508
Anal. Chem. 1980, 52, 508-511
Figure 6 shows a A T vs. urea concentration response curve for the sample loop system. T h e sensitivity, a(An/aC,. is 5 X "C M-'; this is of the order of magnitude expected for a diffusionally limited device ( 4 ) and is about two orders of magnitude less sensitive than devices that carry the reaction to completion (7-14). In conclusion, this study demonstrates the ability to make relatively high resolution AT measurements with an open-flow channel configuration and under conditions compatible with immobilized enzyme use. This configuration has allowed us to address several significant effects on A T measurements: (a) the common mode rejection ratio to temperature (CMRRT) of a thermistor pair, which partially decouples variations in temperature of the incoming stream from A T measurements; (b) the CMRRT coupling between flow variations and A T caused by the thermistors' dissipation constants; (c) the motion or nucleation of bubbles near the thermistors; and (d) the thermal cross-talk that can occur between closely spaced thermistors. These findings suggest that future TEP devices, employed as immersible probes in a well-stirred solution, will need to address several problems. First, since temperature variations with time will occur for turbulent flow, better CMRRT values are needed. Second, since turbulent flow also imposes a spatial, fluctuating temperature gradient, closely spaced thermistors appear to be desirable. Therefore, higher average velocities and smaller excitation powers are required to reduce thermal cross-talk. However, reduced excitation power renders other sources of noise significant (17). Low-duty cycle, pulsed excitation may help, since the average power could be significantly reduced. In short, while experiments that directly address these issues in turbulent flow are desirable, the present study suggests that a limit for AT measurements in T E P s that are immersed in a well-stirred flow may not have been reached. Further, this work is also relevant to column devices; given their intrinsically
higher sensitivity of about 10 "C M-I, if comparable AT measurements can be made in more open columns with quiet, laminar flow and degassing where necessary and permissible, then significantly improved detection limits (below lo4 M) should be obtained.
LITERATURE CITED (1) Rich, S.; Ianniello, R. M.; Jespersen, N. D. Anal. Chem. 1979, 57, 204-206. (2) Tran-Minh, C.; Vallin, D. Anal. Chem. 1978, 50, 1874-1878. (3) Cooney, C. L.; Weaver, J. C.; Tannenbaum, S. R.; Faller, D. V.; ShieMs, A . In "Enzyme Engineering", Vol. 2, Pye, E. K.. Wingard, L. B., Jr., Eds.; Plenum: New York, 1974; pp 411-417. (4) Weaver, J. C.; Cooney, C. L.; Fulton, S. P.; Schuler, P.; Tannenbaum, S. 13.Biochim. Biophys. Acta 1976, 452,285-291. (5) Weaver, J. C.; Cooney, C. L.; Tannenbaum, S. R.; Fulton, S. P. I n "Biomedical Applications of Immobilized Enzymes and Proteins", Vol. 2. Chang, T. M. S., Ed.; Plenum: New York, 1977: pp 191-205. ( 6 ) Cooney, C. L.; Weaver, J. C.; Fulton, S. P.; Tannenbaum, S. R. I n "Enzyme Engineering", Vol. 3, Pye, E. K.. Wingard, L. B., Jr., Eds.; Plenum: New York, 1978; pp 432-436. (7) Mosbach, K.; Danielsson, B.; Borgerud, A,; Scott, M. Biochim. Biophys. Acta 1975, 403, 256-265. (8) Mattiasson, B.; Danielsson, B.; Mosbach, K . Anal. Lett. 1976, 9 , 2 17-234. (9) Mattiasson, 8.; Danielsson, B.; Mosbach, K . Anal. Lett. 1978, 9 , 867-889. (10) Danielsson, E.; Gadd, K.; Mattiasson, B.; Mosbach, K . Anal. Lett. 1978, 9 , 987-1001. (11) Mattiasson, 8. FEBS Lett. 1978, 85, 203-206. (12) . . Bowers. L. D.: Cannina, L. M.; Savers. C. N.; Carr. P. W. Clin. Chem. 1976, 22, 1314-131g (13) Bowers, L. D.; Carr, P. W. Clin. Chem. 1976, 22, 1427-1433. (14) Schmidt, H.-L.: Krisam, G.;Grenner, G. Biochim. Biophys. Acta 1978, 429. 283-290. . -~ (15) Mattiasson, B. FEBS Lett. 1977, 77, 107-1 10. (16) Mattiasson, B.; Borrebaeck, C. FEBS Lett. 1978, 85, 119-123. (17) Bowers, L. D.; Carr, P. W. Thermochim. Acta 1974, 70, 129-142. (18) Fulton, S. P. Master's Thesis, Massachusetts Institute of Technology, Cambridge, Mass., 1977. (19) Weaver, J. C.; Abrams, J. H. Rev Sci. Instrum. 1979, 50, 478-481. ~
RECEIVED for review September 19,1979. Accepted November 12, 1979. This work was supported by United States Public Health Service Biomedical Research Support grant no. S07RR-007047-10.
Oxalate Determination by Immobilized Oxalate Oxidase in a Continuous Flow System Renze Bais," Nicholas Potezny, John B. Edwards, Allan M. Rofe, and Robert A. J. Conyers Division of Clinical Chemistry, Institute of Medical and Veterinary Science, Frome Road, Adelaide, South Australia 5000
A rapid, one-step procedure is described for the determination of oxalate. Oxalate oxidase, immobilized in a continuous-flow system, converts oxalate to hydrogen peroxide which is detected by a color reaction with 4-aminophenazone. The assay Is Sensitive to oxalate concentrations as low as 5 1 M and is hlghty reproducible. A range of inorganic ions, alcohols, mons and polycarboxylic acids, sugars, and nltrogenous substances do not interfere with the assay. Although ascorbate and NADH did not affect the enzyme reaction, they did reduce the color formation with 4-aminophenazone. The immobilized enzyme system can, however, be readily adapted to more specific detection systems.
Oxalate is found in a wide range of biological and nonbiological materials ( I ) and is used in a variety of manufacturing ( 2 ) and analytical procedures ( 3 ) . In the determination of oxalate, techniques such as volumetric analysis, precipitation, 0003-2700/80/0352-0508$0 1 OO/O
solvent extraction, colorimetry, fluorimetry, chromatography, radioisotope methods, ion selective electrodes, enzymes, and mass spectroscopy have all been used ( I ) . Most of the published methodologies have the disadvantage that the sample must be pretreated before the actual oxalate analysis can be performed. This increases the complexity of the analytical procedure and consequently the sensitivity, specificity, and reproducibility of the determination often suffers. As analytical reagents, enzymes provide a degree of specificity and sensitivity available only in other more expensive chemical and physical methods such as mass spectroscopy. In addition, immobilization of the enzyme on an inert surface permits reutilization of the enzyme thus reducing the cost when used routinely ( 4 ) . If the immobilized enzyme is coupled with a continuous flow system, handling is minimized and reproducibility is enhanced. Oxalate decarboxylase (E.C. 4.1.1.2) and oxalate oxidase (E.C. 1.2.3.4) are two enzymes used in the determination of oxalate. However, the methods described involve either a S 1980 American Chemical Society
ANALYTICAL CHEMISTRY, VOL. 52, NO. 3, MARCH 1980
509
Immobilized 10 per hr
0 32 ml min f l o w c e l l
recorder
15 m m flowcell 520 nm colorimeter
Figure 1. Schematic diagram for the continuous flow analysis of oxalate
separate secondary enzyme reaction t o monitor product formation ( 5 , 6) or the use of radioactive tracer ( 7 , 8 ) ,and both are time-consuming and laborious. Oxalate oxidase prepared from plant tissue (9, 1 0 ) catalyzes t h e reaction:
(COOH)2 +
0 2
+
2C02
+ HzOZ
T h i s paper describes t h e immobilization of t h e purified enzyme onto an inert surface and its use as part of a continuous flow system for the determination of oxalate. We believe that the method outlined is the first, rapid, one-step procedure for t h e analysis of samples containing oxalate.
EXPERIMENTAL Chemicals and Materials. ['4C]oxalic acid (60 mCi per mmol) was purchased from the Radiochemical Centre, Amersham, England. Oxalic acid, .l-aminophenazone, 1,2-diaminoethane and glutaraldehyde (Electron Microscopy grade) were obtained from British Drug Houses, Poole, England; hyamine and succinic acid were obtained from Calbiochem-Behring (Aust.). Pty Ltd, N.S.W., Australia; l-chloro-2,3-epoxypropane (epichlorohydrin) and boron trifluoride diethyletherate (boron trifluoride etherate) from Aldrich Chemical Co., Milwaukee, Wis. Dichloromethane (Ajax Chemicals, N.S.W., Australia) and diethyl ether (May and Baker) were redistilled and stored over calcium hydride. Horseradish peroxidase (Type 1)was obtained from the Sigma Chemical Co., St. Louis, Mo. Nylon 6 tubing (1-mm internal bore) was from Portex Ltd, Hythe, Kent, U.K. Procedures. A s s a y of Oxalate Oxidase. The enzyme activity was assayed using the same method as described by Rofe et al. (11) for oxalate decarboxylase. The I4CO2which is formed from ['4C]oxalic acid, is absorbed into hyamine and counted after the addition of scintillant. Unless otherwise stated, enzymic assays were carried out in 0 05 M sodium succinate buffer, pH 3.5, containing 1 mM ['4C]oxalate (0.2 ICi per mL). Preparation of Oxalate Oxidase. Oxalate oxidase was prepared from the roots of 10-day old barley ( H o r d e u m uulgare) seedlings by a modification of the method of Chiriboga (12). Roots (1500 g) were collected and homogenized with two volumes of water in a Waring Blendor for 2 min. The extract was stirred for 6 h with a n overhead stirrer and the insoluble material removed by filtration through gauze, followed by centrifugation. The supernatant was concentrated using an Amicon pressure concentrator and dialyzed overnight against 0.005 M potassium phosphate buffer, pH 7.0. Insoluble material was removed by centrifugation and the supernatant loaded on a DEAE-Sephadex column (24 cm X 2.5 cm). The column was washed with the dialysis buffer until the eluant was devoid of protein and then the bound protein was eluted with 0.1 M KC1 in 0.005 M potassium phosphate buffer,
pH 7.0. The peak fractions were concentrated using the Amicon concentrator and loaded onto a Sephadex G-200 column (60 cm X 3.5 cm) equilibrated with 0.02 M potassium phosphate buffer, pH 7.0, containing 0.02 M KC1. The active fractions were again concentrated using the Amicon concentrator and stored a t -20 "C. Prior to immobilization, any catalase in the preparation was denatured by heating the material to 80 "C for 3 min and removing the precipitated material by centrifugation. This preparation gave 20 mg of purified enzyme with a specific activity of 22 units per mg protein which represented a 16% yield of the initial extract. I m m o b i l i z a t i o n of Oxalate Oxidase. A modification of the method described by Morris et al. (13) was used. (a) 0-Alkylation of Nylon Tubing. Three meters of nylon tubing were filled with a 12.5% (w/v) solution of triethyloxonium tetrafluoroborate (14) in dichloromethane and incubated a t 25 OC for 15 min. The tubing was washed *ith dry dichloromethane and used immediately. (b) Amine Substituted Nylon Tubing. The 0-alkylated tubing was filled with diaminoethane and incubated for 2 h a t room temperature. Excess diamine was removed by washing the tube thoroughly with water. (c) Preparation of Glutaraldehyde-Activated Tubing. A 5% (w/v) solution of glutaraldehyde in 0.2 M borate buffer, pH 8.5, was pumped through the substituted nylon tube at room temperature for 15 min. The tube was then washed by perfusion with 0.15 M NaCl in the borate buffer for a further 15 min. (d) Enzyme Immobilization. The glutaraldehyde activated tubing was immediately filled with a solution of enzyme (60 units) in 0.02 M potassium phosphate buffer, pH 7.0, containing 0.02 M KC1. This was incubated overnight a t 4 "C and the tubing then washed with 0.05 M sodium succinate buffer, pH 3.5, for 15 min. Eighty percent of the activity remained bound to the tubing; 1.5 m of tubing was used in the continuous flow system and stored in a sealed damp plastic bag at 4 "C when not in use. Under the experimental conditions described in this paper, we have observed no noticeable deterioration in activity of the bound enzyme over 5 months of usage. Instrumentation. A Technicon AutoAnalyzer AAII system was assembled as shown in Figure 1. A sampler I1 operating a t 10/h with a 1:2 sample-to-wash ratio was used to obtain the data in this report. After being passed through the tubing containing the immobilized enzyme, the sample is mixed with the color reagent. This reagent is modified from that described by Trinder (15) and consists of 0.4 M sodium phosphate buffer, pH 7.0, containing 0.05 g of 4-aminophenazone, 0.1 g phenol, and 1 mg of peroxidase per 100 mL. The AAII colorimeter was equipped with a 520-nm filter and a 15-mm (1.5-mm i.d.) flowcell. A Yokogawa Technicorder F recorder with a chart speed of 20 cm per h was adapted to the AAII colorimeter.
510
ANALYTICAL CHEMISTRY, VOL. 52, NO. 3, MARCH 1980
12c
1oc
8C
W
m c
5
6C
(0
n
40
20
3
5
4
7
6
8
9
10
PH
Figure 2. pH profile for the hydrogen peroxide color reaction. The buffers used were citrate (O),potassium phosphate (A)and Tris-CI (0)
RESULTS T h e system as described is based on the conversion of oxalate by the immobilized oxalate oxidase to hydrogen peroxide which is detected by a color reaction. In the original paper describing the color reagent ( 1 5 ) , Trinder did not mention whether p H had any effect on the color production, and as our system had a buffer of pH 3.5 mixing with the color reagent, the p H effect was investigated further. Samples of hydrogen peroxide were passed through the system, mixed with the color reagent at various p H values, and the resultant color was determined. The results (Figure 2 ) indicate that the optimum pH is between 6 and 7. In addition to the effect of p H , it was possible to show that between the concentrations of 0.1 and 0.5 M potassium phosphate buffer a t pH 7.0, there was less than a 4% variation in color formation. Therefore, it was decided to use a 0.4 M potassium phosphate buffer, pH 7.0, as the color reagent buffer
I
because there was negligible change in the pH when this was mixed with the 0.05 M sodium succinate buffer, pH 3.5, and secondly, this concentration did not affect the resultant color formation. The effect of temperature on the activity of the immobilized enzyme is another variable of the system which was examined. This was assessed by placing the coil in a water bath, allowing equilibration a t the required temperature, and then determining the oxalate concentration. Between 20 and 37 "C there was less t h a n 8% variation in activity, so careful control of the enzyme coil temperature was not warranted. This was acceptable as the assay is an end-point determination and, provided the timing is adequate, the reaction is independent of temperature. Figure 3 is a typical recording from the immobilized oxalate oxidase procedure, and illustrates the characteristics of continuous flow sampling, standards, reproducibility, and interaction between high and low samples. The standards shown are between 0.1 and 0.5 mM oxalate which is in the range for urinary oxalate determinations ( 5 ) . When the criterion that the peak height should be a t least twice the noise level is applied, the lower limit of detection with the system is 5 pM oxalate; and this is below the reported levels in serum (16). Moreover, there is a lack of carryover from high to low samples (Figure 3) and a linear relationship between the oxalate concentration and the color formation of the indicator reaction. For 20 assays a t a level of 0.25 m M oxalate, the intrabatch coefficient of variation is less t h a n 2% and the interbatch variation is 'i7 ' ~ . T h e mechanism by which reducing compounds interfere with the peroxidase coupled indicator reaction has been described (17). In order to determine whether certain compounds interfered with either the enzymic activity or the color formation, the following compounds (20 mM) were run through the system individually and also added to a 0.2 m M standard oxalate solution: lactate, pyruvate, citrate, acetate, ascorbate, NAD, NADH, glucose, fructose, xylitol, urea, creatinine, glycine, serine, glutamate, glycollate, glyoxylate, ethylene glycol, ethanol, SO4'-, PO?-, CO;', HC03 , C1-, Mg2+, Na+, K+, and Li+. Only NADH and ascorbate had any effect on the oxalate results and both were shown to reduce the color reaction and not affect the enzyme activity. When the determination of oxalate in aqueous samples is required, the method described reduces the mundane task of sample preparation and subsequent oxalate analysis to one
0.5
Oxalate rnM
6
I
11
M I-
0 Y
1
Figure 3.
~
.
_
_
_
_
Representative recording of the oxalate determination showing standards followed by high-to-low samples
_
-
Anal. Chem. 1980, 52, 511-514
of simply loading the sample plate of the analyzer. Specificity, sensitivity, and reproducibility result from the combination of immobilized oxalate oxidase and continuous flow analysis. The specificity is conferred by the use of the enzyme, oxalate oxidase, which does not react with substances that have been reported to interfere with other procedures ( I ) . This specificity may be compromised in complex matrices by the presence of interfering substances such as ascorbic acid which affect the indicator reaction. This possibility, however, does not detract from the potential usage of the method. Indeed, if required, the continuous flow system is capable of being further improved through the use of more selective dialysis membranes or by the inclusion of alternative detection systems based on either specific secondary enzymes or selective electrode systems.
51 1
Vogel. A . I. "A Textbook of Quantitative ::norganic Analysis", Longmans: London, 1962: pp 243, 249, 320, 577. Gray, D. N.; Keyes, M. H.: Watson, B. A,oal. Chem. 1977, 49, 1067A1078A. Costello, J.: Hatch, M.; Bourke. E. J . Lab. Clin. Med. 1976, 8 7 , 903-908. Kohlbecker, G.; Butz, M.; Heinz. F . P r o c X Int. Congr., Clin. Chem. 1978, 107. Hallison, P. C.: Rose, G. A . Clin. Chim. Acta 1974, 55, 29-39. Kohlbecker, G.;Richter. L.; Butz, M. J Clin. Chem. Clin. Biochem. 1979, 17, 309-314. Srivastava. S. K.: Krishnan, P. S. Biochem. J . 1962, 8 5 , 33-38. Chiriboga, J. Biochem. Biophys. Res. Commun. 1963, 11, 277-282. Rofe. A . M.: Edwards. J. B. Biochem. Med. 1976. 16. 277-283. Chiiboga, J. Arch. biochem. biophys. 1966, 716, 516-523. Morris, D. L.: CamDbell, J.: Hornby. W. E. Biochem. J . 1975, 147, 593-603. Meerwein, H. Org. Synth. 1966, 4 6 , 120-121. Trinder, P. A n n . Clin. Biochem. 1969, 6 , 24-27. Hodakinson. A,: Wilkinson. R. Clin. Sci. Mol. Med. 1974. 46. 61-73. Gochman, N.; Schrnitz, J. M. Clin. Chem 1972, 13, 943-950
L I T E R A T U R E CITED
RECEIVED for review August 6, 1979. Accepted November 26,
(1) Hcdgkinson, A. "Oxalic Acid in Biology and Medicine", Academic Press: New York, 1977. ( 2 ) Florio, F. A., Pakel, G. "Kirk-Othmer Encyclopedia of Chemical Technoiogy", pnd ed,, vel, 14; Wiley and Sons: New York, 1967: pp 356-373.
1979. T h e financial assistance of the National Health and Medical Research Council of Australia and Roche Products P t y Ltd of Australia is gratefully acknowledged.
Kinetics and Mechanism of the Extraction of Copper with 2-Hydroxy-5nonylbenzophenone Oxime Stephen P. Carter and Henry Freiser" Department of Chemistry, University of Arizona, Tucson, Arizona
8572 1
The kinetics of the extraction of Cu(I1) into CHCI, solutions of 2-hydroxy-5-nonylbenzophenone oxime, a high molecular weight hydroxyoxime chelating agent, has been found to be first order in metal ion, second order in extractant, and inverse first order in hydrogen ion. The results are consistent with a mechanism in which the rate-determining step involves chemical reactions in the aqueous phase, as was found in other chelate extraction systems we have studied. Interfacial reactions, invoked by earlier workers for such extractions, are not required by this mechanism.
Solvent extraction processes represent a separation technique of continuing high significance for inorganic and organic trace analysis as well as for large scale process recovery (1-5). T o gain insight into existing processes, new studies of equilibrium and kinetic aspects are of essential importance. In addition, a greater understanding of solvent extraction systems can be used for the optimization of processing and design. While much useful information has been reported on details of extraction equilibria, the study of extraction kinetics has not been as thoroughly investigated as its importance demands. Elucidation of extraction behavior a t the molecular level could be applied to the theoretical understanding of modern analytical methods such as liquid chromatography, a n d also to the large scale commercial metal recovery processes, as well as to analytical extraction methodology. In earlier work from this laboratory, we reported a series of kinetic studies of chelating extractions in which it was demonstrated that, even using poorly water soluble extractants such as dithiazone (distribution constant K b = IO"), the slow rate of extraction was attributable to the homogeneous chemical reactions, namely, the formation of metal chelates 0003 2/OO/80/0352-05 11$01 O O / O
in the aqueous phase (6). These studies resulted in the felicitous discovery of a unique and simple method for the study of the kinetics and mechanism of metal chelate formation and dissociation particularly when, due to very low water solubility of neutral chelates, other fast kinetic techniques are not applicable (7-10). An interesting and important fundamental question to be posed is that of the possible change in reaction mechanism that might occur on increasing the hydrophobicity (Le., greater KD,) of the ligand. At what point will the aqueous phase extractant concentration be so low as to relocate the reaction site from the bulk aqueous phase to either the interface or the organic phase? Such questions will have great bearing on the design of any new extractants. T o investigate this point, we decided to make a detailed study of the extraction of Cu(I1) with LIX Reagents. These materials, which are hydroxyoximes, were introduced in the 1960s by General Mills for the hydrometallurgical refining of copper ( 5 , 11). T h e kinetics of copper-LIX extractions are of great practical importance and have been the subject of some study. Most of the theoretical studies have been performed with 2-hydroxy-5nonylbenzophenone oxime (LIX65N) and 5,8diethyl-7-hydroxy-dodecan-6-one oxime (LIX63), which are shown below.
-0li
LIX63
LIX65N
Previous workers, pointing to the low aqueous solubility of c 1980 American Chemical Society
