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Immobilization of alcohol dehydrogenase enzyme in a Langmuir-Blodgett film of stearic acid: its application as an ethanol sensor
138
Thin Solid Films, 239 (1994) 138-143
Immobilization of alcohol dehydrogenase enzyme in a Langmuir-Blodgett film of stearic acid: its application as an ethanol sensor Prabir Pal, Dipa Nandi and Tridibendra N. Misra* Spectroscopy Department, Indian Association for the Cultivation of Science, Calcutta 700 032 (India)
(Received June 6, 1993; accepted September 2, 1993)
Abstract A stable monolayer of alcohol dehydrogenase (ADH) enzyme has been prepared by spreading an aqueous solution of ADH on a water subphase containing stearic acid monolayer. This ADH-stearic acid monolayer has been successfully transferred onto a conducting polypyrrole-coated glass electrode by the Langmuir-Blodgett (LB) technique. ADH catalyses the reaction between the coenzyme//-NAD ÷ (fl form of oxidized nicotinamide adenine dinucleotide) and ethanol to produce NADH and acetaldehyde. Conducting polypyrrole acts as a mediator for transferring electrons produced on NADH oxidation. This LB-immobilized polypyrrole-mediated enzyme electrode can be used as an ethanol sensor. It shows better sensitivity than sensors consisting of ADH immobilized by chemical cross-linking with glutaraldehyde. This LB sensor can be used for low ethanol concentration (in the micro-molar range).
1. Introduction The use of Langmuir-Blodgett (LB) f i l l s for chemical sensing applications is very attractive. Their large surface-to-volume ratios suggest high sensitivity as detectors. For biosensor applications, LB films of enzymes show high selectivity for substrates. Recently glucose sensors [1,2] comprising LB films of glucose oxidase enzyme~lipid complex have been developed. It has been reported by many researchers [3,4] that the electrochemical response of glucose can be improved by using conducting polypyrrole combined with the glucose oxidase enzyme, giving a so-called proteinmodified electrode which may facilitate the electron transfer from the reduced enzyme to the electrode. An ethanol sensor using an electrode modified by immobilizing alcohol dehydrogenase ( A D H ) enzyme on a conducting organic charge transfer salt, N M P . T C N Q , by cross-linking has been developed [5,6]. In an electrochemical sensor it is important that electron transfer between the enzyme and electrode material be facilitated. Thus in our work we have concentrated on techniques to immobilize the enzyme that will promote proximity between the enzyme active site and the conducting surface of the electrode. In this paper we report the LB film technique of immobilization of A D H in the
*To whom all correspondenceshould be addressed.
0040-6090/94/$7.00 SSDI 0040-6090(93)02906-T
stearic acid matrix and its use as an ethanol sensor on a conducting polypyrrole-modified electrode. We also compare our results with that where A D H is immobilized by chemical cross-linking with glutaraldehyde.
2. Experimental details 2.1. Preparation o f conducting polypyrrole electrode
Polypyrrole film on SnO2-coated conducting glass electrode was prepared by the electrochemical oxidation of a solution containing pyrrole monomer. In the process, at first an extremely reactive n-radical cation is formed. It then reacts with neighbouring pyrrole species to produce a polymer that is predominantly ct,ct'-coupied, although some branching of the polymer chain is thought to take place by fl coupling. The resulting polymer has a net positive charge and it incorporates anions from solution during the film growth process. This makes the polymer film highly conductive. In the area of enzyme electrode biosensors, most of the activities were devoted towards improvement of surface morphology and adhesion of the conducting polypyrrole film to the substrate. Electropolymerization at high temperature gives a highly adhesive, fine-grain, uniform and less conducting film (10 -2 S cm-~), whereas electropolymerization at low temperature gives a less adhesive, coarse-grain, highly conducting (1 S cm-~) polymer film. A double-layer structure with a
© 1994--- Elsevier Sequoia. All rights reserved
P. Pal et aL / LB-immobilized ADH enzyme as ethanol sensor
thin high-temperature underlayer and a thick lowtemperature overlayer gives a highly adhesive and conducting polypyrrole film ideal for device application [7]. For polymerization, SnO2-coated conducting glass as a working electrode (WE), platinum foil as the counterelectrode (CE) and a saturated calomel electrode (SCE) as the reference electrode (RE) were used. 0.5 M pyrrole (Aldrich, purified by redistillation before use) and 0.5 M tetraethylammonium toluene 4-sulfonate were dissolved in 25 ml acetonitrile to form the electrolyte. The solution was deoxygenated by passing dry N2 gas for 1 h prior to the polymerization. A voltage of 0.65 V(SCE) was applied to the working electrode. A current of 0.5 mA for 30 s at temperature 70 °C yielded a thin (20-40 nm) polypyrrole sublayer (doped with CH3C6HaSO 3- ion). Then a thick (300-400 nm) overlayer was synthesized at low temperature (20 °C) by passing the same current for 5 min. The thickness was estimated from the electrode area and the amount of the polymer deposited. 2.2. Preparation o f the enzyme electrode by chemical cross -linking The conducting polypyrrole electrode was thoroughly rinsed in highly pure water (Millipore Q-plus) and was carefully dried. 10 units of ADH (EC 1.1.1.1., Sigma) was dissolved in 15 ]~1 of buffer solution (pH = 9.2) containing 0.6 M of tris buffer, and 0.01 M of flNAD ÷ (Sigma). This ADH solution was added to the electrode surface uniformly by drops (I0 pl) and dried carefully in air. Finally 1% (v/v) aqueous solution Of glutaraldehyde (10 /al) was added on this enzymeloaded electrode surface and dried again in air. The electrode was then washed carefully with purified water to remove excess glutaraldehyde and dried again. ADH was immobilized on the electrode surface both by adsorption in the micropores on the polymer surface and also by cross-linking with glutaraldehyde. The enzyme electrode was now ready for use. 2.3. Preparation o f the enzyme electrode by L B technique For LB film preparation we have used a computercontrolled Joyce-Loebl (UK) LB trough (model Langmuir Trough 4). Highly purified water by Milli-Q plus (Millipore Co. Ltd.) was used as a subphase (resistivity 18 M fl cm). 60/~1 of chloroform (spectrograde) solution (0.5 mg ml-~) of stearic acid (Sigma) was spread on the water subphase in a teflon-coated trough of area 430 cm 2. After 15 min (the time to evaporate the chloroform) an aliquot of freshly prepared aqueous solution of ADH was again spread on the surface containing stearic acid. The surface pressure was found to increase with time and the monolayer stabilized after 20 min, indicating the adsorption of ADH on the stearic acid film. The pressure-area (n-A) isotherm was recorded under a barrier compression/expansion
139
rate of 5 cm 2 m i n - ' . The compressed (35 mN m-~) monolayer was transferred onto the substrate (polypyrrole-coated conducting glass slide) by Y-type deposition technique. Dipping rate of the substrate was 5 mm min-~ and the drying time was 20 min. Five to ten layers were transferred onto the substrate by this technique. This LB-immobilized polypyrrole-modified electrode was subsequently used as a working electrode. 2.4. Detection o f ethanol Ethanol detection was done by the traditional amperometric method [5]. A three-electrode system consisting of the enzyme electrode as the working electrode, a platinum foil as the counter electrode and the saturated calomel electrode as the reference electrode were used in the electrochemical cell containing a buffer solution (20 ml, pH = 9.2) comprising 0.6 M of tris buffer, 0.4 M of lysine and 0.01 M of fl-NAD *. A potential of + 0.24 V(SCE) was applied to the enzyme electrode. A steadystate current value of few microamperes was attained in 30 min. Ethanol was added in the electrolyte in varied doses and the current enhancement was noted. The buffer solution was replaced after eight to ten measurements. For overnight preservation, the enzyme electrode was kept in the same buffer solution.
3. Results and discussion 3.1. Methodology o f the enzyme electrode
Alcohol dehydrogenase is an enzyme which catalyses the reaction between the coenzyme, fl-NAD + and ethanol to give NADH and acetaldehyde. CH3CH2OH+NAD + ADH C H 3 C H O + N A D H + H +
(1) Aldehyde produced is consumed by the lysine present in the buffer. The reduced coenzyme NADH is not strongly bound to ADH. At the working enzyme electrode the NADH produced is reoxidized to NAD + NADH -, NAD ÷ + H ÷ + 2e-
(2)
Figure 1 shows the cyclic voltammogram for NADH oxidation in the enzyme-loaded polypyrrole-mediated electrode. It is found that the oxidation of NADH proceeds rapidly at around + 0.24 V(SCE). The presence of CH3CrH4SO3 - ion in polypyrrole film facilitates the oxidation of NADH to N A D - and hence the oxidation potential of NADH is reduced to +0.24 V(SCE) from that of 1.1 V(SCE) in the naked electrode. The conducting polypyrrole electrode thus seems to work as a mediator. Such mediating activity of polypyrrole electrode has been reported [3]. Although it is possible to regenerate NAD ÷ from NADH at a naked electrode, this approach does have its disadvan-
140
P. Pal et al.
/ L B - i m m o b i l i z e d A D H e n z y m e as ethanol sensor
i,0F\\\\ 4O
I 0
I
I
'oh \\ \ \
I
0.1 0.2 0.3 Potentl(al vs. SCE ( V )
~ ol
Fig. 1. Cyclic voltammogram (100 mV s-1 sweep rate) of ADH enzyme-loaded polypyrrole-mediated electrode in a buffer solution (pH = 9.2) containing 0.6 M of tris buffer, 0.4 M lysine and 0.01 M of fl-NAD + and 1 mM ethanol.
tages. A large overvoltage is needed, i.e. 1.1 V(SCE) for the regeneration of N A D + at a platinum electrode, and prolonged usage results in fouling of the electrode surface owing to the accumulation of high-molecularweight oxidation products [8]. To overcome this difficulty we use the conducting polypyrrole surface which acts as a mediator. The current flowing at the working electrode is then related to the concentration of the ethanol present. 3.2. Langmuir-Blodgett film characterization Figure 2 shows the change in surface pressure with time after the spreading of A D H solution in the water subphase at different pH containing stearic acid monolayer at room temperature (20 °C). It is found that surface pressure increases with time and ultimately reaches saturation. This indicates the adsorption of A D H in the stearic acid monolayer on the subphase. Figure 2 also indicates that, with the same amount of A D H spreading, surface pressure increases more in the subphase with lower pH. This indicates that adsorption
pH =4.0
7
_~ 20 pH =6.0 in
,.. 10 -~ o"
pH ,7.5
ca
tn 0 5
I
10 Time (rain)
I
15
I
20
Fig. 2. Surface pressure rs. time plot after the spreading of ADH enzyme solution (60 #1 of 10 units per ml aqueous solution) in the water subphase at different pH containing stearic acid monolayer. The stearic acid monolayer is produced by the spreading of 60/~1 of 5 mg ml - ' chloroform solution in the trough of area 450 crn 2.
0.2
~\\ 0,3
",,,\ 0,4
A r e a / M o l e c u l e [ n m 2) Fig. 3. n-.4 isotherms (temperature = 20 °C, pH = 6) of pure stearic acid and stearic acid-ADH complexes with different amount of ADH: curve 1, pure stearic acid; curves 2 to 4, stearic acid-ADH complexes with increasing ADH amount. Area per molecule is calculated in terms of stearic acid molecules.
50
"E 40
-
"c
N 0
0.25 0.5 Area / Molecute (nm 2)
Fig. 4. Cyclic n A isotherms ( t e m p e r a t u r e = 2 0 ° C , pH = 6 ) of stearic acid-ADH complexes measured for two consecutive cycles.
of A D H with stearic acid is facilitated in the acidic medium. In acidic medium A D H becomes positively charged and the adsorption may take place by the electrostatic interaction of A D H with stearic acid. Denaturization of A D H also increases in the lower pH. Therefore, we concentrated our work at pH ~ 6 to keep the enzyme more active. Figure 3 shows the pressure-area (n-A) isotherm (temperature = 20 °C, pH = 6) of pure stearic acid and stearic acid-ADH complexes with different amounts of ADH. The area per molecule is calculated in terms of stearic acid molecules. It is found that area per molecule increases with the increase of A D H amount. Figure 4 shows the cyclic n - A isotherm for ADHstearic acid monolayer between 0 and 50 mN m surface pressure. It is found that almost the same path is traversed in each cycle. This indicates that the monolayer is stable. A clear hysteresis behaviour is observed between compression and expansion. This hysteresis
P. P a l et al.
/
3OO
0.~0
0.35
~ 1 0 m N / m
%
E o 200 ~
C
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141
L B - i m m o b i l i z e d A D H e n z y m e as ethanol sensor
pH =,6
-
,¢ pH =7'.5
0.30
E
10C 0
8
I
1
1000
2000
3000
Time (sec) 0.25 ~
50mN/m
I
0.2( 0
40
1
80 ADH amount (jUt)
Fig. 6. Area us. time plot of ADH-stearic acid monolayer compressed at a surface pressure of 35 mN m - J at different subphase pH.
I 120
Fig. 5. Area per molecule us. ADH amount spread in the subphase at different surface pressures. Concentration of ADH solution is 10 units per ml.
behaviour may arise because of the difference in the time taken for self-reorganization of the molecules in the monolayer during compression and expansion or because of the enzyme being squeezed off the monolayer during compression and its subsequent return upon the monolayer expanding. However, the expanding isotherm in Fig. 4, though similar to that of pure stearic acid in Fig. 3, is different. Isotherm 1 of Fig. 3 yields an area of 0.235 nm 2 per molecule at zero pressure in condensed phase, whereas the isotherm during expansion in Fig. 4 yields an area of 0.275 nm 2 per molecule. We therefore believe that the hysteresis in Fig. 4 arises because of self-reorganization of the enzyme molecule. The n - A isotherms also show that there is a transition from liquid to condensed phase at the surface pressure around 20 mN m-~ and this phase transition is reversible. Figure 5 shows the area per molecule vs. A D H amount plotted at different surface pressures. The results show that the change is linear, which indicates the proportional incorporation of A D H in stearic acid matrix. Figure 6 shows the area vs. time plot of ADH-stearic acid monolayer compressed at a pressure of 35 mN m - ~ at different pH of the subphase. It is observed that initially there is a small loss in the surface area, but after that the loss of area is negligible. The same behaviour is observed for both low and high pH of the subphase. This indicates that the monolayer is almost stable. Initial loss may be due to the self-organization of the molecules after the compression is stopped. The LB film is successfully transferred onto both the SnO2-coated conducting glass electrode and the polypyrrole-coated glass electrode by Y-type deposition at the surface pressure of 35 mN m-~. The transfer ratios are found to be 0.95 and 0.90 respectively. At
lower surface pressure (20 mN m - ' ) the film can be transferred but the transfer ratio is small (0.50). Indeed, both the conducting glass electrode and the polypyrrole electrode are unlikely to have a highly regular surface to accommodate distinct LB layers. The satisfactory transfer ratio, however, suggests that this can be achieved at high surface pressure, when the film acts more as a solid layer than as a molecular film. Five to ten layers of ADH-stearic acid LB film is deposited on the bare and the polypyrrole-coated conducting glass surfaces to get the LB-immobilized enzyme electrode. 3.3. S e n s o r c h a r a c t e r i z a t i o n Three different arrangements of the biosensor on the electrode were used: 1. polypyrrole-modified electrode with ADH immobilized by chemical cross-linking with glutaraldehyde, 2. polypyrrole-modified electrode with ADH-immobilized stearic acid LB film, and 3. bare conducting glass electrode with ADH-immobilized stearic acid LB film. The third arrangement needed large overvoltage (1.1 V(SCE)) for the regeneration of NAD + at the electrode and thus electrode poisoning occurred and the specificity for ethanol detection was lost. Figure 7 shows the schematic diagram of the first two types of arrangements together with the response characteristics of the sensors against ethanol concentration. It is found that the LB-modified sensor shows better response for low ethanol concentration, though the amount of ADH present in the LB film is much less than that present in the sensor immobilized by chemical cross-linking. This indicates that in the LB film the electron transfer to the electrode is more efficient. That the LB electrode, in spite of less enzyme loading, gives higher response for a specific substrate concentration can be understood from the fact that the substrate principally interacts with the surface layer of the electrode. More ADH and fl-NAD- molecules are at the surface layer in the LB electrode, whereas in the glutaraldehyde cross-linked polypyrrole electrode more of
142
P. Pal et al. / LB-immobilized A D H enzyme as ethanol sensor
20 LB immo b i [ i $ ~ . ' ; ' . - ~ l
......
t'.* ~
~, /
~ 1 0 ._.PoIypyr role
H-gtutaratdt, hy
/ 0, ~ . 0
-o- - - -
PO 2, ~
~
------
1
25
50
75
Ethcmot (./a mole)
Fig. 7. Response characteristics of the two types of sensors against ethanol concentration (in micro-molar range): curve 1, sensor containing ADH immobilizedby chemical cross-linkingwith glutaraldehydeand; curve 2, sensor containing five layers of ADH-stearic acid LB films. Insets show the schematic diagrams of these sensors.
200 ,
Ethanot (.juM) t, O0 ,
~12
600
3rd doy
/ / ; S;
o..
0
25
50
o
E#.honot (juM)
Fig. 8. Response characteristics and stability of the sensors containing ADH ( ) immobilized by chemical cross-linking, stored at 0 °C in the buffer solution, and ( . . . . . . ) immobilized by LB trapping, stored at 0 :'C in the dry condition.
these are sealed deep into the polymer film. Further, the LB film being an order of magnitude thinner, the electron transfer from the reduced coenzyme to the electrode material is faster in this case. Excellent linearity is observed at low ethanol concentration. For the LB sensor it is up to 40 # M and for chemical cross-linked A D H sensor it is up to 500 # M (as observed from Fig. 8). This linear behaviour is expected when a substrate diffusion membrane is incorporated and the range of linearity can be controlled by changing the permeability of the membrane [9]. In our case, though we have not used any diffusion-limiting membrane, the linearity may arise because of the self-
diffusion of ethanol. At higher ethanol concentration, saturation behaviour is observed. This saturation behaviour can be explained by the saturated enzyme kinetics [10] when the reduction of N A D + to N A D H does not depend on ethanol concentration but is governed by the enzyme activity, as at high ethanol concentration the number of enzymatic reaction sites is less than the number of ethanol molecules present. Response time of these sensors is almost the same and very fast (30 s). Figure 8 shows the response characteristics and stability of the two sensors. It is found that, where A D H is immobilized by chemical cross-linking, the sensitivity is reduced by 25% after three days at 400 /zM concentration and after five days there is practically no response. The LB sensor, on the other hand, remains active even after ten days. It is known that A D H is very unstable. The degradation may be due to leaching out or denaturization of the enzyme or isomerization of the coenzyme.
4. Conclusions
1. Alcohol dehydrogenase is successfully immobilized in the stearic acid matrix by the LB technique. It is found that A D H is quite active after immobilization. 2. Conducting polypyrrole acts as a mediator for N A D H oxidation. The oxidation potential is reduced to + 0.24 V owing to incorporation of polypyrrole-mediated electrode. 3. The LB-modified sensor shows better response compared to the sensor where A D H is immobilized by chemical cross-linking with glutaraldehyde. This LB sensor can be used for low ethanol concentration.
P. Pal et al. [ LB-immobilized ADH enzyme as ethanol sensor
4. T h e chemically cross-linked A D H sensor is quite stable for three days. A t longer times it degrades rapidly. The LB sensor r e m a i n s active even after ten days.
Acknowledgment The a u t h o r s are t h a n k f u l to the D e p a r t m e n t o f Science a n d T e c h n o l o g y , G o v e r n m e n t o f India, for financial s u p p o r t t h r o u g h g r a n t no. F 1 2 1 / T S D [ D S T / 8 9 .
Reference 1 Y. Okahata, T. Tsuruta, K. Ijiro and K. Ariga, Thin Solid Films, 180 (1989) 65.
143
2 G. G. Guilbault, Analytical Uses o f Immobilized Enzymes, Dekker, New York, 1984. 3.J.R. Li, M. Cai, T. F. Chen and L. Jiang, Thin Solid Films, 180 (1989) 205. 4 H. Shinohara, M. Aizawa and H. Shirakawa, J. Chem. Soc. Jpn., 3 (1986) 465. 5 W. J. Albery and P. N. Bartlett, J. Chem. Soc., Chem. Commun., (1984) 234. 6 J. J. Kulys, Enzyme Microb. Technol., 3 (1981) 342. 7 A. F. Diaz and B. Hall, I B M J. Res. Dev., 27(1983) 477. 8 M. F. Cardosi and A. P. F. Turner, in A. P. F. Turner, I. Karube and G. S. Wilson (eds.), Biosensors, Fundamentals and Applications, Oxford University Press, New York, 1987, p. 265. 9 W. J. Albery and D. H. Craston, in A. P. F. Turner, I. Karube and G. S. Wilson (eds.), Biosensors, Fundamentals and Applications, Oxford University Press, New York, 1987, p. 193. 10 P. N. Bartlett, in A. G. E. Cass (ed.), Biosenso~'s: A Practical Approach, Oxford University Press, New York, 1990, p. 47.
Thin Solid Films, 239 (1994) 138-143
Immobilization of alcohol dehydrogenase enzyme in a Langmuir-Blodgett film of stearic acid: its application as an ethanol sensor Prabir Pal, Dipa Nandi and Tridibendra N. Misra* Spectroscopy Department, Indian Association for the Cultivation of Science, Calcutta 700 032 (India)
(Received June 6, 1993; accepted September 2, 1993)
Abstract A stable monolayer of alcohol dehydrogenase (ADH) enzyme has been prepared by spreading an aqueous solution of ADH on a water subphase containing stearic acid monolayer. This ADH-stearic acid monolayer has been successfully transferred onto a conducting polypyrrole-coated glass electrode by the Langmuir-Blodgett (LB) technique. ADH catalyses the reaction between the coenzyme//-NAD ÷ (fl form of oxidized nicotinamide adenine dinucleotide) and ethanol to produce NADH and acetaldehyde. Conducting polypyrrole acts as a mediator for transferring electrons produced on NADH oxidation. This LB-immobilized polypyrrole-mediated enzyme electrode can be used as an ethanol sensor. It shows better sensitivity than sensors consisting of ADH immobilized by chemical cross-linking with glutaraldehyde. This LB sensor can be used for low ethanol concentration (in the micro-molar range).
1. Introduction The use of Langmuir-Blodgett (LB) f i l l s for chemical sensing applications is very attractive. Their large surface-to-volume ratios suggest high sensitivity as detectors. For biosensor applications, LB films of enzymes show high selectivity for substrates. Recently glucose sensors [1,2] comprising LB films of glucose oxidase enzyme~lipid complex have been developed. It has been reported by many researchers [3,4] that the electrochemical response of glucose can be improved by using conducting polypyrrole combined with the glucose oxidase enzyme, giving a so-called proteinmodified electrode which may facilitate the electron transfer from the reduced enzyme to the electrode. An ethanol sensor using an electrode modified by immobilizing alcohol dehydrogenase ( A D H ) enzyme on a conducting organic charge transfer salt, N M P . T C N Q , by cross-linking has been developed [5,6]. In an electrochemical sensor it is important that electron transfer between the enzyme and electrode material be facilitated. Thus in our work we have concentrated on techniques to immobilize the enzyme that will promote proximity between the enzyme active site and the conducting surface of the electrode. In this paper we report the LB film technique of immobilization of A D H in the
*To whom all correspondenceshould be addressed.
0040-6090/94/$7.00 SSDI 0040-6090(93)02906-T
stearic acid matrix and its use as an ethanol sensor on a conducting polypyrrole-modified electrode. We also compare our results with that where A D H is immobilized by chemical cross-linking with glutaraldehyde.
2. Experimental details 2.1. Preparation o f conducting polypyrrole electrode
Polypyrrole film on SnO2-coated conducting glass electrode was prepared by the electrochemical oxidation of a solution containing pyrrole monomer. In the process, at first an extremely reactive n-radical cation is formed. It then reacts with neighbouring pyrrole species to produce a polymer that is predominantly ct,ct'-coupied, although some branching of the polymer chain is thought to take place by fl coupling. The resulting polymer has a net positive charge and it incorporates anions from solution during the film growth process. This makes the polymer film highly conductive. In the area of enzyme electrode biosensors, most of the activities were devoted towards improvement of surface morphology and adhesion of the conducting polypyrrole film to the substrate. Electropolymerization at high temperature gives a highly adhesive, fine-grain, uniform and less conducting film (10 -2 S cm-~), whereas electropolymerization at low temperature gives a less adhesive, coarse-grain, highly conducting (1 S cm-~) polymer film. A double-layer structure with a
© 1994--- Elsevier Sequoia. All rights reserved
P. Pal et aL / LB-immobilized ADH enzyme as ethanol sensor
thin high-temperature underlayer and a thick lowtemperature overlayer gives a highly adhesive and conducting polypyrrole film ideal for device application [7]. For polymerization, SnO2-coated conducting glass as a working electrode (WE), platinum foil as the counterelectrode (CE) and a saturated calomel electrode (SCE) as the reference electrode (RE) were used. 0.5 M pyrrole (Aldrich, purified by redistillation before use) and 0.5 M tetraethylammonium toluene 4-sulfonate were dissolved in 25 ml acetonitrile to form the electrolyte. The solution was deoxygenated by passing dry N2 gas for 1 h prior to the polymerization. A voltage of 0.65 V(SCE) was applied to the working electrode. A current of 0.5 mA for 30 s at temperature 70 °C yielded a thin (20-40 nm) polypyrrole sublayer (doped with CH3C6HaSO 3- ion). Then a thick (300-400 nm) overlayer was synthesized at low temperature (20 °C) by passing the same current for 5 min. The thickness was estimated from the electrode area and the amount of the polymer deposited. 2.2. Preparation o f the enzyme electrode by chemical cross -linking The conducting polypyrrole electrode was thoroughly rinsed in highly pure water (Millipore Q-plus) and was carefully dried. 10 units of ADH (EC 1.1.1.1., Sigma) was dissolved in 15 ]~1 of buffer solution (pH = 9.2) containing 0.6 M of tris buffer, and 0.01 M of flNAD ÷ (Sigma). This ADH solution was added to the electrode surface uniformly by drops (I0 pl) and dried carefully in air. Finally 1% (v/v) aqueous solution Of glutaraldehyde (10 /al) was added on this enzymeloaded electrode surface and dried again in air. The electrode was then washed carefully with purified water to remove excess glutaraldehyde and dried again. ADH was immobilized on the electrode surface both by adsorption in the micropores on the polymer surface and also by cross-linking with glutaraldehyde. The enzyme electrode was now ready for use. 2.3. Preparation o f the enzyme electrode by L B technique For LB film preparation we have used a computercontrolled Joyce-Loebl (UK) LB trough (model Langmuir Trough 4). Highly purified water by Milli-Q plus (Millipore Co. Ltd.) was used as a subphase (resistivity 18 M fl cm). 60/~1 of chloroform (spectrograde) solution (0.5 mg ml-~) of stearic acid (Sigma) was spread on the water subphase in a teflon-coated trough of area 430 cm 2. After 15 min (the time to evaporate the chloroform) an aliquot of freshly prepared aqueous solution of ADH was again spread on the surface containing stearic acid. The surface pressure was found to increase with time and the monolayer stabilized after 20 min, indicating the adsorption of ADH on the stearic acid film. The pressure-area (n-A) isotherm was recorded under a barrier compression/expansion
139
rate of 5 cm 2 m i n - ' . The compressed (35 mN m-~) monolayer was transferred onto the substrate (polypyrrole-coated conducting glass slide) by Y-type deposition technique. Dipping rate of the substrate was 5 mm min-~ and the drying time was 20 min. Five to ten layers were transferred onto the substrate by this technique. This LB-immobilized polypyrrole-modified electrode was subsequently used as a working electrode. 2.4. Detection o f ethanol Ethanol detection was done by the traditional amperometric method [5]. A three-electrode system consisting of the enzyme electrode as the working electrode, a platinum foil as the counter electrode and the saturated calomel electrode as the reference electrode were used in the electrochemical cell containing a buffer solution (20 ml, pH = 9.2) comprising 0.6 M of tris buffer, 0.4 M of lysine and 0.01 M of fl-NAD *. A potential of + 0.24 V(SCE) was applied to the enzyme electrode. A steadystate current value of few microamperes was attained in 30 min. Ethanol was added in the electrolyte in varied doses and the current enhancement was noted. The buffer solution was replaced after eight to ten measurements. For overnight preservation, the enzyme electrode was kept in the same buffer solution.
3. Results and discussion 3.1. Methodology o f the enzyme electrode
Alcohol dehydrogenase is an enzyme which catalyses the reaction between the coenzyme, fl-NAD + and ethanol to give NADH and acetaldehyde. CH3CH2OH+NAD + ADH C H 3 C H O + N A D H + H +
(1) Aldehyde produced is consumed by the lysine present in the buffer. The reduced coenzyme NADH is not strongly bound to ADH. At the working enzyme electrode the NADH produced is reoxidized to NAD + NADH -, NAD ÷ + H ÷ + 2e-
(2)
Figure 1 shows the cyclic voltammogram for NADH oxidation in the enzyme-loaded polypyrrole-mediated electrode. It is found that the oxidation of NADH proceeds rapidly at around + 0.24 V(SCE). The presence of CH3CrH4SO3 - ion in polypyrrole film facilitates the oxidation of NADH to N A D - and hence the oxidation potential of NADH is reduced to +0.24 V(SCE) from that of 1.1 V(SCE) in the naked electrode. The conducting polypyrrole electrode thus seems to work as a mediator. Such mediating activity of polypyrrole electrode has been reported [3]. Although it is possible to regenerate NAD ÷ from NADH at a naked electrode, this approach does have its disadvan-
140
P. Pal et al.
/ L B - i m m o b i l i z e d A D H e n z y m e as ethanol sensor
i,0F\\\\ 4O
I 0
I
I
'oh \\ \ \
I
0.1 0.2 0.3 Potentl(al vs. SCE ( V )
~ ol
Fig. 1. Cyclic voltammogram (100 mV s-1 sweep rate) of ADH enzyme-loaded polypyrrole-mediated electrode in a buffer solution (pH = 9.2) containing 0.6 M of tris buffer, 0.4 M lysine and 0.01 M of fl-NAD + and 1 mM ethanol.
tages. A large overvoltage is needed, i.e. 1.1 V(SCE) for the regeneration of N A D + at a platinum electrode, and prolonged usage results in fouling of the electrode surface owing to the accumulation of high-molecularweight oxidation products [8]. To overcome this difficulty we use the conducting polypyrrole surface which acts as a mediator. The current flowing at the working electrode is then related to the concentration of the ethanol present. 3.2. Langmuir-Blodgett film characterization Figure 2 shows the change in surface pressure with time after the spreading of A D H solution in the water subphase at different pH containing stearic acid monolayer at room temperature (20 °C). It is found that surface pressure increases with time and ultimately reaches saturation. This indicates the adsorption of A D H in the stearic acid monolayer on the subphase. Figure 2 also indicates that, with the same amount of A D H spreading, surface pressure increases more in the subphase with lower pH. This indicates that adsorption
pH =4.0
7
_~ 20 pH =6.0 in
,.. 10 -~ o"
pH ,7.5
ca
tn 0 5
I
10 Time (rain)
I
15
I
20
Fig. 2. Surface pressure rs. time plot after the spreading of ADH enzyme solution (60 #1 of 10 units per ml aqueous solution) in the water subphase at different pH containing stearic acid monolayer. The stearic acid monolayer is produced by the spreading of 60/~1 of 5 mg ml - ' chloroform solution in the trough of area 450 crn 2.
0.2
~\\ 0,3
",,,\ 0,4
A r e a / M o l e c u l e [ n m 2) Fig. 3. n-.4 isotherms (temperature = 20 °C, pH = 6) of pure stearic acid and stearic acid-ADH complexes with different amount of ADH: curve 1, pure stearic acid; curves 2 to 4, stearic acid-ADH complexes with increasing ADH amount. Area per molecule is calculated in terms of stearic acid molecules.
50
"E 40
-
"c
N 0
0.25 0.5 Area / Molecute (nm 2)
Fig. 4. Cyclic n A isotherms ( t e m p e r a t u r e = 2 0 ° C , pH = 6 ) of stearic acid-ADH complexes measured for two consecutive cycles.
of A D H with stearic acid is facilitated in the acidic medium. In acidic medium A D H becomes positively charged and the adsorption may take place by the electrostatic interaction of A D H with stearic acid. Denaturization of A D H also increases in the lower pH. Therefore, we concentrated our work at pH ~ 6 to keep the enzyme more active. Figure 3 shows the pressure-area (n-A) isotherm (temperature = 20 °C, pH = 6) of pure stearic acid and stearic acid-ADH complexes with different amounts of ADH. The area per molecule is calculated in terms of stearic acid molecules. It is found that area per molecule increases with the increase of A D H amount. Figure 4 shows the cyclic n - A isotherm for ADHstearic acid monolayer between 0 and 50 mN m surface pressure. It is found that almost the same path is traversed in each cycle. This indicates that the monolayer is stable. A clear hysteresis behaviour is observed between compression and expansion. This hysteresis
P. P a l et al.
/
3OO
0.~0
0.35
~ 1 0 m N / m
%
E o 200 ~
C
g
141
L B - i m m o b i l i z e d A D H e n z y m e as ethanol sensor
pH =,6
-
,¢ pH =7'.5
0.30
E
10C 0
8
I
1
1000
2000
3000
Time (sec) 0.25 ~
50mN/m
I
0.2( 0
40
1
80 ADH amount (jUt)
Fig. 6. Area us. time plot of ADH-stearic acid monolayer compressed at a surface pressure of 35 mN m - J at different subphase pH.
I 120
Fig. 5. Area per molecule us. ADH amount spread in the subphase at different surface pressures. Concentration of ADH solution is 10 units per ml.
behaviour may arise because of the difference in the time taken for self-reorganization of the molecules in the monolayer during compression and expansion or because of the enzyme being squeezed off the monolayer during compression and its subsequent return upon the monolayer expanding. However, the expanding isotherm in Fig. 4, though similar to that of pure stearic acid in Fig. 3, is different. Isotherm 1 of Fig. 3 yields an area of 0.235 nm 2 per molecule at zero pressure in condensed phase, whereas the isotherm during expansion in Fig. 4 yields an area of 0.275 nm 2 per molecule. We therefore believe that the hysteresis in Fig. 4 arises because of self-reorganization of the enzyme molecule. The n - A isotherms also show that there is a transition from liquid to condensed phase at the surface pressure around 20 mN m-~ and this phase transition is reversible. Figure 5 shows the area per molecule vs. A D H amount plotted at different surface pressures. The results show that the change is linear, which indicates the proportional incorporation of A D H in stearic acid matrix. Figure 6 shows the area vs. time plot of ADH-stearic acid monolayer compressed at a pressure of 35 mN m - ~ at different pH of the subphase. It is observed that initially there is a small loss in the surface area, but after that the loss of area is negligible. The same behaviour is observed for both low and high pH of the subphase. This indicates that the monolayer is almost stable. Initial loss may be due to the self-organization of the molecules after the compression is stopped. The LB film is successfully transferred onto both the SnO2-coated conducting glass electrode and the polypyrrole-coated glass electrode by Y-type deposition at the surface pressure of 35 mN m-~. The transfer ratios are found to be 0.95 and 0.90 respectively. At
lower surface pressure (20 mN m - ' ) the film can be transferred but the transfer ratio is small (0.50). Indeed, both the conducting glass electrode and the polypyrrole electrode are unlikely to have a highly regular surface to accommodate distinct LB layers. The satisfactory transfer ratio, however, suggests that this can be achieved at high surface pressure, when the film acts more as a solid layer than as a molecular film. Five to ten layers of ADH-stearic acid LB film is deposited on the bare and the polypyrrole-coated conducting glass surfaces to get the LB-immobilized enzyme electrode. 3.3. S e n s o r c h a r a c t e r i z a t i o n Three different arrangements of the biosensor on the electrode were used: 1. polypyrrole-modified electrode with ADH immobilized by chemical cross-linking with glutaraldehyde, 2. polypyrrole-modified electrode with ADH-immobilized stearic acid LB film, and 3. bare conducting glass electrode with ADH-immobilized stearic acid LB film. The third arrangement needed large overvoltage (1.1 V(SCE)) for the regeneration of NAD + at the electrode and thus electrode poisoning occurred and the specificity for ethanol detection was lost. Figure 7 shows the schematic diagram of the first two types of arrangements together with the response characteristics of the sensors against ethanol concentration. It is found that the LB-modified sensor shows better response for low ethanol concentration, though the amount of ADH present in the LB film is much less than that present in the sensor immobilized by chemical cross-linking. This indicates that in the LB film the electron transfer to the electrode is more efficient. That the LB electrode, in spite of less enzyme loading, gives higher response for a specific substrate concentration can be understood from the fact that the substrate principally interacts with the surface layer of the electrode. More ADH and fl-NAD- molecules are at the surface layer in the LB electrode, whereas in the glutaraldehyde cross-linked polypyrrole electrode more of
142
P. Pal et al. / LB-immobilized A D H enzyme as ethanol sensor
20 LB immo b i [ i $ ~ . ' ; ' . - ~ l
......
t'.* ~
~, /
~ 1 0 ._.PoIypyr role
H-gtutaratdt, hy
/ 0, ~ . 0
-o- - - -
PO 2, ~
~
------
1
25
50
75
Ethcmot (./a mole)
Fig. 7. Response characteristics of the two types of sensors against ethanol concentration (in micro-molar range): curve 1, sensor containing ADH immobilizedby chemical cross-linkingwith glutaraldehydeand; curve 2, sensor containing five layers of ADH-stearic acid LB films. Insets show the schematic diagrams of these sensors.
200 ,
Ethanot (.juM) t, O0 ,
~12
600
3rd doy
/ / ; S;
o..
0
25
50
o
E#.honot (juM)
Fig. 8. Response characteristics and stability of the sensors containing ADH ( ) immobilized by chemical cross-linking, stored at 0 °C in the buffer solution, and ( . . . . . . ) immobilized by LB trapping, stored at 0 :'C in the dry condition.
these are sealed deep into the polymer film. Further, the LB film being an order of magnitude thinner, the electron transfer from the reduced coenzyme to the electrode material is faster in this case. Excellent linearity is observed at low ethanol concentration. For the LB sensor it is up to 40 # M and for chemical cross-linked A D H sensor it is up to 500 # M (as observed from Fig. 8). This linear behaviour is expected when a substrate diffusion membrane is incorporated and the range of linearity can be controlled by changing the permeability of the membrane [9]. In our case, though we have not used any diffusion-limiting membrane, the linearity may arise because of the self-
diffusion of ethanol. At higher ethanol concentration, saturation behaviour is observed. This saturation behaviour can be explained by the saturated enzyme kinetics [10] when the reduction of N A D + to N A D H does not depend on ethanol concentration but is governed by the enzyme activity, as at high ethanol concentration the number of enzymatic reaction sites is less than the number of ethanol molecules present. Response time of these sensors is almost the same and very fast (30 s). Figure 8 shows the response characteristics and stability of the two sensors. It is found that, where A D H is immobilized by chemical cross-linking, the sensitivity is reduced by 25% after three days at 400 /zM concentration and after five days there is practically no response. The LB sensor, on the other hand, remains active even after ten days. It is known that A D H is very unstable. The degradation may be due to leaching out or denaturization of the enzyme or isomerization of the coenzyme.
4. Conclusions
1. Alcohol dehydrogenase is successfully immobilized in the stearic acid matrix by the LB technique. It is found that A D H is quite active after immobilization. 2. Conducting polypyrrole acts as a mediator for N A D H oxidation. The oxidation potential is reduced to + 0.24 V owing to incorporation of polypyrrole-mediated electrode. 3. The LB-modified sensor shows better response compared to the sensor where A D H is immobilized by chemical cross-linking with glutaraldehyde. This LB sensor can be used for low ethanol concentration.
P. Pal et al. [ LB-immobilized ADH enzyme as ethanol sensor
4. T h e chemically cross-linked A D H sensor is quite stable for three days. A t longer times it degrades rapidly. The LB sensor r e m a i n s active even after ten days.
Acknowledgment The a u t h o r s are t h a n k f u l to the D e p a r t m e n t o f Science a n d T e c h n o l o g y , G o v e r n m e n t o f India, for financial s u p p o r t t h r o u g h g r a n t no. F 1 2 1 / T S D [ D S T / 8 9 .
Reference 1 Y. Okahata, T. Tsuruta, K. Ijiro and K. Ariga, Thin Solid Films, 180 (1989) 65.
143
2 G. G. Guilbault, Analytical Uses o f Immobilized Enzymes, Dekker, New York, 1984. 3.J.R. Li, M. Cai, T. F. Chen and L. Jiang, Thin Solid Films, 180 (1989) 205. 4 H. Shinohara, M. Aizawa and H. Shirakawa, J. Chem. Soc. Jpn., 3 (1986) 465. 5 W. J. Albery and P. N. Bartlett, J. Chem. Soc., Chem. Commun., (1984) 234. 6 J. J. Kulys, Enzyme Microb. Technol., 3 (1981) 342. 7 A. F. Diaz and B. Hall, I B M J. Res. Dev., 27(1983) 477. 8 M. F. Cardosi and A. P. F. Turner, in A. P. F. Turner, I. Karube and G. S. Wilson (eds.), Biosensors, Fundamentals and Applications, Oxford University Press, New York, 1987, p. 265. 9 W. J. Albery and D. H. Craston, in A. P. F. Turner, I. Karube and G. S. Wilson (eds.), Biosensors, Fundamentals and Applications, Oxford University Press, New York, 1987, p. 193. 10 P. N. Bartlett, in A. G. E. Cass (ed.), Biosenso~'s: A Practical Approach, Oxford University Press, New York, 1990, p. 47.