.4

2.88 3.01

2.81 3.09 3.22 3.49 3.91 3.70

2.35

18.3

98.6 99.5 101.2 101.4 99.1 100.7

14.6

M19

6.00 12.0 18.0 6.00 12.0 20.0

18.4

2.17

a

The methanol content is an average of five tests. b The recovery is an average of three tests.

hyde in the microenvironment around the immobilized enzymes was indeed significantly low due to better solubility of benzaldehyde in n-hexane.30 This meant that the dampening effect of inhibitors could be effectively minimized by entrapping the enzyme in the massive hydrogel and working in highly soluble hydrophobic organic solvent. Furthermore, the biosensor was negligibly affected by L-ascorbic acid or glucose in the acetonitrile/ buffers. Methanol Analysis in Gasoline-Methanol Blend Samples. An attempt to apply the optical alcohol biosensor in gasolinemethanol blends was made by analyzing the methanol content in commercial gasoline-methanol fuel blends. M15 and M19 samples were obtained from local fuel stores. These samples are mainly the mixture of methanol with conventional petroleum. It was observed that there was no effect on the response of the biosensor when the concentration of the conventional petroleum (No. 93 or 97) containing no methanol in n-hexane was below 5% (v/v). Thus, the sample matrixes would not exhibit significant interference to the determination of methanol in the petroleum fuel samples. Samples of 0.30 mL of M15 and 0.25 mL of M19 gasolinemethanol blend were separately diluted to 25.0 mL with n-hexane. The relative signal change (Rs) of each sample n-hexane solution was measured and compared with that of a set methanolic standard solution. To investigate the effect of water in a real sample on the response of the biosensor, several artificial gasoline blends were prepared with No. 97 petroleum containing 15.0 or 20.0% methanol (v/v) and 0.05-0.32% water (v/v). These samples were then subject to the biosensor. The responses of the biosensor did not show any significant change with samples containing water. Hence, it was concluded that a water content in gasoline blends of less than 0.32% (v/v) would not affect the proposed biosensor method. Furthermore, a recovery test was performed and the results obtained from an AOAC GC method41 were also (41) William, S., Ed. Official Methods of Analysis of AOAC, 14th ed.; Association of Official Analytical Chemists: Washington, DC, 1984; p 187.

compared with our proposed biosensor method. Our findings demonstrated that the biosensor showed performance comparable to the standard GC method as depicted in Table 2. The recoveries are good as well for the proposed method. The results also showed only slight differences between these two methods; thus, our proposed biosensor method can provide an alternative simple approach for methanol analysis in the gasoline-methanol blend samples. CONCLUSION An organic-phase optical alcohol biosensor was for the first time developed by coentrapping AOx and HRP in the ionotropy polymer hydrogel matrixes of silica gel/HECMC-PAB and further kneading into the spongiform with ODS particles. This novel immobilization architecture is mostly characterized with high and efficient mass transfer of the reactants and good accessibility and stability of the immobilized enzymes when functioning in both acetonitrile/buffer and n-hexane solvents. Under the stop-flow mode, the biosensor works well analytically and has relatively high sensitivity when the analysis time is kept at 10 min. The response of the biosensor is not affected by interferences including water when performing in n-hexane solvent. The real application of the biosensor for the petrochemical industry is simple and reliable. Future work directed to the fabrication of other enzyme-based biosensors using similar immobilization architecture is anticipated to be promising. More work will proceed in our laboratory, and the findings will be published in the future. ACKNOWLEDGMENT The work described in this paper was supported by a grant from the Research Grants Council of the Hong Kong Special Administrative Region, China (project HKBU 2058/98P). Received for review February 5, 2004. Accepted May 10, 2004. AC049799D

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