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Amperometric determination of choline and acetylcholine with enzymes immobilized in a photocross-linkable polymer
Analytica Chimica Acta, 228 (1989) 49-53 Elsevier Science Publishers B.V., Amsterdam
49 - Printed
in The Netherlands
Amperometric determination of choline and acetylcholine with enzymes immobilized in a photocross-linkable polymer JEAN-LOUIS Research Laboratory of Resources Utilization,
MARTY
a
Tokyo Institute of Technology, 4259 Nagatsuta-cho, KOJI SODE and ISA0
KARUBE
Midoru-ku,
*
Research Center for Advanced Science and Technology, University of Tokyo, 4-6-I Komaba, Meguro-ku, (Received
29th November
Yokohama 227 (Japan)
Tokyo 153 (Japan)
1988)
SUMMARY
Choline and acetylcholine sensors were prepared by using choline oxidase and acetylcholinesterase, entrapped in photocrosslinkable poly(viny1 alcohol) bearing styrylpyridinium (PVA-SbQ). The measurements were based on the detection of hydrogen peroxide liberated by an enzyme reaction (choline oxidase) or two sequential enzyme reactions (acetylcholine esterase and choline oxidase). The determination range for choline was 2.5-150 pmol 1-l and for acetylcholine 20-750 nmol I-‘. The response times were 2-4 mm. The immobilized enzyme membranes stored in a dry state were very stable and no loss of activity was observed after storage for 60 days.
A number of sensors have been developed for the determination of choline and acetylcholine, the detection of which is important in biological media. For example, the choline sensor can be used for the determination of choline-containing phospholipids or choline esterase activity in blood. The acetylcholine sensor may be an excellent tool for the detection of organophosphorus pesticides. Ion-selective microelectrodes [l] and an acetylcholine sensor based on an ion-selective field effect transistor (ISFET) [2] have been reported, but most work has been based on enzyme electrodes with potentiometric detection of acetylcholine [3-51 or amperometric detection of oxygen [6-81
a Permanent address: Groupe d’Etudes et de Recherches Appliquees Pluridisciplinaires, UA CNRS 461, Chemin de la Passio Vella, 66025 Perpignan, France. 0003-2670/90/$03.50
0 1990 Elsevier Science Publishers
B.V.
and hydrogen peroxide [9,10] for choline and acetylcholine. The best sensitivity was obtained with the amperometric detection of hydrogen peroxide. In the enzyme electrodes, the enzymes are immobilized covalently using activated membranes or mineral support [3,4,7,9,10] or by crosslinking with an inert protein such as albumin treated with glutaraldehyde [4,10]. A physical entrapment method using a synthetic gel matrix has also been reported [6,8]. No method has been proposed that is generally applicable for all enzymes. However, the entrapment method has the distinct advantage that since the immobilization requires no covalent bond, the native properties of the enzymes are maintained. Ichimura [ll] developed a novel method for enzyme immobilization by using photocross-linkable poly(viny1 alcohol) bearing styrylpyridinium (PVA-SbQ). The photochemical technique to en-
50
trap bioactive materials is of particular interest owing to the mild conditions for network formation of the polymeric matrix, no change in pH or temperature being necessary. The PVA-SbQ method is rapid and simple and the enzyme structure is hardly affected. The membranes also show excellent mechanical properties. In this study, we attempted to utilize this immobilization method to construct enzyme sensors for the determination of choline and acetylcholine. The membranes with immobilized enzymes were coupled with an amperometric hydrogen peroxide sensor. For the choline sensor, a choline oxidase membrane was used, whereas in the acetylcholine sensor an acetylcholine esterase membrane was combined with the choline oxidase membrane. These two enzymes were also co-immobilized in a single membrane with beneficial results.
EXPERIMENTAL
Reagents Choline oxidase (E.C. 1.1.3.17, 10 U mgg’, from Alcaligenes sp.), acetylcholine esterase (E.C. 3.1.1.7, 1000 U mgg’, from electric eel), peroxidase (E.C. 1.11.1.7, 80 U mg-i, from horseradish), choline chloride, acetylcholine chloride and acetylthiocholine iodide were purchased from Sigma. Poly(viny1 alcohol)-styrylpyridinium (PVA-SbQ; degree of polymerization 1700, degree of saponification 88, SbQ content 1.3 mol-%, solid content ll%, pH 7) was kindly provided by Prof. Ichimura (Research Institute for Polymers and Textiles, Tsukuba). Cellulose nitrate membranes were obtained from Toyo Roshi and other reagents were of analytical-reagent grade. Preparation of enzyme membrane Choline oxidase (20 U) and/or acetylcholine esterase (50 U) dissolved in 0.2 ml of water were added to 1 g of a 11% aqueous solution of PVA-SbQ. The mixture was stirred magnetically at room temperature to give a homogeneous solution of PVA-SbQ. The mixture was spread on a cellulose nitrate membrane using a bar coater and
K. SODE
air dried at room temperature. The air-dried brane was exposed to normal fluorescent light for 2 h.
ET AL.
memroom
Measurement of enzyme activity by a spectrophotometric method Free and immobilized choline oxidase were assayed as follows. The reaction mixture [containing 0.2 ml of 0.1 M choline chloride solution, 1.3 ml of 0.1 M phosphate buffer (pH 8) 0.25 ml of 4-aminoantipyrine solution (0.3% w/v), 0.25 ml of phenol solution (0.2% w/v), 0.25 ml of peroxidase solution (2 U ml-‘) and 0.25 ml of distilled water] and an appropriate amount of the free or immobilized enzyme were incubated for 10 min at 35“C in a shaking machine. The reaction was stopped by adding 2.5 ml of ethanol. The absorbance at 480 nm was read with a spectrophotometer (Jasco Uvidec-510). Free and immobilized acetylcholine esterase were assayed as follows. The reaction mixture [containing 0.5 ml of 4 X 10e3 M acetylthiocholine solution, 1 ml of 0.1 M phosphate buffer (pH 8) and 0.5 ml of 4 x 10m3 M 5,5’-dithiobisnitrobenzoate] and the free or immobilized acetylcholine esterase were incubated for 10 min at 35 o C in a shaking machine. The reaction was stopped by adding 1 ml of ethanol. The absorbance at 405 nm was read with the spectrophotometer as above.
Preparation of the sensor The system consisted of three electrodes: an Ag/AgCl reference electrode, a Pt counter electrode and a Pt working electrode (0.3 cm2 area) covered with an immobilized enzyme membrane and a dialysis membrane. The activity of immobilized choline oxidase was 15 nmol min-’ cmp2, the activity of immobilized acetylcholine esterase was 17 nmol mm’ cme2 and the activities of co-immobilized choline oxidase and acetylcholine esterase were 13 and 8 nmol min ’ cm- 2, respectively (Table 1). A constant potential (+ 700 mV vs. Ag/AgCl) was applied with a potentiostat (Hokuto Denko) and the resulting current from the breakdown of hydrogen peroxide was measured with a strip-chart recorder (TOA Electronic Polyrecord).
AMPEROMETRIC
DETERMINATION
OF CHOLINE
AND
ACETYLCHOLINE
51
TABLE 1 Characteristics of choline and acetylcholine enzyme electrodes (CHO = choline esterase; ACE = acetylcholine esterase) Electrode
CHO membrane CHO membrane + ACE membrane Co-immobilized CHO + ACE
Membrane activity (nmol min-’ cm-‘) (spectrophotometric method)
Current (nA)
CHO
ACE
Choline (1 x 1O-4 M)
Acetylcholine (4 x 1O-4 M)
1s 1s
0 17
180 83
0 54
13
8
168
167
Procedure The sensor is immersed in 10 ml of 0.05 M phosphate buffer (pH 8) and maintained at 35OC with thermostated water circulation. When the current of the sensor has become constant, an appropriate volume of choline or acetylcholine solution is injected into the jacketed vessel. The current increases rapidly and reaches a steady state within 2 min when using a choline oxidase membrane, 4 min when using the two enzymes immobilized simultaneously and 8 min when using two different membranes.
a broad pH range. The highest activity for immobilized choline oxidase was obtained at pH 9.5-10 (carbonate buffer) whereas the free enzyme is optimum at pH 8. A shift of the pH optimum and broadening of enzyme activity after immobilization are common phenomena and have been explained by Goldstein et al. [12]. In this work, the shift in the pH optimum may be due to the effect of the photofunctional group of the polymer. Studies of stability at 35°C showed that the activity of the sensor decreases quickly at pH 6 and above pH 9.5 (Fig. 2). Therefore, the choline sensor should be used between pH 7 and 9.
RESULTS AND DISCUSSION
Acerylcholine sensor The acetylcholine sensor gave a linear calibration plot for the range 2.0 x 10e5-7.5 x 10P4 M with a slope and linear correlation coefficient of
Choline sensor The choline sensor gave a linear calibration plot for the range 2.5 X 10-6-1.5 x 10e4 M choline with a slope and linear correlation coefficient of 1.90 nA 1 pmol-’ and 0.9996, respectively. The concentration range is well below those obtained with oxygen electrode-based sensors [6,7]. Moreover, the linear range is larger than that obtained by Mascini and Moscone [9] with the same type of detection. The average response time was 2 min. We assume that the superior features of this sensor depend on the immobilization method. The high sensitivity and good linearity make the sensor useful for measurements in biological fluids. The effect of pH on the choline electrode (Fig. 1) was studied after 15 min of incubation of free or immobilized choline oxidase in different buffers at 35’C. Free choline oxidase was denaturated at pH lower than 6.5 or higher than 9, whereas the immobilized choline oxidase kept its activity over
I
I
PH
Fig. 1. Response of the choline sensor and free choline oxidase in different buffer solutions at several pH values to addition of choline giving a final concentration of 50 nmol 1-l for the choline sensor and 5 mmol 1-l for free choline oxidase. 0, 0, 0.05 M phosphate buffer; 0, n, 0.05 M pyrophosphate buffer; A, A, 0.05 M carbonate buffer. Open symbols for choline sensor and closed symbols for free choline oxidase.
K. SODE
52
372 nA 1 mmol-’ and 0.9998, respectively. This result is better than the practical range obtained with the pH electrode [3-51 and ion-selective electrodes [l]. This result was obtained by using a membrane containing co-immobilized choline oxidase and acetylcholine esterase. The sensitivity with co-immobilized enzymes is better than that obtained by using membranes with two enzymes immobilized separately (Table 1). This is due to poor diffusion of choline to the choline oxidase membrane across the nitrate cellulose membrane. Indeed, when the choline oxidase membrane is covered with an acetylcholine esterase membrane, the response to choline decreases by 54% (Table 1). The response time of the biosensor for acetylcholine was 4 min with co-immobilized enzymes. By using two enzyme membranes, however, 8 min are necessary to obtain a steady-state current. The results indicate that the sensitivity and response time obtained with a two-enzyme membrane are much superior to those obtained by using a two-membrane system. The ultimate aim of this work is to develop a novel enzyme sensor system for the determination of organophosphorus pesticides. This measurement is based on the inactivation of cholinesterase by such pesticides.
ET AL.
Time (days)
Fig. 3. Stability of the free (0) and immobilized (W) choline oxidase stored at 4” C in 0.05 M phosphate buffer (pH 8) and immobilized choline oxidase in a dry film (A) at 4 o C.
Stability Figure 3 shows the storage stability of the choline oxidase membrane. The immobilization of choline oxidase increases the stability of the enzyme. After storage for 60 days in 0.05 M phosphate buffer (pH 8) at 4°C the decrease in the activity was about 35%. When the enzyme film was stored in a dry state at 4”, no decrease in the activity was observed. Ichimura [ll] also reported that glucose amylase immobilized by PVA-SbQ shows no inactivation even after storage for 1 year in a dry state at 4OC. The stabilities of the acetylcholinesterase and co-immobilized enzyme membranes were similar (results not shown).
REFERENCES
Fig. 2. 35 o C. 0. pH n . pH
Stability of the choline sensor at different pH values at *. pH 6; *, pH 7; 0, pH 8 (0.05 M) phosphate buffer); 9 (0.05 M pyrophosphate buffer): A, pH 9.5; A, pH 10; 10.5 (0.05 M carbonate buffer).
1 A. Jaramillo, S. Lopez, J.B. Justice, J.D. Salomone and D.B. Neill, Anal. Chim. Acta, 146 (1983) 149. 2 M. Gotoh, E. Tamiya, M. Monoi, Y. Kagawa and I. Karube, Anal. Lett., 20 (1987) 857. 3 K. Gibson and G.G. Guilbault, Anal. Chim. Acta, 80 (1975) 245. 4 P. Durand, A. David and D. Thomas, Biochim. Biophys. Acta, 527 (1978) 277. 5 M.F. Suaud-Chagny and J.F. Pujol, Analusis, 13 (1985) 25. 6 K. Matsumuto, H. Seijo, I. Karube and S. Suzuki, Biotechnol. Bioeng., 22 (1980) 1071.
AMPEROMETRIC
DETERMINATION
OF CHOLINE
AND
53
ACETYLCHOLINE
7 L. Campanella, M. Mascini, G. Palleschi and M. Tomassetti, Clin. Chim. Acta, 151 (1985) 71. 8 L. Campanella, M. Tomassetti and M.P. Sammartino, Analyst, 113 (1988) 77. 9 M. Mascini and D. Moscone, Anal. Chim. Acta, 179 (1986) 439.
10 T. Yao, Anal. Chim. Acta, 153 (1983) 169. 11 K. Ichimura, J. Polym. Sci., 22 (1984) 2817. 12 L. Goldstein, Y. Levin and E. Katchalski. Biochemistry, (1964) 1913.
3
49 - Printed
in The Netherlands
Amperometric determination of choline and acetylcholine with enzymes immobilized in a photocross-linkable polymer JEAN-LOUIS Research Laboratory of Resources Utilization,
MARTY
a
Tokyo Institute of Technology, 4259 Nagatsuta-cho, KOJI SODE and ISA0
KARUBE
Midoru-ku,
*
Research Center for Advanced Science and Technology, University of Tokyo, 4-6-I Komaba, Meguro-ku, (Received
29th November
Yokohama 227 (Japan)
Tokyo 153 (Japan)
1988)
SUMMARY
Choline and acetylcholine sensors were prepared by using choline oxidase and acetylcholinesterase, entrapped in photocrosslinkable poly(viny1 alcohol) bearing styrylpyridinium (PVA-SbQ). The measurements were based on the detection of hydrogen peroxide liberated by an enzyme reaction (choline oxidase) or two sequential enzyme reactions (acetylcholine esterase and choline oxidase). The determination range for choline was 2.5-150 pmol 1-l and for acetylcholine 20-750 nmol I-‘. The response times were 2-4 mm. The immobilized enzyme membranes stored in a dry state were very stable and no loss of activity was observed after storage for 60 days.
A number of sensors have been developed for the determination of choline and acetylcholine, the detection of which is important in biological media. For example, the choline sensor can be used for the determination of choline-containing phospholipids or choline esterase activity in blood. The acetylcholine sensor may be an excellent tool for the detection of organophosphorus pesticides. Ion-selective microelectrodes [l] and an acetylcholine sensor based on an ion-selective field effect transistor (ISFET) [2] have been reported, but most work has been based on enzyme electrodes with potentiometric detection of acetylcholine [3-51 or amperometric detection of oxygen [6-81
a Permanent address: Groupe d’Etudes et de Recherches Appliquees Pluridisciplinaires, UA CNRS 461, Chemin de la Passio Vella, 66025 Perpignan, France. 0003-2670/90/$03.50
0 1990 Elsevier Science Publishers
B.V.
and hydrogen peroxide [9,10] for choline and acetylcholine. The best sensitivity was obtained with the amperometric detection of hydrogen peroxide. In the enzyme electrodes, the enzymes are immobilized covalently using activated membranes or mineral support [3,4,7,9,10] or by crosslinking with an inert protein such as albumin treated with glutaraldehyde [4,10]. A physical entrapment method using a synthetic gel matrix has also been reported [6,8]. No method has been proposed that is generally applicable for all enzymes. However, the entrapment method has the distinct advantage that since the immobilization requires no covalent bond, the native properties of the enzymes are maintained. Ichimura [ll] developed a novel method for enzyme immobilization by using photocross-linkable poly(viny1 alcohol) bearing styrylpyridinium (PVA-SbQ). The photochemical technique to en-
50
trap bioactive materials is of particular interest owing to the mild conditions for network formation of the polymeric matrix, no change in pH or temperature being necessary. The PVA-SbQ method is rapid and simple and the enzyme structure is hardly affected. The membranes also show excellent mechanical properties. In this study, we attempted to utilize this immobilization method to construct enzyme sensors for the determination of choline and acetylcholine. The membranes with immobilized enzymes were coupled with an amperometric hydrogen peroxide sensor. For the choline sensor, a choline oxidase membrane was used, whereas in the acetylcholine sensor an acetylcholine esterase membrane was combined with the choline oxidase membrane. These two enzymes were also co-immobilized in a single membrane with beneficial results.
EXPERIMENTAL
Reagents Choline oxidase (E.C. 1.1.3.17, 10 U mgg’, from Alcaligenes sp.), acetylcholine esterase (E.C. 3.1.1.7, 1000 U mgg’, from electric eel), peroxidase (E.C. 1.11.1.7, 80 U mg-i, from horseradish), choline chloride, acetylcholine chloride and acetylthiocholine iodide were purchased from Sigma. Poly(viny1 alcohol)-styrylpyridinium (PVA-SbQ; degree of polymerization 1700, degree of saponification 88, SbQ content 1.3 mol-%, solid content ll%, pH 7) was kindly provided by Prof. Ichimura (Research Institute for Polymers and Textiles, Tsukuba). Cellulose nitrate membranes were obtained from Toyo Roshi and other reagents were of analytical-reagent grade. Preparation of enzyme membrane Choline oxidase (20 U) and/or acetylcholine esterase (50 U) dissolved in 0.2 ml of water were added to 1 g of a 11% aqueous solution of PVA-SbQ. The mixture was stirred magnetically at room temperature to give a homogeneous solution of PVA-SbQ. The mixture was spread on a cellulose nitrate membrane using a bar coater and
K. SODE
air dried at room temperature. The air-dried brane was exposed to normal fluorescent light for 2 h.
ET AL.
memroom
Measurement of enzyme activity by a spectrophotometric method Free and immobilized choline oxidase were assayed as follows. The reaction mixture [containing 0.2 ml of 0.1 M choline chloride solution, 1.3 ml of 0.1 M phosphate buffer (pH 8) 0.25 ml of 4-aminoantipyrine solution (0.3% w/v), 0.25 ml of phenol solution (0.2% w/v), 0.25 ml of peroxidase solution (2 U ml-‘) and 0.25 ml of distilled water] and an appropriate amount of the free or immobilized enzyme were incubated for 10 min at 35“C in a shaking machine. The reaction was stopped by adding 2.5 ml of ethanol. The absorbance at 480 nm was read with a spectrophotometer (Jasco Uvidec-510). Free and immobilized acetylcholine esterase were assayed as follows. The reaction mixture [containing 0.5 ml of 4 X 10e3 M acetylthiocholine solution, 1 ml of 0.1 M phosphate buffer (pH 8) and 0.5 ml of 4 x 10m3 M 5,5’-dithiobisnitrobenzoate] and the free or immobilized acetylcholine esterase were incubated for 10 min at 35 o C in a shaking machine. The reaction was stopped by adding 1 ml of ethanol. The absorbance at 405 nm was read with the spectrophotometer as above.
Preparation of the sensor The system consisted of three electrodes: an Ag/AgCl reference electrode, a Pt counter electrode and a Pt working electrode (0.3 cm2 area) covered with an immobilized enzyme membrane and a dialysis membrane. The activity of immobilized choline oxidase was 15 nmol min-’ cmp2, the activity of immobilized acetylcholine esterase was 17 nmol mm’ cme2 and the activities of co-immobilized choline oxidase and acetylcholine esterase were 13 and 8 nmol min ’ cm- 2, respectively (Table 1). A constant potential (+ 700 mV vs. Ag/AgCl) was applied with a potentiostat (Hokuto Denko) and the resulting current from the breakdown of hydrogen peroxide was measured with a strip-chart recorder (TOA Electronic Polyrecord).
AMPEROMETRIC
DETERMINATION
OF CHOLINE
AND
ACETYLCHOLINE
51
TABLE 1 Characteristics of choline and acetylcholine enzyme electrodes (CHO = choline esterase; ACE = acetylcholine esterase) Electrode
CHO membrane CHO membrane + ACE membrane Co-immobilized CHO + ACE
Membrane activity (nmol min-’ cm-‘) (spectrophotometric method)
Current (nA)
CHO
ACE
Choline (1 x 1O-4 M)
Acetylcholine (4 x 1O-4 M)
1s 1s
0 17
180 83
0 54
13
8
168
167
Procedure The sensor is immersed in 10 ml of 0.05 M phosphate buffer (pH 8) and maintained at 35OC with thermostated water circulation. When the current of the sensor has become constant, an appropriate volume of choline or acetylcholine solution is injected into the jacketed vessel. The current increases rapidly and reaches a steady state within 2 min when using a choline oxidase membrane, 4 min when using the two enzymes immobilized simultaneously and 8 min when using two different membranes.
a broad pH range. The highest activity for immobilized choline oxidase was obtained at pH 9.5-10 (carbonate buffer) whereas the free enzyme is optimum at pH 8. A shift of the pH optimum and broadening of enzyme activity after immobilization are common phenomena and have been explained by Goldstein et al. [12]. In this work, the shift in the pH optimum may be due to the effect of the photofunctional group of the polymer. Studies of stability at 35°C showed that the activity of the sensor decreases quickly at pH 6 and above pH 9.5 (Fig. 2). Therefore, the choline sensor should be used between pH 7 and 9.
RESULTS AND DISCUSSION
Acerylcholine sensor The acetylcholine sensor gave a linear calibration plot for the range 2.0 x 10e5-7.5 x 10P4 M with a slope and linear correlation coefficient of
Choline sensor The choline sensor gave a linear calibration plot for the range 2.5 X 10-6-1.5 x 10e4 M choline with a slope and linear correlation coefficient of 1.90 nA 1 pmol-’ and 0.9996, respectively. The concentration range is well below those obtained with oxygen electrode-based sensors [6,7]. Moreover, the linear range is larger than that obtained by Mascini and Moscone [9] with the same type of detection. The average response time was 2 min. We assume that the superior features of this sensor depend on the immobilization method. The high sensitivity and good linearity make the sensor useful for measurements in biological fluids. The effect of pH on the choline electrode (Fig. 1) was studied after 15 min of incubation of free or immobilized choline oxidase in different buffers at 35’C. Free choline oxidase was denaturated at pH lower than 6.5 or higher than 9, whereas the immobilized choline oxidase kept its activity over
I
I
PH
Fig. 1. Response of the choline sensor and free choline oxidase in different buffer solutions at several pH values to addition of choline giving a final concentration of 50 nmol 1-l for the choline sensor and 5 mmol 1-l for free choline oxidase. 0, 0, 0.05 M phosphate buffer; 0, n, 0.05 M pyrophosphate buffer; A, A, 0.05 M carbonate buffer. Open symbols for choline sensor and closed symbols for free choline oxidase.
K. SODE
52
372 nA 1 mmol-’ and 0.9998, respectively. This result is better than the practical range obtained with the pH electrode [3-51 and ion-selective electrodes [l]. This result was obtained by using a membrane containing co-immobilized choline oxidase and acetylcholine esterase. The sensitivity with co-immobilized enzymes is better than that obtained by using membranes with two enzymes immobilized separately (Table 1). This is due to poor diffusion of choline to the choline oxidase membrane across the nitrate cellulose membrane. Indeed, when the choline oxidase membrane is covered with an acetylcholine esterase membrane, the response to choline decreases by 54% (Table 1). The response time of the biosensor for acetylcholine was 4 min with co-immobilized enzymes. By using two enzyme membranes, however, 8 min are necessary to obtain a steady-state current. The results indicate that the sensitivity and response time obtained with a two-enzyme membrane are much superior to those obtained by using a two-membrane system. The ultimate aim of this work is to develop a novel enzyme sensor system for the determination of organophosphorus pesticides. This measurement is based on the inactivation of cholinesterase by such pesticides.
ET AL.
Time (days)
Fig. 3. Stability of the free (0) and immobilized (W) choline oxidase stored at 4” C in 0.05 M phosphate buffer (pH 8) and immobilized choline oxidase in a dry film (A) at 4 o C.
Stability Figure 3 shows the storage stability of the choline oxidase membrane. The immobilization of choline oxidase increases the stability of the enzyme. After storage for 60 days in 0.05 M phosphate buffer (pH 8) at 4°C the decrease in the activity was about 35%. When the enzyme film was stored in a dry state at 4”, no decrease in the activity was observed. Ichimura [ll] also reported that glucose amylase immobilized by PVA-SbQ shows no inactivation even after storage for 1 year in a dry state at 4OC. The stabilities of the acetylcholinesterase and co-immobilized enzyme membranes were similar (results not shown).
REFERENCES
Fig. 2. 35 o C. 0. pH n . pH
Stability of the choline sensor at different pH values at *. pH 6; *, pH 7; 0, pH 8 (0.05 M) phosphate buffer); 9 (0.05 M pyrophosphate buffer): A, pH 9.5; A, pH 10; 10.5 (0.05 M carbonate buffer).
1 A. Jaramillo, S. Lopez, J.B. Justice, J.D. Salomone and D.B. Neill, Anal. Chim. Acta, 146 (1983) 149. 2 M. Gotoh, E. Tamiya, M. Monoi, Y. Kagawa and I. Karube, Anal. Lett., 20 (1987) 857. 3 K. Gibson and G.G. Guilbault, Anal. Chim. Acta, 80 (1975) 245. 4 P. Durand, A. David and D. Thomas, Biochim. Biophys. Acta, 527 (1978) 277. 5 M.F. Suaud-Chagny and J.F. Pujol, Analusis, 13 (1985) 25. 6 K. Matsumuto, H. Seijo, I. Karube and S. Suzuki, Biotechnol. Bioeng., 22 (1980) 1071.
AMPEROMETRIC
DETERMINATION
OF CHOLINE
AND
53
ACETYLCHOLINE
7 L. Campanella, M. Mascini, G. Palleschi and M. Tomassetti, Clin. Chim. Acta, 151 (1985) 71. 8 L. Campanella, M. Tomassetti and M.P. Sammartino, Analyst, 113 (1988) 77. 9 M. Mascini and D. Moscone, Anal. Chim. Acta, 179 (1986) 439.
10 T. Yao, Anal. Chim. Acta, 153 (1983) 169. 11 K. Ichimura, J. Polym. Sci., 22 (1984) 2817. 12 L. Goldstein, Y. Levin and E. Katchalski. Biochemistry, (1964) 1913.
3