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Anal. Chem. 1987. 59. 2256-2259
simultaneously. Detailed applications of this procedure to the provision of mechanistic information for irreversible electrooxidation reactions will be described elsewhere. It is commonly accepted that the electrooxidation of formic acid on platinum can occur via “dual pathways” involving both weakly and strongly adsorbed intermediates (16). The identity of the latter species has been the subject of much debate; although the presence of adsorbed CO is indicated from surface infrared spectroscopy ( 3 ) ,electrochemical (16) and coupled mass spectrometric (“DEMS”)measurements suggest that the strongly adsorbed intermediate is COH (17, 18). While the present results certainly do not eliminate the latter possibility, they do nonetheless indicate that the initially adsorbed CO is sufficiently reactive to have potential as a possible catalytically active intermediate under voltammetric conditions (15).
CONCLUDING REMARKS Overall, the SPAIR spectra provide unusually direct information on the chemical nature, as well as extent, of the electrochemical reaction induced by a single potential step (or sweep) and how the reactivity of the adsorbed CO intermediate compares with that for the overall conversion of reactants to products. Although the spectra obtained under potential-modulation conditions can provide some information along these lines, such conventional PMIR are clearly of limited value for examining irreversible electrode processes. In addition, the rapid potential modulation that is necessarily employed in PMIRS when using a grating spectrometer (i.e., EMIRS) can have severe deleterious effects on the surface condition and, hence, on the chemical and physical state of the adsorbed species. Admittedly, the likely applicability of SPAIRS is limited, at least using present commercial FTIR spectrometers, to the examination of interfacial species with especially strong infrared absorptions. Nevertheless, a real virtue is that the spectra can be obtained under electrochemically well-defined potential-step conditions on time scales down to a few seconds. This offers the additional prospect of employing time-dependent SPAIRS to extract electrochemical rate data as well as mechanistic information for adsorbed species for com-
parison with kinetic data obtained by conventional electrochemical means. The virtue of such spectral measurements is that they could provide a direct connection between the vibrational properties and electrochemical reactivity of adsorbed species. A variety of experiments in our laboratory along these lines, focusing particular attention on the electrooxidation kinetics of CO and small organic fuels (IS),are under way or are planned for the near future.
ACKNOWLEDGMENT Some experimental assistance was provided by Feng Bao. Registry No. CO, 630-08-0;HC02H, 64-18-6;Pt, 7440-06-4; Au, 7440-57-5;COP, 124-38-9.
LITERATURE CITED (1) Bewick, A.; Pons. S. In Advances in InfraredandRaman Spectroscopy; Clark, R. J. H., Hester, R. E., Eds.; Wiley Heyden: New York, 1985; Vol. 12, Chapter 1. (2) Foiey, J. K.; Korzenlewski, C.; Daschbach, J. L.; Pons, S. In Electroanalytical Chemistry-A Serles of Advances; Bard, A. J., Ed.; Marcel Dekker: New York, 1986; Voi. 14, p 309. (3) Foley, J. K.; Pons, S.;Smith, J. J. Langmulr 1985, 7 , 697. (4) Pons, S.;Datta, M.; McAleer, J. F.; Hinrnan, A . S. J . Elecfroanal. Chem. 1984, 760, 369. (5) Beden, 8.; Bewick, A . ; Larny, C. J . Electroanal. Chem. 1983, 748, 747. (6) Beden, B.; Bewick, A.; Larny, C. J . Necfroanal. Chem. 1983, 150, 505. (7) Kunirnatsu, K. J . Electroanal. Chem. 1982, 740, 205. ( 8 ) Kunirnatsu, K. J . Elecfroanal. Chem. 1983, 745, 219. (9) Kunimatsu, K. J . Electroanal. Chem. 1988, 273,149. 10) Corrigan, D. S.;Weaver, M. J. J . Phys. Chem. 1986, 90,5300. 11) Corrigan, D. S.;Milner. D. F.; Weaver, M. J. Rev. Sci. Instrum. 1985, 56, 1965. 12) Leung, L-W. H.;Weaver, M. J. J . Am. Chem. Soc., in press. 13) Sheppard, N.; and Nguyen, T. T. I n Advances in Infrared and Raman Sllecfroscow; Clark, R. J. H., Hester, R. E., Eds., Heyden: London, 1978; Voi. 5; p 67. 14) Beden, E.; Bewick, A.; Kunirnatsu, K.; Lamy, C. J . Electroanal. Chem. 1982, 142, 345. 15) Corrigan. D. S.;Weaver, M. J., submitted for publication in J . Nectroanal . Chem 16) Capon, A.; Parsons, R. J . Necfroanal. Chem. 1973, 4 5 , 205. 17) Wolter, 0.; Willsau, J.; Heitbaum, J. J . Nectrochem. Soc. 1985, 732, 1635. (18) Wiiisau, J.; Wolter, 0.; Heitbaurn, J., J . Necfroanal. Chem. 1985, 785,163.
.
RECEIVED for review March 6, 1987. Accepted May 28, 1987. This work is supported by the National Science Foundation.
Determination of Sulfite Using a Sulfite Oxidase Enzyme Electrode Vicki J. Smith Phillips Petroleum Company, Biotechnology Division, 21 1 CPL, Phillips Research Center, Bartlesville. Oklahoma 74004 A sulfite-senslng electrode was constructed by physically trapplng sulfite oxidase (EC 1.8.3.1) enzyme at the tip of a dissolved-oxygen electrode. The decrease In oxygen that occurs as sulfite is enzyrnatlcally oxidized to sulfate was measured amperometrlcaiiy. A linear response to three commonly used sulfltlng agents, sodium sulfite, sodlum blsulfite, and sodlum metablsulffte, was obtained. Precision of the measurements expressed as relative standard devlation was 1.0% at 100 ppm sodlum sulflte. Comparative determinations of the sulfite content of dried aprlcots, potato flakes, ros6 wfne, wine vinegar, and lemon julce uslng a colorimetric procedure and the sulflte electrode showed a 0.98 llnear correlation coefflcient. The electrode was stored refrigerated for at least 84 days without loss of response.
Interest in analytical methods capable of determining sulfiting agents has heightened due to recent Food and Drug Administration (FDA) rulings revoking the generally recognized as safe status of sulfiting agents for use on fruits and vegetables sold raw. FDA has also established 10 ppm as the threshold for declaration of sulfites in the labeling of foods, nonalcoholic beverages, and wine products ( I ) . This is the result of adverse and potentially life-threatening reactions in sulfite-sensitive individuals after eating foods treated with these agents (2-4). The mechanism of action and fate of sulfiting agents upon addition to food have been described (5-7) but are not completely understood. Among the analytical methods used for sulfite analysis are
0003-2700/87/0359-2256$01.50/0 42 1987 American Chemical Society
ANALYTICAL CHEMISTRY, VOL. 59, NO. 18, SEPTEMBER 15, 1987
titration by the Monier-Williams method ( I ) , gas chromatography (8), liquid chromatography (9), mercuric sulfide/ mercurous chloride electrodes (IO, II), ion chromatography (I2-14), enzymatic UV (15, 16) flow injection analysis (In, color test strip ( I @ , and spectrophotometry (19). Sulfite analysis is complicated by the instability of the ion in the presence of oxygen (12), loss as sulfur dioxide (gas) which exists in equilibrium with sulfite and bisulfite anion ( 5 ) ,and variability in the amount of “bound” sulfite released upon extraction. The FDA reference method is the Monier-Williams procedure in which samples are refluxed in boiling HCl for 1.75 h. The released sulfur dioxide is purged and trapped under nitrogen for subsequent titration. The method is time-consuming and not practical for numerous routine determinations. Many analytical applications of enzyme electrodes that amperometrically measure the decrease in dissolved oxygen accompanying oxidase-catalyzed substrate conversions have been described. Analytes that can be measured by using this type of bioanalytical device are uric acid (201, alkaline phosphatase @I), L-lactate (22),lactate dehydrogenase (22), acetic acid (23), lysine (24), and many others (25). This technique can potentially be applied to any oxidase system to measure substrate or inhibitor. This paper describes the preparation and use of a highly stable sulfite enzyme electrode built around a commercially available multipurpose biosensor. The sulfite electrode was found to detect as little as 10 ppm sodium sulfite in complex sample matrices and did not require sample clarification or deproteinization. Sulfite assays using the enzyme sensor were compared to results obtained by a spectrophotometric (pararosaniline) method for five sulfited foods. EXPERIMENTAL S E C T I O N Reagents. Enzyme Gel. Sulfite oxidase (EC 1.8.3.1 from chicken liver, 500 EU/mL) was obtained from Sigma Chemical Co. The ammonium sulfate suspension was used as received. Blank gel matrix was obtained from Provesta Corp. Colorimetric Sulfite Assay. Pararosaniline hydrochloride (P-389) was obtained from Fischer Scientific. Mercuric chloride, sodium chloride, and formaldehyde (37% solution) were from Mallinckrodt. Sulfite Standards. Anhydrous sodium sulfite, sodium bisulfite, and sodium metabisulfite were reagent grade chemicals obtained from Mallinckrodt and used without further purification. Buffer Salts. Trizma base from Sigma and monobasic and dibasic potassium phosphate from Mallinckrodt were reagent grade. Lemon juice, mashed potato flakes, dried apricots,wine vinegar and rose wine were obtained at a local market. Assembly of the Enzyme Electrode System. A Provesta Multipurpose Bioanalyzer Model 98 was used (Bartlesville,OK). The unit contains a dissolved oxygen probe, blank gel matrix, and membranes for preparation of enzyme electrodes. The Bioanalyzer processes the enzyme electrode signal which is output to a strip chart recorder. The procedure for preparation of the sulfite oxidase electrode was as follows: 7 pL of enzyme suspension (3.5 EU) was placed on a plastic weighing dish and blank gel matrix was added gradually until a gel formed. The gel was applied to the center of a wet dialysis membrane (provided with the Bioanalyzer). The conical tip of the dissolved oxygen electrode was removed and the sulfite oxidase gel and membrane were centered on the probe. The tip was replaced, sandwiching the sulfite oxidase gel between the Teflon membrane of the oxygen electrode and the dialysis membrane. Procedures. Standard solutions of sodium sulfite, sodium metabisulfite, and sodium bisulfite were prepared at 1000 ppm in 0.1 M Tris buffer, pH 8.5. These solutions were kept on ice and diluted to 10-100 ppm in assay buffer prior to analysis. Measurements with the sulfite oxidase probe were made by setting the Bioanalyzer gain at 6.8 and establishing a steady-statebase-lime reading with the probe tip immersed in a stirred solution of assay buffer, 0.2 M potassium phosphate, pH 7.5. Sample was then
2257
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Flgure 1. Typical response curves of the sulfite electrode at pH 7.5
to (A) 75 pg, (B) 50 pg, and (C) 25 pg of sodium sulfite. added to the buffer solution and the associated decrease in dissolved oxygen was measured. Response was measured on the strip chart recorder trace as the difference (in millimeters), between the original base line and the response curve. Values for standards and samples were measured at a set time, typically 3-5 min from the point of sample addition. The sum of buffer and sample volume was held constant at 4.0 mL. Sample volume was varied with concentration such that 0.1-1.0 pmol of analyte was added for each determination. The effect of temperature on electrode response was determined by using a water-jacketed beaker and a Forma Model 2067 bath and circulator. Stability of the sulfite probe was examined by measuring the response to 50 Kg of sodium sulfite at time intervals after storage: (1)refrigerated in 3.2 M ammonium sulfate, (2) refrigerated in 0.05 M phosphate buffer, pH 7.5, or (3) room temperature in 0.05 M phosphate buffer, pH 7.5. Sulfite in food samples was determined by the sulfite oxidase electrode and by spectrophotometric means using AOAC Method 20.044 and 20.045 (1). Sulfite values are expressed as parts per million of sodium sulfite. The spectrophotometric method measures the regeneration of color of a bleached pararosaniline solution in the presence of sulfite and formaldehyde. A Perkin-Elmer Model 570 spectrophotometer was used to measure absorbance at 550 nm. Lemon juice and rose wine were diluted 1:2 with distilled water prior to analysis. Potato flakes (5.0 g) were shaken vigorously with 50 mL of 0.2 M phosphate buffer, pH 7.5. A filtrate (no. 1 Whatman) was collected for analysis. Dried apricots (10.0 g) were blended with 290 mL of distilled water in a Waring blender for 2 min and filtrate (no. 1 Whatman) was collected. Wine vinegar was not pretreated. All samples were adjusted to pH 7.5-8.5 with NaOH and held at room temperature 10 min prior to analysis. A 1.0-mL sample was mixed with sodium tetrachloromercurate solution for subsequent spectrophotometric analysis. At the same time, 0.5-1.0 mL of sample was added to the sulfite electrode system. RESULTS AND DISCUSSION Electrode Response. Figure 1 shows sulfite electrode response to increasing amounts of sodium sulfite. The signal began to change 20-30 s after the addition of sulfite. Figure 2 shows calibration curves of enzyme electrode response to three commonly used sulfiting agents. Millimolar concentration of the sulfite salt in the bulk solution is shown. A lower detection limit of 10 ppm sulfite salt in food samples was achieved by using a 1.0-mL sample volume and a Bioanalyzer gain of 6.8. Relative standard deviation determined by eight successive analyses of a 100 or 10 ppm sodium sulfite standard were 1.0% and 4.670, respectively. A relative standard deviation of 2.7% was obtained for a lime juice sample which was processed and analyzed eight times (a = 79.16; 2.15 standard deviation).
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Flgure 2. Calibration curves for three sulfiting agents: (A) sodium metabisulfite, (B) sodium bisulfite, and (C) sodium sulfite.
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Figure 4. Relationship between sulfite oxidase gel mass and electrode response.
t
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Figure 3. Relationship between the sulfite oxidase content of the enzyme gel and electrode response. Variables i n Electrode Preparation. Enzyme gel mass was held constant while the sulfite oxidase activity per electrode was varied. Figure 3 shows that larger relative responses were obtained at lower concentrations of enzyme. This may be due to rapid oxidation of substrate as it diffuses into the outer enzyme gel layer a t high enzyme concentrations. Substrate would not reach the inner gel layer which is in immediate contact with the dissolved oxygen sensor, resulting in reduced response. Sulfite oxidase was held constant a t 1.8 EU per electrode while gel mass was varied. Each electrode was prepared by use of 5 p L of sulfite oxidase plus 0-30 WL of 3.2 M (NH4)&304. Blank gel matrix was added as required to form a gel. Figure 4 shows the %fold reduction in response which occurred as gel mass was increased. Excess gel may slow mass transfer of substrate and products resulting in reduced response. Effect of B u f f e r Concentration. Buffer (potassium phosphate, p H 7.5) concentrations ranging from 0.025 to 0.2 M had no effect on sulfite electrode response. This agrees with results of Cohen and Fridovich (26) showing little effect of phosphate concentration on oxygen consumption by sulfite oxidase, whereas cytochrome c reduction by the enzyme is reduced a t high phosphate levels. Effect of pH. MacLeod et al. (27) report a pH optimum for soluble sulfite oxidase of 8.5 with cytochrome c as electron acceptor. With oxygen as the electron acceptor, Cohen (26)
PH
Figure 5. Effect of pH on sulfiie electrode response; 0.1 M potassium phosphate buffer.
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Flgure 6. Effect of temperature on sulfite electrode response.
reports a pH optimum of 8.6. The effect of pH on electrode response was studied from p H 5.5 to 8.5. As shown in Figure 5 , the optimum pH was approximately 7.0. Within the pH range of 6.5-8.5 response remained at least 75% of maximum. A buffer pH of 7.5 was selected for sulfite electrode assays.
ANALYTICAL CHEMISTRY, VOL. 59, NO. 18, SEPTEMBER 15, 1987
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Flgure 7. Long-term stability of the sulfie electrode when stored: (A) at 4 “C In 3.2 M (NH4)2S04r (B) at 4 ‘C In phosphate buffer, and (C)
at room temperature in phosphate buffer. A decrease in response with multiple determinations was observed at pH less than 7.0. This is in agreement with reports (26) that the activity of sulfite oxidase decreases rapidly below pH 7.2 and is irreversibly inactivated at pH 5.0 or lower. Temperature Effect. Figure 6 shows the increase in electrode response obtained as the probe and buffer were equilibrated at various temperatures prior to addition of the sodium sulfite sample. For convenience, analyses were routinely performed a t room temperature (25 “C). Stability. The operational stability of the sulfite electrode under appropriate storage conditions was quite good, as shown in Figure 7. Electrodes stored at room temperature lost half their initial response in less than 10 days, whereas electrodes stored refrigerated between uses retained full initial response for 84 days. Although these results show that it is feasible to store the sulfite electrodes, fresh probes were routinely prepared each day. Interferences. Ascorbic acid interferes with this assay. Compounds such as ascorbic acid spontaneously consume oxygen giving a positive response in the absence of sulfite. The response of the sulfite electrode to 100 ppm ascorbic acid was approximately equal to the response to 100 ppm sulfite. Ascorbic acid also interferes with the enzymatic UV assay of sulfite (15,161. In the UV assay, sulfite oxidase converts sulfite to sulfate with the production of hydrogen peroxide. The latter product is subsequently reacted with NADH in the presence of peroxidase and the disappearance of NADH is followed spectrophotometricallyat 340 nm. It is reported that concentrations of ascorbate in excess of 100 mg/L give low sulfite values, presumably due to inhibition of one of the enzymes used. Ascorbic acid can be removed by pretreatment of samples with ascorbate oxidase (16). Assay of Sulfite in Food Samples. Figure 8 compares sulfite assays of five food samples by enzyme electrode and colorimetric methods. The linear correlation coefficient is 0.98. Each point represents a simultaneous determination on the same extract by the two methods. A set of assays of a particular food represents extracts prepared and assayed on different days. Slight variation of the pH used to prepare the extract is expected to result in variation of the assayed value due to the effect of pH on the release of “bound” sulfite from the food (7). Sulfite content decreases with time of storage. For example, an aqueous extract of dried apricota showed over 6000 ppm sodium sulfite when assayed immediately. This extract was stored refrigerated and reassayed several weeks later, at which time the sulfite value had fallen to about 4000
600
400
Enzyme Electrode.
ppm
Flgure 8. Correlation of the results of the sulfite electrode with those of a colorimetric procedure (AOAC 20.044) for the analysis of (W) wine vinegar, (X) lemon juice, (0) rose wine, (0)potato flakes, and (A)dried apricots (XlO-’).
ppm. These data demonstrate the potential utility of the enzyme electrode for determination of the sulfite in food samples. Procedures would need to be optimized for particular sample types. Advantages of the method include short analysis time, minimum equipment expense, and the ability to readily determine additional analytes using alternate enzyme gels.
ACKNOWLEDGMENT The author thanks R. A. Green for his technical assistance and T. R: Hopkins for encouragement and advice.
LITERATURE CITED (1) Fed. Regist. 1988, 57(131) 25012-25021. (2) Baker, G. J.; Coliett. P.; Allen, D. H. Med. J . Aust. 1981, 2 , 614. (3) Simon, R. A.; Green, L.; Stevenson, D. D. J . Allergy Clin. Immunol. 1982, 69, 118. (4) Stevenson, D. D.; Simon, R. A. J . Allergy Clin. Immunol. 1981, 88, 26. (5) Barnett, D. Food Technol. Aust. 1985, 37(1 l), 503-505. (6) Report by Institute of Food Technologists Food Technol. (Chicago) 1986, (June), 47-52. (7) Modderman, John P. J . Assoc. Off. Anal. Chem. 1986, 69, 1-3. (8) Hamano. T.; Mitsuhashi, Y,; Matsuki, Y; et ai, 2. Le6ensm.-Unters. -Forsch. 1979, 768, 195-199.
.
(9) Imaizumi, N.; Hayakawa, K.; Okubo, N.; Miyazakl, M. Chem. fharm Bull. 1981, 29(12),3755-2757. (10) Tseng, Paul; Gutknecht, W. F. Anal. Chem. 1978, 4 8 , 1996-1998. (11) Marshall, G. 6.; Mldgley, D. Analyst (London) 1983, 708, 701-711. (12) Lindgren, M.; Cedergren, A.; Llndberg, J. Anal. Chim. Acta 1982, 74 I, 279-286. (13) Sullivan, D. M.; Smith, R. L. Food Technol. (Chicago) 1985, (July), 45. (14) Anderson, C.;Warner, C. R.; Daniels, D. H.; Padgett, K. L. J . Assoc. Off. Anal. Chem. 1988, 69,14-19. (15) Beutler, H. 0.FoodChem. 1984, 75, 157-164. (16)Technical Update, sulfite: Test for the Enzymatic Determination of Sulfurous Acid in Foods; Boehrlnger Mannhelm Biochemicals, 1985. (17) Sullivan, J. J.; Holiingworth, T. A.; Wekell, M. M.; Newton, R. T.; LaRose, J. E. J . Assoc. Off. Anal. Chem. 1988, 69,542-546. (18) Markley, 8.; Meioan, C. E.; Lambert, J. L. Anal. Lett. 1988, 79(1-2),
37-46. (19) Official Methods of Analysis of the Association of Official Analytical Chemists, 14th ed.; Williams. S.,Ed.; AOAC: Washington, DC, 1984; p 379. (20) Nanjo, M.; Gullbault, G. G. Anal. Chem. 1974, 46, 1769-1772. (21) Kumar, A.; Christian, G. D. Anal. Chem. 1978, 4 8 , 1283-1286. (22) Mitzutanl, F.; Sasaki, K.; Shimura, Y. Anal. Chem. 1983, 55, 35-38, (23) Hlkuma, M.; Kubo, T.; Yasuda, 1.;Karube, I . ; Suzukl, S. Anal. Chim. Acta 1970, 709, 33-38. (24) Romette, J. L.; Yang, J. S.; Kusakabe, H.; Thomas, D. Siotechnolcgy and Bioengineering; Wlley: 1983;Vol. 25,pp 2557-2566. (25) Provesta Multipurpose Bioana!~zerhhnual; Provesta: Bartlesvllle, OK, 1986. (28) Cohen, H. J.; Frldovlch, I . J . Biol. Chem. 1971, 246,359-366. (27) MacLeod, R. M.; Farkas, W.; Fridovich, I.; Handler, P. J . Biol. Chem. 1961, 236, 1841-1846.
RECEIVED for review March 13,1987. Accepted May 18,1987.
