523
Anal. Chem. 1986, 58,523-526 (13) Booman, G. L.; Holbrook, W. B. Anal. Chem. 1963, 35, 1793. (14) Brown, E. R.; Smith, D. E.; Booman, G. L. Anal. Chem. 1968, 4 0 , 141 1. (15) Brown, E. R.;Hung, H. L.; McCord, T. G.; Smith, D. E.; Booman, G. L. Anal. Chem. 1988, 4 0 , 1424. (16) Bewlck, A. Nectrochim. Acta 1968, 73,825. (17) Garreau, D.; Saveant, J. M. J . Nectroanal. Chem. 1972, 35, 309. (18) Brown, E. R.; McCord, T. G.; Smith, D. E.; DeFord, D. D. Anal. Chem. 1966, 38, 1119. (19) Schroeder, R. R.; Shain, I. Chem. Instrum. 1969, 7 , 233. (20) delevie, R.; Husovsky, A. A. J . Elecfroanal. Chem. 1969, 2 0 , 181. (21) Piiia, A. A.; Roe, R. 6.; Herrmann, C. C. J . Nectrochem. Soc. 1969, 116, 1105. (22) Piila, A. A. J . Electrochem. SOC. 1971, 718,702. (23) Weiis, E. E., Jr. Anal. Chem. 1971, 4 3 , 87. (24) Sarma, N. S.; Sankar, L.; Krishnan, A.; Rajagopaian, S. R. J . Electroanal. Chem. 1973, 4 7 , 503.
(25) Deroo, D.; Diard, J. P.; Guitton, J.; Gorrec, B. J . Nectroanal. Chem. 1976. 67. 269. (26) Gabrkili, C : Ksouri, M.; Wlart, R. Electrochim. Acta 1977, 22, 255. (27) Garreau, D.; Saveant, J. M. J . Nectroanal. Chem. 1978, 86, 63. (28) Yarnitzky, C.; Friedman, Y. Anal. Chem. 1975, 4 7 , 876. (29) Yarnitzky. C.; Klein, N. Anal. Chem. 1975, 4 7 , 880. (30) He, P.; Avery. J. P.; Fauikner, L. R. Anal. Chem. 1982, 12, 1313A. (31) He, P. Ph.D. Thesis, University of Illinois at Urbana-Champaign, 1984. (32) Sedra, A. S.;Smith, K. C. "Microelectronic Circuits"; CBS College Publishing: New York, 1982. (33) Shea, R. F. "Amplifier Handbook"; McGraw-Hill: New York, 1966.
RECEIVED for review June 25, 1985. Accepted October 10, 1985. We are grateful to the National Science Foundation for supporting this work through Grant CHE-81-06026.
Immobilized Enzyme Electrode for the Determination of Oxalate in Urine Mohammad A. Nabi Rahni and George G. Guilbault*
Department of Chemistry, University of New Orleans, Lakefront, New Orleans, Louisiana 70148 Net0 Graciliana de Olivera
Instituto de Quimica, Unicamp, Campinas, Brazil
An lmmobllized enzyme electrode for the assay of oxalate, particularly In urlne, was constructed. I t Is based on the lmmoblllzatlon of the enzyme oxalate oxidase on pig Intestine, mounted on the tip of an oxygen electrode. The oxygen partial pressure was amperometrlcally monitored, the current change being converted to a voltage through an adapter, whlch could then be monitored directly on a dlgttal voltmeter. The experimental parameters were all optimlzed, and callbration curves based on both the initial rate and the steady state were constructed. The enzyme electrode response to oxalate in urlne samples from 14 patlents was compared to that obtained by the establlshed spectrophotometric method. The proposed method exhibits high sensitivity and speclflclty to oxallc acid with almost no loss of relative activity due to interferences studied. There Is no sample pretreatment required and the method can be modlfled to be equally useful for the assay of oxallc acid in any blologlcal or nonblologlcal samples.
During recent years, there has been an increasing interest in the determination of oxalic acid in a wide variety of biological and nonbiological materials (1);oxalate has also been used in analytical (2) and manufacturing procedures (3). It has also been shown that increased urinary oxalate excretion may lead to the development and formation of renal calculi and urinary tract stones ( 4 , 5 ) . The determination of oxalate in urine is also clinically important for the diagnosis of various forms of hyperoxaluria, a genetic disorder of oxalate metabolism characterized by the early onset of calcium oxalate nephrolithiasis and nephrocalcinosis (6, 7). Current methods for determination of oxalic acid can be divided into three main groups: (1) solvent extraction and precipitation; (2) isotopic dilution; and (3) enzymatic analyses. Excellent reviews of these methods can be found elsewhere
(8, 9). Among these, enzymatic methods generally seem to be promising in terms of specificity, selectivity, and sensitivity where sample preparation (preconcentration, extraction, etc.) is minimal. Two enzymes have now been identified in purified form: oxalate decarboxylase [EC 4.1.1.21 from the wood rot fungus (Colybia velutipes ref lo), which has been shown to be highly specific in catalyzing the stoichiometric reaction: oxalate COz + formate; and oxalate oxidase [EC 1.2.3.41 prepared from plant tissue (11,12), which catalyzes the reaction
-
(C0OH)Z + 0
oxalate oxidase 2
2C02
+ HzOz
The immobilization of enzymes has found ever increasing popularity, since in most cases the enzyme stability is increased, the effects of enzyme inhibitors greatly reduced or eliminated (13, 14), and the immobilized enzyme useful for many assays. Enzyme electrodes have opened a new era in clinical analysis (13). The method presented herein utilizes the purified enzyme oxalate oxidase, immobilized on an oxygen electrode, for the determination of oxalate in urine. An amperometric method is more sensitive than a potentiometric based oxalate decarboxylase electrode method (15,16). It can be applied to small volumes of untreated urine and is therefore not subject to errors introduced by preliminary treatments. The method outlined here is a rapid, one-step procedure suitable for the analysis of any samples containing oxalate. EXPERIMENTAL SECTION Apparatus. A PHM 84 Research pH meter coupled to a REC 61 Servograph recorder (Radiometer America, Inc.) through a dc offset module and potentiometricamplifier (Model EV-200-1and 2, Schlumberger) was used for all measurements. To eliminate the necessity of using a polarograph to monitor the amperometric oxygen electrode, an adapter (Model No. CP-960, Universal Sensors, Inc., P.O. Box 736, New Orleans, LA) was used. It is a device that will simultaneously apply the desired potential to the amperometric enzyme electrode, take the resulting current
0003-2700/86/0358-0523$01.50/0 0 1986 American Chemical Society
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ANALYTICAL CHEMISTRY, VOL. 58, NO. 3, MARCH 1986
generated, and convert it to a voltage that can then be read on the voltmeter. A Radelkis combination O2electrode (type A) was employed for the construction of the enzyme electrode. Pig intestine type H was obtained from Universal Sensors. Reagents and Chemicals. All of the following chemicals were obtained from Sigma Chemical Co., St. Louis, MO: oxalic acid dihydrate, succinic acid, 8-hydroxyquinoline,EDTA sodium salt, a-chymotrypsin from bovine pancreas [EC 3.4.21.13, bovine albumin, glutaraldehyde (25% grade 11), oxalate oxidase [EC 1.2.3.4.1, oxalate kit (procedure No. 590), L-lysine, (dihydroxypheny1)acetic acid, L-cysteine, L-lactic acid, acetylsalicylic acid, and acetaminophen (Paracetamol). Any other chemicals mentioned were reagent-grade available in our laboratory. Double distilled deionized water was used for all solution preparations. Procedure. The enzyme electrode was prepared by a covalent cross-linking method described by Guilbault et al. (17, 18). At first, a pig intestine type H membrane was mounted on the tip of a Radelkis O2electrode by an "0''-ring. It was then activated by a few drops of chymotrypsin solution (0.5 mg/0.5 mL of H,O). Then, 20 yL of 5% bovine serum albumin solution containing 2.50 mg (3.0 units) of lyophilized,oxalate oxidase was put at the center of the pig intestine membrane. Then, 2.0 yL of 6.25% glutaraldehyde solution was added as a cross-linking reagent, and the resulting layer was stirred rapidly by a thin nylon rod for 2 min and let dry at room temperature for 3 h. The mounted pig intestine was then removed from the electrode jacket, inverted, and mounted again such that the immobilized enzyme was now sandwiched between the intestine and the polypropylene oxygen membrane. Besides the chemical cross-linking immobilization, the physical entrapment of the enzyme layer may help maintain enzyme activity and perhaps reduce some of the interference effects. The electrode was then equilibrated in freshly prepared 0.05 M succinate buffers, pH 3.50, containing 1 mM EDTA and 0.65 mM 8-hydroxyquinoline. This was the optimum buffer solution and was used throughout the experiments. In all measurements, the total volume was kept at 1.5 mL. All solutions reached a thermal equilibrium (25 "C) and a steady base line before the substrate of appropriate concentration (oxalic acid stock solution, 0.01 M) was introduced. The signal was recorded for both the initial rate and the steady state. The enzyme electrode jacket was kept at 5 "C in buffer for later use.
RESULTS AND DISCUSSION Electrode Response. The Universal Sensors adapter simultaneously applies a constant potential of -0.65 V to the oxygen electrode and converts the current output of the electrode to a potential that can then be monitored on a regular p H meter. Every 1-nA change in the current due to oxygen concentration change (the change in the oxygen partial pressure) in the bulk solution is equal to a 10 mV potential change monitored on the voltmeter. Therefore, a linear relationship exists between the current and voltage changes, and either one can be used to construct the calibration curve. For every assay, the enzyme electrode was placed into a stirred solution of 0.05 M succinate buffer, pH 3.50 (containing 1mM EDTA and 0.65 mM 8-hydroxyquinoline) to reach a steady base line. Substrate solution with appropriate volume (stock solution of substrate, 0.01M) was then added to the buffer to obtain the desired concentration of oxalic acid in a total volume of 1.5 mL. In the bulk solution, both the initial rate of disappearance of dissolved oxygen current and the total current change a t steady state were proportional to the substrate concentration. Steady-state currents are obtained due to an equilibrium between the dissolved oxygen consumption by the enzymatic reaction and the supply of oxygen from the bulk solution to the enzyme layer on the surface of the electrode. The buffer solution is first allowed to reach a steady base line, which is due to a rather constant oxygen concentration. This gives rise to a residual signal of about 100-120 nA (1000-1200 mV). The assay signal is on top of this background curlent. Effects of Experimental Parameters. A substrate bulk
1 .o
100
rLE D T Ad1 ,
2.0 1
(A)
mM
3.0
4.0
I
1
5.0
c 80
I
1
1
0.05
0.10
0.15
EUCCINAT~,
0.20
0.25
M
Flgure 1. Effectsof (A) EDTA and (B) succinate concentrations on the relative activity of the enzyme electrode. Oxalic acid concentration was 2.91 X low4M at optimum conditions.
solution of 2.91 X M was used to optimize the experimental parameters. I t was found that p H 3.5 was well within the optimum range in a p H profile. Also, 0.65 mM 8hydroxyquinoline gave the highest relative enzyme activity. Figure 1A illustrates the EDTA concentration profile, where a 1.0 m M concentration, in conjunction with 0.65 mM 8hydroxyquinoline, resulted in the highest relative activity. In the same manner, a t pH 3.5, different succinate concentrations were tried, and a 0.05 M concentration was found to give rise to the best relative activity (Figure 1B). I t has been reported that the purified enzyme does not need a cofactor and that succinate buffer is the only one suitable for oxalate assays (19). Thus, the effects of the EDTA and 8-hydroxyquinoline (as well as succinate to a lesser extent) at low concentrations are probably due to their binding of calcium and other metal ions present. This leads to more free oxalate, to be acted upon by the enzyme electrode. At higher concentrations these diverse substances affect the enzyme reaction through depression of some side reactions or exert a salt effect. Finally, temperature effect studies on the reaction rate were performed; there was less than a 6% variation in relative activity when going from 26 "C (room temperature) to 37 "C. Hence, 26 "C was quite sufficient for all studies. Construction of Calibration Curve(s). The total volume for every assay was kept a t 1.5 mL. Aliquots of stock solution were added to initiate the enzyme-catalyzed reaction; both the initial rate and steady state were recorded. A calibration curve obtained a t steady state for the assay of oxalic acid showed linearity of current change vs. oxalate concentration M. The response time for M to 2.2 X between 1.3 X the steady-state attainment is generally slow and quite subjective (7-12 min for low concentrations and 15-25 min for high concentration). Alternatively the initial rate of reaction can be monitored. Such a calibration curve obtained shows a linearity between 1.2 x M and 2.58 X lo4 M oxalic acid concentration (Figure 2B) for a freshly prepared enzyme electrode. The calibration was repeated when the enzyme electrode was 10 days old and the results are shown in Figure 2A. I t was found that not only the slope of the calibration improved by about 15% but also the sensitivity increased. The linear analytical region is now extended to 2.85 X M. Every assay and base line recovery takes only about 4 min when the initial rate is measured. Stability and Selectivity of the Electrode. The stability of the enzyme electrode was monitored by constructing a
ANALYTICAL CHEMISTRY, VOL. 58, NO. 3, MARCH 1986 ---703
1300
525
Table 11. Recovery Studies of Randomly Spiked Aqueous and Urine Samples of Oxalic Acid concn added, M 1.0 x 3.0 X 6.0 x 1.0x 1.3 x 2.2 x av
10-5 10-5
10-4 10-4
10-4
recovery," % found (aqueous) found (added to urine) 105.0 98.2 97.6 98.7 99.9 99.0 99.7
97.0 98.0 106.4 93.5 98.7 98.5 98.7
"Every assay is the average of three measurements. Table 111. Reproducibility Studies of Urine Samples (Obtained from Three Patients and Analyzed on Nine Separate Occasions) Flgure 2. Standard calibration curve of oxalic acid using Initial rate of current vs. concentration when (A) enzyme electrode is 10 days old
run no.
and (B) electrode is only 3 days old. Table I. Effect of Diverse Substances on the Activity of the Oxalate Oxidase Reaction compound"
enzyme activity,b%
ascorbic acid acetaminophen dihydroxyphenylaceticacid L-cystine L-lycine L-lactic acid D-glUcOSe
99 99 96 95 89 100 100
'Concentrations: 2.91 X lo4 M in 1 mM EDTA + 0.65 mM 8-Hydroxyquinoline in succinate buffer, pH 3.50. *The enzyme activity was set at 100% using 2.91 X M oxalic acid (no interference present). calibration curve every 3 days. Figure 3 illustrates the percent relative activity vs. the number of days, obtained when a 2.91 x lo4 M substrate solution has been assayed. The electrode lost part of its enzymic activity during the first several days and then increased by about 25% relative activity. Finally it stabilized to about 85-90% of original activity. The enzyme electrode relative activity stayed stable for almost 2 months and the electrode can be used for well over 200 assays during this time. Table I summarizes the effects of some potential interM oxalic ferences on the relative activity. Again, a 2.91 X M acid concentration was used, and the effects of 2.91 X added concentrations of each of these potential interferences on the assay were studied. L-Lysine seems to have the highest (- 11% on the relative activity), whereas ascorbic acid, which is the precursor of oxalate in the metabolic pathways, seemed to have no effect. This is probably due to the effective immobilization and selectivity of the modified pig intestine matrix used. Precision Studies. The reproducibility and recovery studies were conducted using randomly spiked aqueous and urine samples. The corresponding oxalate acid concentration obtained from a calibration curve similar to that in Figure 2A was compared to the amount originally added. Table I1 shows the percent recoveries of such assays of oxalic acid: 97.6-105.0% for aqueous and 93.5-106.4%, for urine samples, respectively. Urine samples from three patients were analyzed for oxalic acid on nine separate occasions. The mean values obtained for each patient were 27.5,18.53, and 21.5 mg/L, with coefficient of variations of 3.2,4.20, and 5.16, as shown in Table 111.
oxalic acid concn (mg/L) for patient no. 1 2 3
4
27.5 28.6 26.8 28.8
5
27.4
6 7
26.5 28.2 26.4 27.3 27.5 0.88 3.2
1 2
3
8 9 mean, mg/L std dev % coeff var
18.4 18.8 18.1 19.2 19.4 17.5 17.7
18.2 19.5 18.53 0.78 4.20
21.9 22.7 21.1
20.8 21.4
19.8 20.2 22.5 22.9
21.5 1.11 5.16
Table IV. Comparison of the Proposed Enzyme Electrode and Spectrophotometric Methods
specimen no.
oxalic acid concn, mg/L spectrophotomenzyme electrode etric method ( Y)" method ( X ) n
14
32.6 20.0 18.1 18.4 40.0 38.7 33.7 24.6 27.5 30.4 18.7 13.1 17.4 12.5
mean
24.7
1
2
3 4
5 6 7
8 9 10 11
12 13
34.1 22.8 17.5 24.7
37.5 38.3 36.6 26.7
29.5 34.8 26.8 16.7 16.4 14.4 26.9
"Regression of X on Y , X = -2.89 + 1.025Y; standard error of estimate of Y on X , SY,X= 2.904; correlation coefficient, r = 0.95; coefficient of determination, r2 = 0.903; coefficient of alienation, 1 - r2 = 0.097; t test between means of paired samples, t = -2.897.
Comparison of the Proposed Immobilized Enzyme Electrode and Spectrophotometric Methods. The spectrophotometric kit method (Sigma) is one of the most reliable currently available. This was used as a reference method to minimize any error associated with reagent preparation. Urine specimens from 14 adult donators (both men and women) were analyzed, using both the established spectrophotometric and the proposed enzyme electrode methods. The comparison results are outlined and presented in Table IV. A regression equation, X = -2.89 + 1.025Y, a standard error of estimate,
526
Anal. Chem. 1986, 58,526-532
50-99-7.
LITERATURE CITED
81OC
*
c >
b-
o 4
5c
w
z
b-
4 4
w
cc 1
4
1
1
1
12
2 0
28
--I
3 6
L 44 E
DAYS Figure 3. Long-term stability of the enzyme electrode.
Sy,x= 2.904, and a correlation coefficient of r = 0.95 were obtained. Registry No. EDTA, 60-00-4;oxalate oxidase, 9031-79-2;oxalic acid, 144-62-7; succinic acid, 110-15-6; ascorbic acid, 50-81-7; acetaminophen, 103-90-2;dihydroxyphenylaceticacid, 102-32-9; ccystine, 56-89-3;L-lycine,56-87-1;L-lactic acid, 79-33-4; D-glUCOSe,
(1) Hodgkinson, A. "Oxalic Acid in Biology and Medicine"; Academic Press: New York, 1977. (2) Vogel, A. I. "Textbook of Quantitative Inorganic Analysis"; Longmans: London, 1982, pp 243, 320, 577. (3) Pakel, G.; Florio, F. A. "Kirk-Othmer Encyclopedia of Chemical Technology", 2nd ed.; Wiley: New York, 1967; Vol. 14, pp 356-373. (4) Peacock, M.; Heyburn, P. J.; Robertson, W. G. Br. J . Uroi. 1978, 50, 449-454. (5) Galosy, R.; Clarke, L.; Ward, D. I . ; Pak, Cyc J . Urol. (Baltimore) 1879, 723, 320-323. (6) Wyngaarden, J. B.; Elder, T. D. I n "The Metabolic Basis of Inherited Disease"; Stanbury, J. B., Fredrickson, D. S., Eds: McGraw-Hill: New York, 1960; pp 449. (7) Archer, H. E.; Dormer, A. E.; Scowen, E. F.; Watts, R . W. E . Lancet 1957, ii:320. (8) Hodgkinson, A. Clin. Chem. (Winston-Salem, N . C . ) 1970, 76, 547-557. (9) Zerwekh, J. E.; Drake, E.; Gregory, G.; Griffith. 0.; Hofmann, A. F.; Menon, M.; Pak, C. Y. C. Clin. Chem. (Winston-Salem, N . C . ) 1983, 29, 1977-1980. (10) Shimazono, M.; Hayaishi, 0. J . Biol. Chem. 1857, 227, 151. (11) Srivastava, S.K.; Krishnan, P. S. Biochem. J . 1982, 8 5 , 33-389 (12) Chiriboga, J . Arch Blochem. Blophys. 1986, 116, 516-523. (13) Guilbault, G. G. "Handbook of Enzymatic Methods of Analysis"; Marcel Dekker: New York, 1977; pp 497. (14) Klibanov, A. M. Anal. Biochem. 1979, 93, 1. (15) Kobos, R. K.; Ramsey, T . A. Anal. Chlm. Acta 1980, 721, 111-118. (16) Fonong, T.; Rechnitz, G. A. Anal. Chim. Acta 1984, 758, 357-362. (17) Mascini, M.; Guilbault, G. G. Anal. Chem. 1877, 49, 795-798. (18) White, W. C.; Guilbault, G. G. Anal. Chem. 1878, 5 0 , 1481-1486. (19) Chiriboga, J. Biochem. Blophys Res. Commun. 1983, 7 7 , 277-282.
.
RECEIVED for review July 29, 1985. Accepted October 8, 1985.
Direct Observation of Trypto phan Biosynthesis in Escherichia coli by Carbon-I3 Nuclear h, agnetic Resonance Spectroscopy Dennis J. Ashworth,* Chi 5.Chen, and Desmond Mascarenhas Western Research Center, Stauffer Chemical Company, 1200 South 47th Street, Richmond, California 94804
Carbon-I 3 nuclear magnetic resonance (NMR) spectroscopy has been applled to the dlrect monltorlng of L-tryptophan biosynthesis In genetically modlfled E . GO//. Growth of the bacteria In the presence of ~-[3-'~C]serlne followed by NMR analysis of the culture supernatant generated a spectrum contalnlng resonances from nonmetabollred [3-"C]serlne as well as resonances from serlne-derived [3-"C]tryptophan and [2-13C]acetate. Growth In the presence of [2-%]glycIne resulted In a spectrum contalning SIXmajor resonances. A comparison of the chemlcal shlfts to those of L-tryptophan allowed assignment of two resonances to [2-%]tryptophan and [3-13C]tryptophan. The remalnlng four resonances, generated by one-bond 13C-'3C coupllng ( J = 33.8 Hr), were asslgned to [2,3-13C]tryptophanand verlfied by two-dlmenslonal homonuclear ( ''C) correlated spectroscopy. Growth of the bacteria In the presence of [6-'3C]glucose resulted In the labeling of the C-3, C-4', and C-7A' posltlons of tryptophan. To monitor tryptophan productlon In a fermentor, a device was constructed that allowed the contlnuous pumping of ferment dlrectly Into and out of a speclal NMR tube whlle growth of the culture was maintained.
Nuclear magnetic resonance (NMR) spectroscopy has emerged as one of the more powerful methods for studying biological processes. This is due primarily to the technique's
nondestructive nature and the ability of a single NMR spectrum to reveal the many potential metabolites of a given substrate. Its ability to probe the biochemical pathways of plants (1-3), animals (4-6), yeast (7-9), and bacteria (10-12) is now well documented. In E. coli the immediate precursors of tryptophan, indole-3-glycerol phosphate and L-serine, have been well established from a number of investigations (13-15). The required serine is generally considered to be derived from 3phosphoglycerate. Following dehydrogenation, transamination, and finally phosphate hydrolysis of 3-phosphoserine, L-serine is obtained (16,17). An alternative route to serine is from glycine (18). Indole-3-glycerol phosphate is generated by the combination of erythrose 4-phosphate and phosphoenolpyruvate ultimately produce chorismic acid. Conversion of chorismate to anthranilate followed by addition of 5-phosphoribosyl l-pyrophosphate and indole ring formation then leads to indole-3-gycerol phosphate (19,ZO). Glucose, being the primary bacterial source of reduced carbon, is the origin of both serine and indole biosynthesis.
L-Tryptophan
0003-2700/88/0358-0526$01.50/00 1986 American Chemical Society
