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Anal. Chem. lQ92, 64, 1051-1055
Dual-Enzyme Fiber-optic Biosensor for Glutamate Based on Reduced Nicotinamide Adenine Dinucleotide Luminescence Ae-June Wang and Mark A. Arnold*
Department of Chemistry, University of Iowa, Iowa City, Iowa 52242
Response characterlstics are presented for a dual-enzyme flberoptlc biosensor for glutamate. An enzyme layer composed of glutamate dehydrogenase (GDH) and glutamatepyruvate transaminase (OPT) Is used to produce reduced nkotlnamlde adenlne dlnucleotide (NADH) at the tlp of a flber-optlc probe. NADH luminescence is monitored through thk probe and the measured fluorescence intensity k related to the Concentration of glutamate. GDH catalyzes the formation of NADH, and OPT drlves the GDH reaction by removing a reactlon product and regeneratlng glutamate. Op tbnal response b obtalned in a pH 7.4 Tris-HCi buffer maintalned at 25 OC In the presence of 4 mM NAD' and 10 mM L-alanlm. The temperature proflle reveals a strong negative temperature effect whlch Is attributed to the temperature dependency of NADH lumlneclcence. Under optlmal condlHans, the "or 8ondtMly Is 0.127 W p M ovw the 1-10 pM concentratlon range, the detectlon llmit k 0.13 pM, and response timer range from 4 to 8 min. The semi response k stable for 12 day8 when stored at 4 OC. Sectlvlty for glutamate k exover most of the common amino adds as well as ascorbk acid, uric acid, taurine, and GABA. Only slight were observed for glutamhe and lydne. The effect of ammonia on the glutamate response was found to be minknal at total ammonla nitrogen concentrations as high as 200 MY.
INTRODUCTION Considerable attention has been given to the development of a glutamate-selective biosensor. The need to monitor glutamate in the food industry,'" biorea&rs,4 and biomedical research is the major impetus for this activity. We are interested in biosensors capable of measuring glutamate in the extracellular fluid during neurochemical experiments. Such a sensor must possess sufficient selectivity to measure glutamate accurately in the 1-10 pM concentration range under physiological condition^.^ Our initial effort was an evaluation of a biosensor constructed by immobilizing glutamate decarboxylase at the tip of a carbon dioxide gas-sensing electrode.6 This electrode system was judged unsuitable for neurochemical applications because of an insufficient detection limit, long response times, and poor selectivity. In addition, this glutamate biosensor does not respond at pH values greater than 5.5, which renders it incompatible with physiological measurements. Glutamate oxidase has been used for the construction of glutamate biosensors based on either the consumption of oxygen4s7or the production of hydrogen peroxide.*1° The analytical merits of the amperometric version of this biwnsor have been established by Guilbault and co-workers with an eye on potential neurochemical applications.' Sensors based on amperometric oxygen detection lacked the detection limit needed for such measurements. Sensors based on hydrogen peroxide detection suffered from interference by oxidizable endogenous species such as ascorbate and urate. Regardless 0003-2700/92/0364-1051$03.00/0
of the detection mode, any bioseneor based on an oxidasecatalyzed reaction would be susceptible to fluctuations in oxygen tension. This latter point is a major concern for neurochemicalmeasurementa where changes in oxygen tension have been measured during synaptic events." Fiber-optic biosensors (FOBS'S) based on the fluorometric detection of reduced nicotinamide adenine dinucleotide (NADH)12J3offer an alternative approach for a glutamate biosensor. The concept of this type of biosensor has been demonstrated previously for lactate and pyruvate with a FOBS constructed by immobilizing lactate dehydrogenase at the tip of a bifurcated fiber-optic probe.14 NADH luminescence is measured through this fiber-optic probe and the magnitude of the resulting fluorescence signal is related to the bulk concentrations of these enzyme substrates. A FOBS for glutamate can be constructed in an analogous fashion by immobilzing glutamate dehydrogenase (GDH) at the probe tip and monitoring the production of NADH from the following reaction: glutamate NAD+ a-ketoglutarate NH4+ NADH
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GDH has been shown to possess sufficient selectivity for neurochemical measurements of glutamate,'m6 which suggests that the correapondingFOBS would also possess the selectivity required for such measurements. The major limitation of using this reaction for a biosensor is the large thermodynamic barrier associated with the production of NADH. The thermodynamically favored products of the GDH reaction are glutamate and NAD'. The equilibrium constant with NADH as the reaction product is approximately 10-14.17Such a large thermodynamic barrier suggests that an approach based on GDH done would be unsuitable for glutamate measurements in the micromolar concentration range. We have constructed and evaluated a novel dual-enzyme NADH-based FOBS for glutamate that combinea the activities of GDH and glutamate-pyruvate transaminase (GPT) at the tip of the fiber-optic probe. The dual-enzyme reaction scheme is as follows: glutamate NAD+ a-ketoglutarate NH4+ NADH a-ketoglutarate alanine pyruvate glutamate In this scheme, GPT drives the GDH reaction in the desired direction by removing a reaction product and regenerating glutamate. The addition of GPT significantly improves both the sensitivity and detection limit for glutamate. The optimal operating conditions and analytical response characteristics are presented for this GDH/GPT dual-enzyme NADH-based FOBS for glutamate. EXPERIMENTAL SECTION Apparatus and Reagents. The optical arrangement used for the fluorescence measurements is shown in Figure 1. A 100-W tungsten-halogen lamp (Oriel, Model 77825) with a constantvoltage transformer (Oriel, Model 6393) was used as the source. The excitation radiation was isolated by passing the source radiation through an infrared blocking filter and a 50-nm band-pass
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@ 1992 American Chemical Society
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ANALYTICAL CHEMISTRY, VOL. 64, NO. 9, MAY 1, 1992
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Fiber-Optic Probe
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t K Figure 1. Schematic diagram of the optical arrangement: (A) source, (B) I R blocking filter, (C) 340nm filter, (D) chopper, (E) lockin amplifier, (F)450-nm filter, (G) power supply, (H) photomultiplier tube, (I) sensor unit, (J) water bath, (K) cross-sectional view of the fiber-optic probe.
filter with maximum transmission at 340 nm (Earling, UG11). The excitation radiation then passed through a mechanical chopper (Stanford Research Systems, Model SR 540) set at 390 Hz and was then focused onto the central fiber of a custom-made bifurcated fiber-optic probe. This probe consisted of a center 1000-pm-diametersilica fiber surrounded by a ring of 12 484-pm plastic fibers (see Figure 1).The total diameter of the fiber probe was 3 mm. The excitation radiation was directed to the biocatalytic layer by the silica fiber, and NADH luminescence was collected by the ring of plastic fibers. A 450-nm interference filter (Corion,P10-450-S-G985)was used to isolate the emitted radiation before detection by a photomultiplier tube (PMT, Oriel, Model 70680) in combination with a lock-in amplifier (Princeton Applied Research Co., Model 5209). The power supply of a PMT readout device (Oriel, Model 7070) supplied the operating voltage for the PMT. An IBM-XT computer in conjunction with an Analog Connection Jr. A/D board (StrawberryTree,Inc., Sunnyvale, CA) was used to collect intensity data as a function of time. L-Glutamic acid (monopotassium salt), L-alanine, @-NAD+, bovine serum albumin (BSA, 96-99% albumin),L-glutamic dehydrogenase (E.C.1.4.1.3,36 units mg-l protein from bovine liver), glutamiepyruvic transaminase (E.C.2.6.1.2,80 units mg-l protein from porcine heart), all amino acids used in the interference study, ascorbic acid, uric acid, ammonium chloride, and GABA were obtained from Sigma Chemical Co., St. Louis, MO. Glutaraldehyde (25% solution) was purchased from Aldrich Chemical Co., Milwaukee, WI. Poly(viny1 alcohol) (PVA, Elvanol, 99.0-99.8% hydrolysis) was a gift from Dr. Eric Fogt, Promeon, Division of Medtronic, Inc., Brooklyn Center, MN. Silica fiber with a core diameter of 1mm (numerical aperture of 0.4) was obtained from Quartz Products Corp., Plainfield, NJ. Plastic fibers with 484-pm core diameter (numerical aperature of 0.47) were purchased from Mitsubishi Rayon America Inc., New York, NY. Enzyme Immobilization. The biocatalytic layer was constructed by cross-linking both enzymes and BSA with glutaraldehyde in a thin layer of PVA at the tip of a sensing unit. The sensing unit was constructed by cutting the end of a plastic pipet tip so that it had a diameter of approximately 6 mm. Epoxy was used to fix a thin layer of glass over the end of the pipet tip, and the resulting glass surface served as the support for the enzyme layer. An enzyme solution was prepared by combining GDH (6 mg, 144 units)and GPT (2 mg; 160units)with 20 pL of a Tris-HC1 buffer (pH 7.4) and 20 pL of a 10% BSA solution. After mixing, 10 pL of a 1%PVA solution was added. A 10-pL portion of the enzyme/PVA solution was transferred to the sensing unit and spread over the glass surface. Cross-linkingwas initiated by mixing 1pL of a 2% glutaraldehyde solution. The membrane was allowed to form at room temperature for 90 min. The sensing unit was placed at the end of the bifurcated fiber-optic probe such that the distance between the probe tip and the enzyme layer was 1 mm. The sensing unit was stored at 4 "C in pH 7.4 Tris-HC1 buffer when not in use. Sensor Response Measurements. Sensor responses were measured by immersing the sensor tip in 5 mL of a stirred 0.1
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NAD+ G L UTAMAT E alanine Flgure 2. Sensor schematic showing the biocataiyzed reactions for the dualsnzyme glutamate FOBS.
M Tris-HC1 buffer solution which contained NAD+ and alanine. Initially, the background intensity was recorded as the baseline signal. A known amount of glutamate was then added to the buffer solution, and the fluorescence intensity was monitored as a function of time until a steady-statesignal was attained. The reported sensor response corresponds to the difference between the steady-state and baseline signals. Afterward, the sensing tip was placed in a fresh aliquot of buffer to reestablish the baseline. Response and recovery times were measured as the time necessary for the sensor to achieve 95% of the total response. Unless stated otherwise, all measurements were carried out at 25 "C in a thermostated cell controlled by a common water bath (Fisher, Model 80).
RESULTS AND DISCUSSION The operating principle of the NADH-based glutamate biosensor is illustrated in Figure 2. During operation, glutamate, NAD+,and alanine diffuse from the bulk solution into the enzyme layer. In this layer, GDH catalyzes the formation of NADH from NAD+ during the oxidation of glutamate and GPT catalyzes a subsequent transamination reaction that re-forms glutamate and drives the GDH-catalyzed reaction toward the formation of NADH. This dual-enzyme reaction scheme enhances the amount of NADH generated a t the sensor tip, thereby increasing the magnitude of the sensor response. A steady-state concentration of NADH is obtained when the rate of NADH production is equal to the rate of NADH diffusion away from the sensing region. NADH luminescence generates a corresponding steady-state fluorescence intensity which is measured through the bifurcated fiber-opticprobe. The magnitude of the NADH fluorescence is related to the concentration of glutamate in the bulk solution as long as the rate of NADH production is first order with respect to glutamate. Several key parameters have been investigated in order to optimize the response properties of this dual-enzyme glutamate biosensor. These parameters include concentration of NAD+ and alanine as cosubstrates, pH, and temperature. The magnitude of the sensor response for 5 pM glutamate is plotted as functions of NAD+ and alanine concentrations in Figures 3 and 4, respectively. These experiments were carried out in pH 7.4 Tris-HC1 buffer. For the effect of NAD+ concentration, the alanine concentration was constant a t 10 mM and the concentration of NAD+ was varied from 0 to 6 mM. Similar experiments were performed for the effect of alanine, except the concentration of alanine was varied from 0 to 12 mM while the NAD+ level was held constant at 4 mM. In both cases, increasing the concentration of the cosubstrate resulted in an asymptotic approach to a maximum response. The sensor response is essentiallyindependent of cosubstrate concentrations when the NAD+ concentration is above 3 mM and the alanine concentration is above 8 mM. The concentrations of NAD+ and alanine used in all subsequent experiments were 4 and 10 mM, respectively.
ANALYTICAL CHEMISTRY, VOL. 64, NO. 9, MAY 1, 1992
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Figure 5. Effect of pH on the relative sensor response. 120
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Effect of temperature on the sensor response (0)and NADH fluorescence (W).
Concentration of Alanine [mM) Flgure 4. Effect of alanine concentration on the sensor response.
The effect of pH has been determined by comparing the relative sensor response to 5 pM glutamate over the pH range from 6.5 to 8.5. Figure 5 shows the pH profile obtained from this experiment. "he maximum sensor response was obtained over the pH range from 7.4 to 7.8. Only 75% of the maximum response is obtained at pH 8.5, and only 37% of the maximum is present at pH 6.5. The optimum response over the physiological pH range is a major advantage for this glutamate biosensor compared to the glutamate biosensor based on glutamate decarboxylase which cannot be used when the pH is greater than 5.5.6 All subsequent measurements were performed at pH 7.4 by using a Tris-HC1 buffer.
The effect of temperature on the response to 5 pM glutamate is shown in Figure 6. The relative sensor response decreased as the temperature was increased over the range from 20 to 40 OC. This experiment was repeated three times with a fresh sensor each time, and the plotted values represent the average of these measurements. This type of temperature effect is unusual for a biosensor. Typically, biosensors possess a maximum response within this temperature range. Normally, the enzyme activity and/or the diffusion properties are enhanced as the temperature increases, thereby giving greater signals. This enhancement continues until the temperature begins to thermally denature the enzyme which adversely affects the sensor response. In this case, however, a steady decrease in sensor response is observed which does not cor-
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ANALYTICAL CHEMISTRY, VOL. 64, NO. 9, MAY 1, 1992 1
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Concentration of Glutamate ( p M ) Figure 7. &tarnate calibration cwves using the duaknzyme (0)and singleenzyme )(. modes of operation. respond to a degradation of the enzyme. In fact, the decrease in response is caused by the change in luminescence properties of NADH as a function of temperature. Also plotted in Figure 6 is the relative fluorescence signal measured from standard NADH solutions at different temperatures. The relative fluorescence intensity decreases with temperature owing to a decrease in quantum efficiency of NADH.ls As shown in Figure 6, the decrease in biosensor response generally follows the decrease in NADH fluorescence. The deviation between 30 and 35 "C is caused by the enhancement of the biocatalytic activity which partially compensated for the decrease in NADH luminescence. The difference in the biosensor response when going from 35 to 40 "C is larger than that of the NADH fluorescence intensity which means that simple degradation of the luminescence properties of NADH are insufficient to explain the decrease in biosensor response. Degradation of enzyme and loss of biocatalytic activity are likely involved in this temperature range. Temperature also had a significant effect on sensor response time. Shorter response times were displayed at higher temperatures. Regponse times to 5 pM glutamate were 10,5, and 4 min at 20,25, and 30 OC, respectively. Although the sensor response was less at 25 OC, this temperature was selected for subsequent work because it represented a reasonable compromise between sensitivity and response time. The overall strong temperature dependency indicates that rigorous temperature control is needed to assure accuracy and precision. The glutamate response curve obtained under optimal conditions is presented in Figure 7. Measured steady-state fluorescence intensities are plotted versus the corresponding glutamate concentrations over the 0-10 pM range. From this reaponse w e , the sensitivity of this biosensor is 0.127 nA/pM and the detection limit (SIN = 3) is 0.13 pM. Response times were concentration dependent with shorter response times observed at higher concentrations. Response times ranged from 4 to 8 min for concentrations ranging from 0 to 10 pM. The response time was 5 min for 5 pM glutamate. Long time periods were required to recover to the baseline condition between measurementa. Recovery times were approximately
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Time (daya) Flgurr 8. Response to 5 pM glutamate as a function of sensor age. 25 min in most cases. The recovery time can be reduced by using a thinner enzyme layer, but such a configuration adversely affecta the sensitivity and detection limit of the sensor. The thickness of the enzyme layer was approximately 25 pm in the sensors tested here. The utility of the dual-enzyme configuration has been evaluated by comparing the response to that for the single enzyme system. The response from a sensor based solely on GDH activity is plotted in Figure 7 for comparison. The single-enzyme biosensor response was obtained by deleting alanine from the bulk solution, thereby prohibiting the GPT cycling reaction. Clearly, the sensor response is greatly enhanced by the addition of GPT. The sensitivity and detection limit of the single-enzymesystem are 0.011 nA/pM and 0.80 pM,respectively. These results indicate an 11.5-fold improvement in sensitivity and a 6.2-fold improvement in detection limit for the dual-enzyme arrangement. Responses were slightly faster for the single-enzyme system; for example, the response time was 4 min for 5 pM glutamate. Sensor stability has been measured by comparing sensor responses for 5 pM glutamate over a period of several days. Measurements were perfoded three times each day, and the biocatalytic component of the sensor was stored at 4 OC when not in use. The mean sensor response is plotted as a function of sensor age in Figure 8. The relative standard deviation for each m e a s ~ e m e nwas t less than 2%. This figure shows that the sensor response decreases slightly over the first 12 days but that the sensor retains 93% of the original response on the twelfth day. After day 12, the sensor response drops dramatically and only 12% of the original activity remains after 15 days. The sharp drop in response after 12 days corresponds to a decrease in enzyme activity below the critical value required for diffusion-controlledkinetics at the sensor tip. Changes in the sensor response were not caused by changes in the integrity of the enzymelpolymer layer. Such layers were firmly attached to the glass surface with no apparent change in structure for periods as long as 30 days. Selectivity has been examined by recording the sensor response to compounds known to be present in neurochemically relevant solutions. In this experiment, potential interferences
ANALYTICAL CHEMISTRY, VOL. 64, NO. 9, MAY 1, 1992
Table I. Relative Response from Potential Interferences" compd L-glutamate L-alanine L-arginine L-asparagine L-aspartate L-cysteine L-cystine L-glutamine L-glycine hydroxy-L-pinoline L- histidine L-isoleucine L-leucine
% response
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GABA urate taurine
Response from 100 pM of the tested compound relative to the response from 5 ctM glutamate.
were added to the buffer solution individually and the corresponding sensor response was recorded. The final ooncentration of the potential interference was 100 pM. The magnitude of the response has been compared to that obtained for 5 pM glutamate. The relative responses are summarized in Table I. No responses were observed for the majority of compounds tested, including ascorbic acid, uric acid, GABA, and taurine. Glutamine and lysine were the only compounds tested to give a measurable response. In the m e of glutamine, the responae is likely caused by the presence of a small amount of glutamate in the standard glutamine solution due to the natural hydrolysis of glutamine. For lysine, the measured response for 100 pM lysine was only 1%of that obtained for 5 pM glutamate. The corresponding selectivity fador is 2000 for glutamate over lysine. Endogenous alanine and NAD+ would not interfere with glutamate measurements, provided that both cosubstrates are present in saturating amounts in the working buffer. In addition, potential interference from ammonia has been evaluated. Our major concern was that high levels of ammonia would alter the extent of the glutamate dehydrogenation reaction, thereby cawing a negative interference and a systematic error in glutamate measurements. Evaluation of such an interference is particularly important before neurochemical applications because endogenous ammonia levels are known to be significant in such samples. In fact, total ammonia nitrogen levels as high as 200 pM have been measured in various neurochemically relevant samples.19 The effect of ammonia on glutamate measurements was determined in two ways. First, the sensor response was monitored while standard additions of an ammonium chloride solution were added to a 5 pM solution of glutamate. No effect was observed when the total ammonia nitrogen level was 56 pM. With 100 and 200 p M total ammonia nitrogen present, the sensor response for 5 pM glutamate was 98.9 and 97.8%, respectively, of that in the absence of ammonia. The second way in which the effect of ammonia was assessed involved preparing a series of glutamate calibration curves over the 1-10 pM concentration range with ammonia added to the working buffer. Calibration curves were made in the presence of 50, 100, and 200 pM total ammonia nitrogen. Resulta from this experiment were the same as those found before. In the presence of 200 pM total ammonia nitrogen, over 97.4% of the original response was observed at all concentrations of glutamate tested. While the effect of ammonia is minimal, maximum accuracy requires that an appropriate amount of ammonia be added to the working buffer during calibration.
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CONCLUSIONS The dual-enzyme biosensor for glutamate reported here is clearly superior to the single-enzyme case and offers high sensitivity, low detection limits,and excellent selectivity. The principal limitations of this sensor are long response and recovery times. The measured effect of temperature illustrate a general effect for all fiber-optic biosensors based on the fluorometric detection of NADH. The temperature dependency of NADH fluorescence results in a strong negative temperature effect for theae devices and demands temperature control during measurements. The requirement to add significant amounts of NAD+ and alanine to the analyte solution will restrict the use of this glutamate biosensor. From a neurochemistry standpoint, this sensor cannot be used for direct in vivo measurements where the levels of NAD+ and alanine cannot be controlled. This sensor can be used, however, to analyze discrete samples collected from microdialysis or tissue slice perfusion type experiments. In addition, this sensor can potentially be used to monitor in situ glutamate levels in extracellular fluids during tissue slice perfusion experiments when NAD+ and alanine are added to the perfusion buffer. Of course any cellular perturbations caused by adding these reagenta must f i t be identified with appropriate blank experiments. Overall, the excellent selectivity, sensitivity, and limit of detection of this biosensor suggest that it can be used as an alternative to chromatographic techniques for the determination of glutamate in neurochemically relevant solutions. Our next step will be to establish the utility of this sensor to measure glutamate released from photoreceptor cells during potassium-evoked depolarization experiments.
ACKNOWLEDGMENT This work was supported by a grant from the National Science Foundation (BNS-8716768).
REFERENCES (1) Vlllarta, R. L.; Cunningham, D. D.; Gullbault, 0. 0. Talenta 1991, 38, 49-55. (2) Kusakabe, H.; Mldorlkawa, Y.; Fujishima, T. A M c . Bld. Chem. 1984, 48, 181-184. (3) Yao, T.; Kobayashl, N.; Wasa. T. Anal. Chlm. Acta 1990, 231, 121-124. (4) Chn, C. Y.; SU, Y. C. Anal. C h h . Acta 1991, 243, 9-15. (5) Shank, R. P.: Campbel, 0. L. I n Mindbook of "&~IY; A., Ed.; Plenum Press: New York, 1983; Vol. 3, Chapter 14. (8) Dhr, 0. V.; ECIssa, L. H.: Arnold, M. A.; MUlec. R. F. J . Nevoscl. % I1988, 23, 83-89. (7) Dremal, E. A. A.; Schmld, R. D.; Wolfbels, 0. S. Anal. CMm. Acta 1991, 248, 351-359. (8) Wdlenberger, U.: Schellec, F.; Pawbwa, M.: MMer, H. 0.:Rislnger, L.: GOttOn, L. In @FMonogaphs: Schmid, R. D., Scheller, F., Eds.; VCH PubllsherS: New York, 1989; VOI. 13, pp 33-38. (9) Hale. P. D.; Lee, H. Sa; Okamoto, Y.; Skothelm, T. A. Anal. Lett. 1991. 24, 345-358. IO) Yao, T.: Yamamoto, H.: Wasa, T. Anal. C h h . Acta 1990, 236. 437-440. 11) Zhmennan, J. E.: Wlghtman, R. M. Anal. Chem. 1991, 63, 24-28. 12) Amold, M. A. I n Chemkal Sensors and MWnslrumenteMon; Murray, R. W., Dessy, R. E., Heineman, W. R., Janata, J., Seb, W. R., Eds.: ACS Symposium Serbs No. 403: American Chemical Society: Washlf@On, Dc, 198% p~ 303-317. 13) Amold, M. A.: Wangsa, J. I n F l k OpUc Chemical Sensors and Biosensff: Wolfbels, 0. S., Ed.; CRC Press: Bow Raton, FL, 1991: Vol. 2, Chapter 18. 14) Wangsa, J.; Arnold, M. A. Anal. Chem. 1988. 60, 1080-1082. 15) Ayoub, 0. S.: Copenhagen, D. R. J . Neuroscl. Methods 1991, 37, 7-14. (18) N M k , D. G. J . NeIXOChm. 1989, 52, 331-341. (17) Subramanlan, S. Mphys. Chem. 1978, 7 , 375-378. (18) Weng, J. L.; HO, M. H. Anal. Lett. 1990, 23, 2155-2173. (19) knjamh. A. M. In Mindbook of Newochemlsby; Lajtha. A,, Ed.; Plenum Press: New York. 1983; Vol. 1, Chapter 4.
Lam,
RECEIVED for review November 12,1991. Accepted January 31, 1992.
