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(17) Monroe, D. Anal. Chem. 1984, 56, 920A-931A. (18) Kelly, T. A.; Christian, G. D. Talanta 1982, 29, 1109-1112. (19) Worsfokl, P. J.; Hughes, A.; Mowthorpe, D. J. Analyst 1985, 110, 1303- 1305. (20) Ismail, A. A. A.: West, P. M.; Goldie, D. J. Clin. Chem. 1978. 2 4 , 571-579. (21) Nolan, J. P.: DlBenedetto, G.; Tarsa, N. J. Clin. Chem. 1981, 2 7 , 730-747. (22) Bernard, A. M.; Lauwerys, R. R. Clin. Chem. 1983, 2 9 , 1007-1011. (23) De Alwis, W. U.; Wilson, G. S. Anal. Chem. 1985, 5 7 , 2754-2756. (24) De Alwis, W. U.; Wilson, G. S. Anal. Chem. 1987, 59, 2786-2789. (25) Lee, I. H.; Meyerhoff, M. E. Mikrochim. Acta 1988, III, 207-221. (26) Hara, T.; Toriyama, M.; Imaki, M. Bull. Chem. Soc. Jpn. 1985, 58, 1299-1303. (27) Mattiasson, B.; BerdBn, P.; Ling, T. G. I Anal. Biochem. 1989, 181, 379-302. (20) Bachas, L. G.; Meyerhoff, M. E. Anal. Chem. 1988, 58, 956-961. (29) Daunert, S.; Bachas, L. G.; Meyerhoff, M. E. Anal. Chim. Acta 1988, 208, 43-52. (30) Daunert, S.; Payne, B. R.; Bachas, L. G. Anal. Chem. 1989, 61, 2 160-2 164. (31) Rubenstein, K. F.; Schneider, A. S.;Ullman, E. F. Biochem. Biophys. Res. Commun. 1972, 37, 046-851. (32) Kjellstrom, T. L.; Bachas, L. G. Anal. Chem. 1989, 61, 1728-1732. (33) Zettner, A.; Duly, P. E. Clln. Chem. 1974, 2 0 , 5-14.
Sylvia D a u n e r t Leonidas G.Bachas* Department of Chemistry University of Kentucky Lexington, Kentucky 40506-0055 Genevieve S. Ashcom M a r k E. Meyerhoff Department of Chemistry The University of Michigan Ann Arbor, Michigan 48109-1055 RECEIVED for review July 20, 1989. Accepted November 2, 1989. This research was supported by grants from the National Institutes of Health (Grant No. R29-GM40510) (L.G.B.) and the National Science Foundation (CHE-8813952) (M. E.M.). G.S.A. acknowledges support by the Program in Scholarly Research for Urban/Minority High School Students of the University of Michigan.
TECHNICAL NOTES Polishable and Robust Biological Electrode Surfaces J o s e p h Wang* a n d K u r i a n Varughese Department of Chemistry, New Mexico State University, Las Cruces, New Mexico 88003 Tremendous research efforts are being devoted to the surface immobilization of appropriate biological entities (1, 2). Electrochemical probes have been exploited for biosensing more extensively than other devices (3). Depending on the specific biocomponent, a variety of surface immobilization procedures have been explored in the fabrication of electrochemical biosensors. Most commonly, the biological element is placed on top of the electrode surface, where it is being held physically (behind membranes) or chemically (via intermediate linkage). Alternatively, biocomponents can be incorporated directly into a carbon paste matrix to produce rapidly responding, inexpensive, and miniature sensing devices. The cumbersome renewal procedures and/or the poor mechanical and chemical stability of such biosurfaces often limit their practical bioanalytical utility. The motivation of the work described in this note was to develop biologically modified electrodes that would be mechanically robust, polishable, durable, and inexpensive. For example, to enhance the day-to-day practicality of electrochemical biosensors, it is highly desirable to renew their surfaces by simple polishing procedures, common with conventional (solid) electrodes. One promising concept is that of bulk modified electrodes (4-7). In this new class of chemically modified electrodes, the modifier is incorporated into the bulk of a robust carbon/polymer matrix. The bulk of the electrode thus serves as a “reservoir” of the modifier in a manner analogous to modified carbon paste electrodes (8-10). (The latter, however, are soft, nonpolishable, and not stable in most organic solvents.) The successful incorporation of various chemical modifiers (electrocatalysts, preconcentrating agents) into composite electrodes containing carbon black ( 4 , 51, graphite epoxy (61, or carbon fiber (7) have been reported recently. A similar avenue for the fabrication of reusable biologically modified electrodes and the challenges involved in the use of biological entities for this task are explored in the following sections.
EXPERIMENTAL SECTION Apparatus. The 10-mL electrochemical cell (Model VC-2, Bioanalytical Systems) was joined to the working electrode, reference electrode (Ag/AgCl, Mode RE1, Bioanalytical Systems), and platinum wire auxiliary electrode through holes in its Teflon cover. The three electrodes were connected to a Princeton Applied Research Model 174A polarographic analyzer, the output of which was displayed on a Houston Omniscribe stripchart recorder. The flow injection was described elsewhere (8). Electrode Preparation. Biologically modified graphite-epoxy electrodes were prepared by adding the desired quantity of the biocomponent (7.5-25% (w/w)) to the 1:l resin/accelerator mixture of a commercial epoxy-bonded graphite (Grade RX, Dylon, Cleveland, OH); a through mixing proceeded for 10 min. The enzyme/cofactor electrode were prepared in a similar manner, by adding the glucose oxidase/ 1,l’-dimethylferrocene mixture to the resin/accelerator mixtwe. A portion of the electrode material was packed into the end of a 5-mm4.d. glass tube. The electrode was then cured at room temperature for 15-20 h. The surface was polished with silicon carbide papers (150 and 320 grit) for a time period of 5 s each and then with 0.05-pm alumina slurry for 30 s. Residual polishing material was removed from the surface after each step by thoroughly rinsing with doubly distilled water. Such polishing procedure was repeated before each experiment. Between experiments, the bioelectrodes were stored at 4 “C. Reagents. All solutions were prepared with doubly distilled water. Supporting electrolytes were 0.05 M phosphate buffer (pH 7.4) and 0.1 M acetate buffer (pH 5.0). Dopamine, catechol, glucose, 8-nicotinamide adenine dinucleotide (NAD+) (Sigma), ascorbic acid, potassium ferrocyanide, copper nitrate (Baker), 1,l’-dimethylferrocene(Aldrich),ethanol (U.S. Industrial Chemicals), and methanol (Fisher) were used without further purifiStar”,Universal Food, cation. The dry active yeast granules (“M Milwaukee, WI) and the brown alga Eisenia bicyclis (Westbrae Natural Food, Berkeley, CA) were ground with a mortar and pestle. Tyrosinase (EC 1.10.3.1), horseradish peroxidase (EC 1.11.1.7),and glucose oxidase (EC 1.1.3.4)were obtained from Sigma.
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Table I. Summary of Experimental Conditions
% in
biocomponent tyrosinase's* horseradish peroxidasea dry yeasta Eisenia bicyclis' glucose oxidase'*'
graphite epoxy
working solution
operating potential, V
7.5'
0.05 M phosphate buffer (pH 7.4)
-0.2
20b 25
0.05 M phosphate buffer (pH 7.4), containing 10 mM K,Fe(CN)6
-0.2
20 20 20
0.05 M phosphate buffer (pH 7.4) containing 1 mM NAD' and 1 mM F,Fe(CN)6 +0.6 0.1 M acetate buffer (pH 5.0) scan from +0.6 to -0.7 +0.5 0.5 M phosphate buffer (pH 7.4)
'Batch experiments with 400 rpm stirring. bFlow injection analysis with 20-pL samples flowing at 1.0 mL/min. 'Also containing 26% 1,l-dimethylferrocene.
--
7
1
-
--Bl
A
---_------------.
I
1 1
I TIME
Figure 1. Current-time recording upon increasing the ethanol concentration in 5 X lo-' M steps: unmodified (A) and yeast-modified (B) graphke-epoxy electrodes. Batch experiment with a 400 rpm solution stirring and 4-0.6V operating potential. Other conditions are given in Table I. Procedure. Amperometric detection was performed by applying the desired potential and allowing the background current to decay to a steady-state value. Solution stirring (400 rpm) or flow (1.0 mL/min) were used in batch and flow injection experiments. The preconcentration/medium-exchange/voltammetric scheme for algae-containing electrodes was described elsewhere (9). Specific details of the experimental conditions are summarized in Table I.
RESULTS AND DISCUSSION The major challenge in fabricating reusable and robust bioelectrodes based on the bulk modification concept is the heating requirement of the curing/polymerization process. While this has no effect upon the incorporation of most chemical modifiers (4-7),the high temperature may have a deleterious effect on the activity of biological entities. Unlike carbon black or carbon-fiber composite electrodes ( 4 , 5, 71, the graphite-epoxy fabrication strategy (6) offers the advantage that the curing process can take place at room temperature. Although this requires longer (12-20 h) curing times (vs high-temperature curing), it is essential when biocomponents are concerned. The electrodes are very rigid upon formation. They render their bioactivity upon polishing for immediate reuse and appear to be smooth to the naked eye. The incorporation of four different biological entities (of relevance to bioanalysis) into the graphite epoxy material is used in the following sections to illustrate the concept of polishable and robust biosurfaces. Figure 1demonstrates the response of conventional (A) and yeast-containing (B) graphite-epoxy electrodes, to successive standard additions of ethanol, each addition effecting a 5 x M increase in concentration. In the absence of an enzymatic reaction, the former is not responding to additions of ethanol. In contrast, the yeast electrode (with its alcohol dehydrogenase activity) responds rapidly to the change in ethanol concentration, producing the steady-state current response within 28 s. The fast response is attributed to the
Figure 2. Differential pulse voltammograms for 1 X M copper obtained at the Eisenia /graphite-epoxy electrode using the preconcentration/medium-exchange/voltammetricapproach. Five minutes accumulation in a stirred copper solution, followed by rinsing and placing the electrode in a blank (electrolyte) solution to record the voltammogram. Scan rate was 10 mV/s. The electrode was polished prior to each cycle. The dotted line represents the response in the absence of copper. Other conditions are given in Table I.
intimate contact of the biocatalytic and graphite sites. The yeast actually becomes an integral part of the rigid sensing element, in a manner analogous to nonpolishable carbon paste bioelectrodes (8). Algae represent another class of microorganisms that can be incorporated in a stable and active manner into robust graphite-epoxy surfaces. The bioaccumulation of metal ions by alga-modified electrodes has been exploited for designing new sensing devices (9). Figure 2 demonstrates the ability to polish an alga-modified electrode to reproducibly renew its surface. I t shows a series of six successive copper measurements, each recorded with a freshly polished Eisenia-containing surface. The effective collection of the metal by the surface-bound alga is indicated from the well-defined voltammograms obtained after transfer of the electrode to the blank solution. The shape of the voltammogram and overall signal-to-background characteristics are similar to those obtained at nonpolishable alga surfaces (9). Both faradaic and background currents are unaffected by polishing. This series yielded a mean peak current of 38.4 PA, with a range of 38.s39.2 PA and a relative standard deviation of 1.6%. Such precision indicates that the electrode preparation results in homogeneous dispersion of the biological modifier within the graphite epoxy matrix and that the polishing step does not adversely affect the bioaccumulation process.
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ANALYTICAL CHEMISTRY, VOL. 62, NO. 3, FEBRUARY 1, 1990
C
t
W
K 0:
l
J
5
0
Pmin n *
3
4min
3
U
TIME
Figure 3. Amperometric measurements of catechol (A, B) and glucose (C) at enzyme/graph#e-epoxy elechodes: (A) current-time recording upon increasing the catechol concentration in 1 X M steps; (B) flow-injection detection of catechol solutions of increasing concentration from 2.5 X to 7.5 X M (a-c); flow rate, 1.0mL/min; (C) current-time recording upon increasing the glucose concentration in 5 X lo4 M steps; sdutions, 50:50 (v/v) methad-phosphate buffer (pH 7.4) (A) and 0.05 M phosphate buffer (pH 7.4) (B, C). Other conditions are given in Table I.
le5
Isolated enzymes were subsequently tested as biological modifiers, to examine whether the room-temperature curing process has a detrimental effect on their bioactivity. Figure 3 A,B shows the batch and flow amperometric response for catechol at the tyrosinase-modified electrode. For both operations, the rigid enzyme electrode responds very rapidly to the change in the substrate concentration. The reductive detection of the enzymatically produced quinone species, with its favorable signal-to-noise characteristics, allows convenient measurements of micromolar concentrations. Besides high sensitivity and speed, the graphite-epoxy enzyme electrode may be used in organic solutions. For example, a 5050 (v/v) methanol-phosphate-buffer solution was employed in part A of Figure 3. Tyrosinase is known for its ability to catalyze reactions in nonaqueous media (11). Unlike mixedenzyme/carbon-paste electrodes, where the oil binder may dissolve upon exposure to organic solutions, the graphiteepoxy biosurface is stable and robust. The high sensitivity and speed of the flow injection data, coupled with the ability to operate in organic media hold prospect for detection in liquid chromatography. Figure 3C illustrates the ability to coimmobilize an enzyme and its cofactor in the graphite-epoxy matrix. The glucose oxidase/dimethylferrocene/graphite-epoxy electrode responds rapidly to successive additions of 5 X loe4M glucose. Horseradish peroxidase represents another enzyme that was incorporated in an active and stable form within the graphiteepoxy matrix. A series of ten concentration increments of 1 X 10"' M hydrogen peroxide yielded a fast amperometric response (18 s for attainment of steady state) and a linear
concentration dependence (cofiditions as in Table I; not shown). The ability to renew by polishing fresh enzyme electrodes was tested for ten calibration plots for dopamine (50-500 rM) obtained at ten individual tyrosinase surfaces (conditions as in Figure 3A). For all surfaces, linearity prevailed up to 3.5 X lo4 M. The mean sensitivity value (slope of linear portion) found was 9.2 nA/pM, with relative standard deviation of 10%. Similar surface-to-surfacereproducibility experiments at the horseradish peroxidase or glucose oxidase/dimethylferrocene containing electrodes, involving additions of hydrogen peroxide or glucose, yielded relative standard deviations of 12% and 14%, respectively. hdeed, single tyrosinase, glucose oxidase, and horseradish peroxidase electrodes were able to be used for periods of several weeks, performing several hundred measurements with no noticeable loss of stability. Hence, the needs to re-create (by immobilization)the enzyme layer each time are eliminated. While the above data indicate no significant loss of bioactivity during the electrode preparation and routine operation, future studies will explore the utility of thermostable enzymes (isolated from thermophilic bacteria or artificial ones) in connection with the bulk modification approach. In conclusion, the experiments described above indicate that carbon composites can be used to prepare robust bioelectrodes for bioanalytical applications. Such electrodes mimic the behavior of their corresponding carbon paste counterparts (&IO), but possess the advantage that they may be polished (to renew the surface reproducibly) and can operate in organic media. Speed of response, simple modification scheme, versatility, miniaturization, and controlled bulk composition represent other advantages of graphite-epoxy bioelectrodes. Interferences (from coexisting electroactive or surface-active substances) are similar to those encountered at modified carbon-paste probes. Even though the concept is presented in terms of four model biocomponents, it could be extended to the incorporation of numerous biological modifiers. The simultaneous incorporation of a second enzyme (for sequence or competitive operations) or of chemical moieties (e.g. electrocatalyst) should further enhance the power of these composite biosurfaces. Registry No. Peroxidase, 9003-99-0;glucose oxidase, 9001-37-0; tyrosinase, 9002-10-2; hydrogen peroxide, 7722-84-1; ethanol, 64-17-5;copper, 7440-50-8;graphite, 7782-42-5;catechol, 120-80-9; glucose, 50-99-7.
LITERATURE CITED (1) Rechnitt, G. A. Chem. Eng. News 1988, Sept 5, 24. (2) Arnold, A. M.; Meyerhoff, M. E. CRCCrft. Rev. Anal. Chem. lB88, 20 (3), 149. (3) Kobos, R. K. TrAC, Trends Anal. Chem. 1987, 6 , 1987. (4) Shaw, B.; Creasy, K. E. Anal. Chem. 1988, 60, 1241. (5) Park, J.; Shaw, B. Anal. Chem. 1989, 67, 848. (6) Wang, J.; Golden, T.; Varughese, K.; El-Rayes, I. Anal. Chem. 1989, 61, 508. (7) Creasy, K. E.; Shaw, B. R. Anal. Chem. 1989, 61. 1460. (8) Wang, J.; Lin. M. S. Anal. Chem. lB88, 60, 1545. (9) Gardea-Torresdey, J.; Darnall, D.; Wang, J. Anal. Chem. 1988. 60, 72. (10) Santos, L. M.; Baldwin, R. P. Anal. Chem. 1988, 58, 848. (1 1) Hall, G. F.; Best, D.; Turner, A. R. F . Anal. Chim. Acta 1988, 213, 113.
RECEIVED for review July 17, 1989. Accepted November 1, 1989. This work was supported by the National Institutes of Health (Grant No. GM 30913-06).
