Amperometric Measurement of Enzyme Reactions with an Oxygen Electrode using Air Oxidation of Reduced Nicotinamide Adenine Dinucleotide Frank S. Cheng and Gary
D. Christian'
Department of Chemistry, University of Washington, Seattle, Washington 98 195
Dehydrogenase reactions are monitored with a membrane oxygen electrode, using aerobic oxidation of the reduced form of nicotinamide adenine dinucleotide (NADH) in the presence of horseradish peroxidase and co-factors. The optimum rate of oxidation of NADH Is obtained in slightly alkaline solution, favorable for analytical coupling of the reaction to dehydrogenase reactions. A linear relationship between the NADH concentration and the maximum rate of oxygen depletion is observed. The feasibility of anatytical determinations is demonstrated by measuring NADH produced in the ethanol-NAD-alcohol dehydrogenase reaction.
T h e electrochemical measurement of the reduced form of nicotinamide adenine dinucleotide (NADH) has been performed by a variety of means. NADH can be electrochemically oxidized at solid electrodes (2-4) for amperometric monitoring. Or it can be coupled to different redox agents for indirect amperometric measurement, such as Bindschedler's Green ( 5 ) , 2,6-dichlorophenol-indophenol(6),and ferricyanide (7). Many bare electrode measurements, however, are subject to slow response, drifting, or fouling in biological fluids. If monitoring of NADH could be made using a membrane covered amperometric electrode, such as the Clark oxygen electrode (8, 9) measurements would be simplified. The membrane protects the electrode from complex biological matrices. This electrode has been demonstrated to be useful for the measurement of small concentrations of oxygen-consuming substances in biological materials (1&13). Its response is rapid and stable. A number of methods has been used since the 1930's for the coupling of NADH to oxygen, largely for manometric assays. Most involve a redox dye as a n electron transfer agent, and these have been reviewed by Thomas (14). Among the most promising systems is the use of a peroxidase enzyme for coupling NADH to oxygen. The remarkable property of purified horseradish peroxidase (HPO) to catalyze the aerobic oxidation of the reduced form of nicotinamide adenine dinucleotide (NADH) and nicotinamide adenine dinucleotide phosphate (NADPH) under certain conditions was first reported by Akazawa and Conn (15). They observed that the Mn'+-dependent aerobic oxidation of NADH and NADPH was greatly accelerated by the addition of certain phenols. Williams-Ashman et al. (16, 17) have reported that certain phenolic estrogens can stimulate the reaction of NADH in the Mn2+-peroxidase-02 system. Similar results have been confirmed by Klebanoff (181,who demonstrated that thyroxine and its related compounds could activate the oxidation of NADH and NADPH, and the effects of estrodiol and thyroxine are additive. In addition to HPO, some other peroxidases, such as uterus peroxidase (1S-211, myeloperoxidase (22), and lactoperoxidase (23), have been shown to be capable of catalyzing the aerobic oxidation of NADH. I n this paper, we report a study of the oxygen-consuming oxidation of NADH in the presence of horseradish peroxidase
for the amperometric measurement of the NADH according to the reaction 2NADH + 0 ,
+
2H'
peroxidase
BNAD'
A
2H,O
The maximum rate of oxygen consumption is measured with a membrane electrode and related to the NADH concentration. It has been demonstrated that this measurement can be used for the rapid analytical monitoring of dehydrogenase or dehydrogenase coupled reactions. EXPERIMENTAL Apparatus. A glucose analyzer (Reckman Instruments, Inc.) with a Clark-type oxygen electrode was used to measure the amperometric current and electronically take its derivative. Both the Air Adjust and Glucose Sensitivity potentiometers were adjusted to maximum gain and the instrument was operated in the V-mode to obtain maximum sensitivity. Measurements were made at the thermostated temperature of 33.0 f 0.1 "C. Both the direct-current output and the derivative output were recorded on a two-pen strip-chart recorder. Reagents. Tris-succinate buffer, phr 8.0. Prepare a solution of 0.1 M tris(hydroxymethy1)aminomethane and 0.033 M succinate. Check and adjust pH by adding small quantities of either tris or succinate. Imidazole buffer, p H 8.0. Prepare a O.:l M solution of imidazole and adjust the pH with 1 M HC1. Horseradish peroxidase (donor: H 2 0 2oxidoreductase, E.C. 1.11.1.7) was obtained from Worthington Biochemical Corp. Alcohol dehydrogenase (Alcohol: NA.D oxidoreductase, E.C. 1.1.1.1) from yeast was purchased from Calbiochem. Procedure Place 0.75 mL of buffered stock solution containing the desired amount of horseradish peroxidase (about 15 IU) and 0.13 mM manganous ion in the cell of the analyzer. Allow the oxygen in the solution to come to equilibrium with atmospheric oxygen. (This is necessary because the analyzer has a temperature higher than room temperature. However, with 0.75 mL reagent, it takes only a few seconds t o equilibrate the oxygen tension). Inject a suitable volume (10 to 50 pL) of aqueous sample containing NADH into the cell to trigger the reaction, and record the rate of oxygen consumption vs. time. The derivative peak, which represents the maximum rate of current change (i.e., the maximum rate of reaction) is obtained in about 10 s after the sample is injected. RESULTS AND DISCUSSION Demonstration of Signals. Figure 1 shows some typical signals obtained for three different concentrations of aqueous NADH stock solution. The signals were recorded in both the direct oxygen uptake and the derivative rate of oxygen consumption modes. The difference between the baseline and the tip of the curve in the direct signals corresponds to the amount of NADH oxidized. Because the reaction is very rapid, the effect of oxygen back-diffusion from the atmosphere on the initial rate is small (24). Assuming complete oxidation, the rate of oxidation of NADH can be approximated by measuring the change in the direct signal. However, with high concentrations of NADH, a t the end of reaction, the difference ANALYTICAL CHEMISTRY, VOL. 49, NO. 12, OCTOBER
1977
1785
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Figure 1. Time course of NADH oxidation. Ten ~ Lof L (A) 1.0 mM NADH,
mL of 0.1 M tris and 0.033 M succinate buffer containing 15 IU horseradish peroxidase, 0.13 mM Mn2+,and 10 mM 2,4dichlorophenol at pH 8.0. The numbers of the peaks indicate the digital readings of the maxima taken from the instrument. Each small arrow indicates the time when the sample was injected into the cell
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Figure 2. Effect of manganous ion concentration on the maximum rate of oxygen depletion. Conditions similar to those for Figure 1 (2.0 mM NADH)
of oxygen tension between air and solution in the cell is large, and oxygen back-diffusion can cause some inaccuracy in the net direct measurements. Therefore, in our experiments, all quantitative measurements were made using the derivative signals, which correspond to the maximum rate of oxygen depletion and are directly related to the amount of NADH added. Effect of Manganous Concentration. Although there are many contradictory results reported in the literature as to the role of metal ions in peroxidase-catalyzed oxidations, it is generally accepted that manganous ion is important to activate the oxidations (25). For the aerobic oxidation of NADH by peroxidases, differing optimal concentrations of Mn2+for maximum activation have been suggested (15,18-20, 26-28). The p H participation of co-factors, and sources of peroxidases all contribute t o the differences. We have examined the effect of Mn2+concentration in the maximum rate of reaction for the NADH-02-peroxidase system under our conditions. Figure 2 shows the plot of the signals obtained as a function of Mn2+concentration. The optimum rate of oxygen depletion lies around 0.13 mM Mn2+. Effect of Enzyme Activity. Figure 3 shows the effect of enzyme concentration on the maximum rate of NADH oxidation. T h e rate of oxygen depletion for 15 nmol of NADH increases significantly up to about 10 IU enzyme, a t which point the reaction becomes nearly zero order with respect to 1786
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(5) 2.0 mM NADH, and (C) 3.0 mM NADH stock solution added to 0.75
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ANALYTICAL CHEMISTRY, VOL. 49, NO. 12, OCTOBER 1977
Figure 4. The pH profile. (A) Tris-succinate buffer. (5)Imidazole buffer. Conditions similar to those for Figure 1. The imidazole buffer was 0.1 M
enzyme and is independent of the enzyme activity. Effects of pH and Buffer. Although most peroxidases are active over a relatively broad range of pH, NADH is said to be unstable in acidic media, in which it undergoes acidcatalyzed decomposition (29). On the other hand, manganese is readily precipitated at high pH. Therefore, the selection of pH may be expected to be important in obtaining reproducible results. Also, the choice of the buffer system is important since the stimulating effect of manganous ion is dependent on the chelating ability of the buffer (30, 31). Optimal pH values for NADH oxidation have been reported to be 6.0 ( 1 5 ) ,6.5 in the presence of thyroxine ( l a ) ,.5.6 in the presence of 2,4-dichlorophenol (19), 7.7 in the presence of estrodiol-17 (20), 7.5 in tris-buffer and 5.0 in succinate buffer (25). Figure 4 shows the effects of pH on NADH oxidation in two buffers in our system: trissuccinate and imidazole. The results indicate that under our experimental conditions, the optimum pH is about 8.0 in the former buffer and only slightly less in the latter. It also shows that while trissuccinate buffer gives larger signals, the imidazole buffer has more tolerance to higher pH. I t should be pointed out that while oscillations in the oxidation reaction can occur in acid solution under proper
Table I. Effect of Co-factors in the Peroxidase-Catalyzed NADH OxidationQ Inert
Inhibitory
2,4-dichlorophenol ascorbic acid phenol resorcinol thyroxine 1 7 p-estrodiol estrone rn-cresol p-cresol o-cresol
hydroquinone guaiacol X
.
The compounds were tested at a concentration of 0.02 mM b y adding 1 0 G L of 1 . 5 mM solution of each compound t o the reaction cell of analyzer, which contained 0.13 mM Mn2+,0.1 M tris, 0.033 M succinate, and 1 5 IU peroxidase a t p H 8.0. solution conditions (23,26-28, 32-35), we do not observe the oscillations in alkaline medium. We have, though, observed oscillations in slightly acidic media when high concentrations of NADH are used, but the phenomena are not reproducible, even under closely controlled experimental conditions, in agreement with previous investigators (27, 28). The use of alkaline medium yields rapid, reproducible, and linear reaction without complicating oscillations and unreliability. E f f e c t o f O t h e r Co-factors. Certain phenolic compounds are known to have a stimulatory effect on peroxidase-catalyzed NADH oxidations, including 2,4-dichlorophenol (15) and estrogens with a phenolic hydroxyl group (16-18). We examined several compounds for possible stimulatory or inhibitory effects in our system, listed in Table I. The compounds are classified into two groups: inert and inhibitory. In contrast to other investigators, we found that most phenolic compounds tested in our procedure (at concentrations o f 2 x 10-5M) did not show a stimulatory effect when Mn2+was used a t its optimum concentration of 0.13 mM. I t must be noted, however, that the activating ability of phenolic cofactors is dependent on their concentration in the reaction medium, since Mn2+ and co-factor concentrations, p H and buffer are all interrelated in the peroxidase-catalyzed oxidation system ( 2 5 ) . We have found that a slight increase in the maximum reaction rate (about 10%) is observed when 2,4dichlorophenol is increased to 10 mM concentration. On the other hand, increased resorcinol concentration (0.5 mM) causes inhibition of the NADH oxidation. E f f e c t of H y d r o g e n P e r o x i d e . The inhibitory effect of catalase on NADH oxidation (catalyzed by H P O and uterine peroxidase) has caused several workers (15, 18-21) to study the possible role of H202in the reaction. H 2 0 2is reported to increase the reaction rate (15) and to induce HPO to form Compound 111, a peroxidase derivative believed to be responsible for observed reaction oscillations (26). We have examined the effect of H 2 0 2on the reaction rate and on oscillations under our procedure. Figure 5 demonstrates that addition of H 2 0 2t o the reaction medium caused a decrease in the maximum rate of oxygen depletion. Although this is in contrast to the result obtained by Akazawa and Conn (151,we did find a similar effect with catalase, with the addition of 50 IU catalase to the reaction system causing a 30% reduction in the signal. As shown in Figure 1, under our experimental conditions, there was no oscillation observed by continued recording of the signal for several minutes during NADH oxidation. Nor did we observe the oscillation signals when H 2 0 2was added to the reaction mixture. L i n e a r i t y . Previous investigators have reported that the rate of HPO-catalyzed aerobic oxidation of NADH is not
Figure 5. Effect of H202concentration on the oxidation of NADH. The reaction mixture contained the Same composition as Figure 1, with the addition of H202concentration indicated in the graph; 10 pL of 1.5 mM NADH solution was added to start the reaction
-
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Figure 6. Recorded signals for different NADH concentrations. The signals were recorded in two recorder sensitivities, 1 and 10 V full scale, respectively. The numbers on the peaks indicate the concentrations of NADH stock solution added, in millimoles per liter
linearly dependent on the concentration of NADH, using spectrophotometric measurement of NADH ( 1 5 , 18,26, 27). Yamakazi and his collaborators (27) have even considered this irregular dependence of the oxidation velocity as a cause of the periodic oscillation. We have found that aqueous NADH stock solutions react with atmospheric oxygen in the presence of HPO with a linear dependence of the maximum rate 011 the NADH concentration. Duplicately recorded signals for different NADH concentrations are shown in Figure 6. The peak heights give the maximum rate of consumption of oxygen and are directly proportional to the concentration of NADH stock solution shown on the peaks. The smallest peak represents 3.8 nmol of NADH added to 0.75 mL buffered reagent in the cell, for a final concentration of 5.1 pM. The maxima are reached within 10 s of initiating the reaction. R e a c t i o n M e c h a n i s m s . Akazawa and Conn (15) proposed a free radical theory for the aerobic oxidation of NADH, in which they suggested that a ternary complex composed of Mn2+,peroxidase, and H 2 0 2catalyzes the oxidation of NADH. Their reaction mechanism has generally been accepted by other workers (17, 18, 21) for reaction a t neutral pH. ANALYTICAL CHEMISTRY, VOL. 49, NO. 12, OCTOBER 1977
1787
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Figure 7. Rate of oxygen depletion vs. NADH generated in vitro. NADH was produced by incubating different amounts of alcohol dehydrogenase (ADH) with 3 m L buffered substrate, containing 10 m M NAD, 0 33 M ethanol. and 0.1 M imidazole at pH 8.8 and 33 OC. Numbers o n curves represent IU of ADH
Yokota and Yamazaki ( 2 6 ) , on the other hand, in their studies of the oscillation of NADH oxidation in acidic pH, have proposed an ingenious reaction mechanism in which a free radical of NADH reacts with peroxidase intermediates. They attributed the oscillatory behavior to the regulatory properties of Compound III. However, this reaction mechanism has been disputed by Degn et al. (34, 35). Despite the extensive investigations of HPO-catalyzed oxidation of NADH in acidic and neutral pH, little study has been done in alkaline pH. Our results indicate that the reaction mechanism perhaps follows a different scheme. First, although Mn“ ion is essential in our procedure for NADH oxidation, phenolic cofactors have little effect on the reaction. This is in contrast to the reaction in neutral p H which requires both Mn2+and a phenolic co-factor, as well as in contrast to the reaction in acidic pH, which does not need any activators. Second, no oscillation was observed, and H 2 0 2seems to have no effect in our system. Finally, the reaction observed by us is very rapid with the maximum rate obtained in about 10 s, and the relationship between NADH concentration and the maximum rate of oxygen depletion (or maximum reaction rate) is linear. These results are in contrast to those obtained by previous investigators for the reaction in acidic and neutral pH, in which irregular relationships have been reported. Although a detailed reaction mechanism to interpret the NADH oxidation is not available a t the present time, our observations indicated that the reaction follows MichaelisMenten type steady-state kinetics. Analytical Application. The linear relationship between the substrate concentration and the reaction rate suggests the possible analytical application of the reaction to NADHproducing reactions. We have demonstrated the feasibility of such an application by measuring the NADH produced in the ethanol-NAD-alcohol dehydrogenase (ADH) reaction. Buffered NAD-ethanol mixtures were incubated in the presence of ADH for different times, and the NADH produced was analyzed amperometrically by the HPO-catalyzed oxidation reaction. Figure 7 shows the results of the signals obtained. At each indicated time, 50-fiL aliquots were withdrawn from an incubation medium containing NAD, imidazole buffer, and different ADH activities as indicated. The withdrawn sample was then immediately added to the measurement cell of the analyzer containing 0.75 mL of a buffered HPO reagent with a composition similar to that given in Figure 1. A calibration graph for ADH, determined for 4 min of incubation, was linear for 0.02-0.4 IU of ADH. This 1788
ANALYTICAL CHEMISTRY, VOL. 49, NO. 12, OCTOBER 1977
clearly indicated that by allowing a short incubation time, this method can be applied to the measurement of dehydrogenase activities. With the sensitivity and rapidity of the NADH oxidation reaction, a direct single-step amperometric measurement of low concentrations of alcohol appeared possible. The feasibility of such measurements was in fact, demonstrated using a reagent mixture similar to the one for the incubation method described above. For each determination, 0.75 mL of buffered enzyme-coenzyme reagent, containing 3.0 mM NAD, 1.5 JU HPO, 20 IU ADH, 0.13 m M Mn2+and tris-succinate buffer with concentrations of 0.1 M and 0.033 M, respectively, at p H 8.0, was placed in the cell of the analyzer. The reaction was started by injecting 10 fiL of aqueous ethanol into the cell, and the signals were then recorded. The peaks, which represent the maximum rate of oxygen depletion of HPO-catalyzed NADH oxidation, and in turn the alcohol concentration, were obtained in about 20 s. The calibration curve obtained by plotting the peak height vs. aqueous alcohol concentration was linear from 0.02 to 0.2 g ethanol per 100 mL stack solution, which is within the physiological range of blood alcohol. The working range can be doubled by decreasing sample size from 10 to 5 pL. For an aqueous alcohol containing 0.15 g ethanol per 100 mL water, 14 successive measurements, using a 1O-kL sample size, gave a coefficient of variation of k3.34%. This experiment demonstrates that the NADH oxidation reaction is sufficiently rapid that the rate-limiting step occurs in the ethanol-NAD reaction. The described measurement system offers the potential for rapid and sensitive measurement of dehydrogenases and dehydrogenase-coupled reactions.
LITERATURE CITED (1) (2) (3) (4) (5) (6) (7) (8) (9) (10) (11) (12) (13) (14) (15) (16) (17) (18) (19) (20) (21) (22) (23) (24) (25) (26) (27) (28) (29) (30) 1311 . , (32) (33) (34) (35)
W. A. Blaedel and R. A. Jenkins, Anal. Chem., 47, 1337 (1975). G. G. Guilbault and T. Cserfalvi, Anal. Lett., 9, 277 (1976). L. C. Thomas and G. D. Christian, Anal. Chim. Acta, 78. 271 (1975). G. D. Christian, in “Ion and Enzyme Electrodes in Biology and Medicine”, M. Kessier, Ed., Urban and Schwarzenberg, Munchen-Berlin-Wien, Germany, 1976. M. D.Smith and C. L. Olson, Anal. Chem., 46, 1544 (1974). M. D. Smith and C. L. Olson, Anal. Chem., 47, 1074 (1975). L. C. Thomas and G. D. Christian, Anal. Chim. Acta, 82, 265 (1976). L. C. Clark and C. Lyons, Ann. N . Y . Acad. Sci., 102, 29 (1962). G.D. Christian, Adv. Biomed. Eng. Med. Phys.. 4, 95 (1971). A. Kumar and G. D. Christian, Clin. Chem. ( Winston-Salem, N.C.), 21, 325 (1975). A. Kumar and G. D. Christian, Anal. Chem., 48, 1283 (1976). F. S. Cheng and G. D. Christian, Analyst (London), 102, 124 (1977). A . Kumar and G. D. Christian, Clin. Chim. Acta, 24, 101 (1977). L. C. Thomas, Thesis, University of Washington, Seattle, Wash., 1975. T. Akazawa and E. E . Conn, J . Biol. Chem., 232, 403 (1958). H. G. Williams-Ashman, M. Cassman, and M. KlaviPs, f e d . Proc., 18, 352 (1959). H. G. Williams-Ashman, M. Cassman. and M. Klavins, Nature (London). 184, 427 (1959). S. J. Klebanoff, J . Biol. Cbem., 234, 2480 (1959). V . P. Hollander and M. L. Stephens, J . Biol. Chem., 234, 1901 (1959). S. Temple, V. P. Hollander, N. Hollander, and M. L. Stephens, J . Biol. Chem., 235, 1504 (1960). J. Beard and V. P. Hollander, Arch. Biochem. Eiophys., 96, 592 (1962). J. Schuttz and S. J. Berger, in “Biochemistry of the Phagocytic Process”. J. Schultz, Ed., John Wiley 8 Sons, New York, N.Y.. 1970, p 115. S. Nakamura, K. Yokota. and I. Yamazaki, Nature(London),222, 794 (1969). L. C. Thomas and G. D. Christian, Anal. Chim. Acta, 89, 83 (1977). J. M. Mudd and R . H. Burris. J . Biol. Chem., 234, 2774 (1959). K. Yokota and I. Yamazaki. Eiochim. Biophys. Acta., 105, 301 (1965). I. Yamazaki, K. Yokota, and R . Nakajima, Biochem. Siophys. Res. Commun., 21, 582 (1965). I. Yamazaki and K. Yokota, Biochim. Biophys. Acta, 132, 310 (1967). 0. H. Lowry, J. V . Passoneau, and M. K. Rock, J . Eiol. Chem.. 236, 2756 (1961). R . H. Kenten, Biochem. J . , 5 9 , 110 (1955). E. R . Wavaood. A. Oaks. and G. A. Maciachlan. Can. J . Botany. 34, 905 (19567. H. Degn, Nature(London), 217, 1047 (1968). H. Degn, in “Biological and Biochemical Oscillator”, B. Chance, E. K. Pve. A. K. Gosh. and E. Hess. Ed.. Academic Press. New York. N.Y., 1973, p 109 H Degn, Biochm Biophys Acta. 180, 271 (1969) H Degn and D Mayer, Biochim Biophys Acta, 180, 291 (1969)
RECEIVED for review April 8, 1977. Accepted June 27, 1977.
