Anal. Chem. 1982, 5 4 , 217-220
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Development and Application of a Histidine-Selective Biomembrane Electrode Paul M. Kovach' and M. E. Meyerhoff* Department of Chemlstty, The Unlversl@of Michigan, Ann Arbor, Michigan 48109
A highly Selective hlstldlne blomembrane electrode has been prepared by lmmoblllzlng the enzyme hlstldlne decarboxylase (E.C. No. 4.1.1.22) at the surface of a potentlometrlc carbon dioxide sensor. The enzyme employed was extracted from Lactobacillus 30a. The use of concentrated enzyme extract rather than Intact bacterial cells Is shown to yield bloelectrodes wlth Improved response characterlstlcs. The resulting enzyme-based sensor responds linearly to the logarlthm of L-hlstldlne concentration between 3 X lo-' and 1 X lo-' md/L with a slope typlcally of 48-53 mV/decade and a useful ltfetlme of over 30 days. The electrode can be used to assay hlstldlne directly In urine samples wlth good analytical recovery (av 104.8%) and correlation ( r = 0.91) wlth a fluorometric procedure.
The development of enzyme-, bacterial-, and tissue-based biomembrane electrodes selective for physiologically important compounds continues to be an expanding and exciting area of research (1-5). T o be of practical analytical utility, such electrodes must possess certain desirable response characteristics including high selectivity, rapid response times, freedom from inhibitor interferences which may be present in real samples, and long-term stability. We now report the development of a new histidine-selective biomembrane electrode which appears to meet these requirements (histidine refers to the physiologically active form, L-histidine). Several workers have previously reported histidine-selective electrodes based on immobilized enzyme or bacterial cell catalyzed reactions (6-8). Buck et al. utilized histidine ammonialyase (E.C. No. 4.3.1.3) from Pseudomonas s p . to develop a histidine electrode which relied on a potentiometric ammonia gas sensor as the detector (6). The intact cells were also employed for the construction of the electrode, but selectivity was poor because of the presence of other deaminating enzymes (7). White (8),used histidine decarboxylase purified from Ch. Welchii in conjunction with a potentiometric carbon dioxide sensor to prepare a histidine electrode. The electrode had reasonable response times (approximately 10 min), a good dynamic response range, and selectivity over other common amino acids. However, no information concerning response to histidine derivatives or similar compounds was provided. In addition, in that work, and in other histidine electrode reports, no real sample analytical utility was demonstrated. Histidine measurements in serum and urine samples are associated with the diagnosis of histidine metabolism disorders, particularly histidemia (9, IO). Elevated levels in physiological fluids signal this hereditary disease. Current ion exchange chromatography (11) or fluorometric reaction procedures (12,13) used to determine histidine in clinical samples are complex and time-consuming and, in the case of fluorescence, require caustic analytical reagents. In this paper, the development, study and analytical apPresent address: Department of Chemistry, Indiana University, Bloomington, IN 47405. 0003-2700/82/0354-0217$01.25/0
plication of a practical histidine electrode is described. The electrode is prepared by immobilizing a concentrated protein extract containing the enzyme histidine decarboxylase (E.C. No. 4.1.1.22) isolated from Lactobacillus 30a, a t the surface of a potentiometric carbon dioxide sensor. Attempts to utilize the intact bacterial cells yield electrodes with poor response characteristics. Isolated enzyme-based electrodes have high slopes, short response times, and excellent long-term stability. The electrode is highly selective for histidine over all other naturally occurring amino acids, imidazole derivatives and similar compounds both in the presence and absence of pyridoxal phosphate (PLP). Evaluation of this electrode for biological measurements is made via recovery studies on spiked urine samples and correlation studies vs. a fluorometric histidine method on pooled urine samples.
EXPERIMENTAL SECTION Apparatus. An Orion Model 95-02 carbon dioxide gas sensing electrode was used to construct all biomembrane electrodes. Potentiometric measurements were made with a Fisher Accumet Model 620 pH meter in conjunction with a Houston Instruments strip chart recorder. All calibration and sample measurements were made at 25 "C. Fluorometric histidine measurements in urine samples were made with a JY-3 spectrofluorometer. Enzyme solutions were concentrated by using a Micro-Pro-Dicon molecular protein dialyzer-concentrator (Bio-molecular Dynamics, Beaverton, OR). Reagents. Lactobacillus 30a, ATCC No. 33222, was obtained from the American Type Culture Collection (Rockville, MD). Chemicals used to prepare the growth medium and all analytical buffers, etc. were reagent grade. All aqueous solutions were prepared with distilled-deionized water. L-Histidine, free base or hydrochloride, o-phthaldehyde, glutaraldehyde, bovine serum albumin and assorted amino acids, and imidazole derivatives were obtained from Sigma Chemical Co. (St. Louis, MO). Working buffers of 0.2 and 0.4 mol/L ammonium-citrate, pH 4.80, were prepared fresh weekly. Ammonium-acetate buffer, 0.2 mol/L, pH 4.80, was used during the initial enzyme extraction step. Bacterial Cell Preparation and Enzyme Extraction. Lactobacillus 30a uiere grown at 37 "C in a double strength crude medium (medium 1111) (14). Optimum growth was obtained without aeration or shaking action. The cells were harvested by centrifugation at 2000g. For initial bacterial electrode studies, the cells were washed several times in working buffer, and subsequently, microliter quantities of the packed cell slurry were immobilized on the C02 electrode. For enzyme extraction, a modification of the procedure outlined by Rosenthaler et al. (15)was employed: Approximately 10 mL of packed cells were dried with acetone and washed with ether. The dried cells were ground into a fine powder and then approximately 1.2 g of dried cells was suspended in 15 mL of ammonium-acetate buffer and stirred for 5 h. The cell debris was then centrifuged and the resulting supernatant containing a crude extract of the enzyme was concentrated to a final volume of 0.750 mL and simultaneously dialyzed vs. 0.2 mol/L ammonium-citrate buffer, pH 4.80, using a Micro-Pro DiCon dialyzer-concentrator. The 0.750 mL of concentrated protein was rediluted to wash debris from the walls of the concentrator and then reconcentrated to a final volume of 0.100 mL. This concentrated enzyme extract was stored at 4 "C and had 156 units/mL of decarboxylating @ 1982 American Chemical Society
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ANALYTICAL CHEMISTRY, VOL. 54, NO. 2, FEBRUARY 1982 enzyme B S A & lutarddehyde solns. m s i l l c omembrane n e
stopper rubber'k
2 \ pipette .
bulb
Figure 1. An illustration of the technique used to immobilize hlstldlne
decarboxylase uniformly at the surface of a carbon dioxlde electrode. activity toward L-histidine, as determined by kinetic experiments using the C02 sensing electrode (16) (one unit liberates 1pmol of COz/min at 25 "C). Preparation of Biomembrane Electrodes. Bacterial electrodes were prepared as originally described by Rechnitz et al. (17). Ten to 15 microliters of Lactobacillus 30a slurry was applied to the surface of a carbon dioxide membrane and then sandwiched in place with a piece of outer dialysis membrane. The resulting electrode was conditioned in working buffer for several hours in an attempt to remove background COP The enzyme-based sensor was prepared by immobilizing the concentrated enzyme extract preparation via the cross-linking method of Mascini and Guilbault (18). To chemically immobilize the enzyme to the silicone rubber membrane of the COz electrode, we inverted the outer body (with membrane). A slight vacuum was applied to the internal portion of the barrel by using a pipet bulb-rubber stopper assembly. Figure 1schematically illustrates this arrangement. Suction applied by the pipet bulb causes the silicone membrane to have a slightly concave shape. Ten microliters of enzyme extract, 10 pL of bovine serum albumin (BSA, 15% in water), and 2 pL of glutaraldehyde (10%) were placed in the center of the membrane and mixed rapidly with a glass rod. The mixture was allowed to stand for 5 min. Suction was then released and the internal solution and pH electrode were placed into the barrel and secured in place. Cross-linking was allowed to continue for an additional 30 mln before following the water-glycinewater washing steps previously described (18). This procedure allows for a much more even distribution of the enzyme at the center of the membrane since the enzyme is prevented from accumulating at the crevices between the membrane and the electrode cap. Following assembly the electrode was conditioned and always stored in the 0.2 mol/L working buffer. Procedure for Evaluating Electrode Response and Performing Histidine Measurements in Urine. The electrode was calibrated daily by making standard additions of 0.1 mol/L histidine into 25 mL of working buffer. Between calibrations or discrete sample determinations, the electrode was placed into a large volume of well-stirred working buffer to gain base line recovery. Selectivity study data were obtained by making a standard addition of a given compound into a known volume of buffer so that the final concentration of the compound was 4 X lo-' mol/L. The change in potential from the base line value (no histidine) upon this addition was noted after 5 min. Recovery data for a urine sample spiked with various amounts of histidine was obtained as follows: A large volume of normal urine was divided into five 50-mL aliquota. Standard additions of histidine were made to four of these portions. Twenty-five milliliters of each sample was then diluted 1:l with 0.4 mol/L ammonium-citrate buffer, pH 4.80. These samples were degassed with N2for 10 min prior to assay. Steady-state potentials recorded for each sample were used to determine histidine values from a prior calibration curve using a least-squares fit of calibration points in the concentration range of the unknowns. Potentiometric determinations on randomly pooled human samples were done in the same manner. Unidentifiable pooled samples were obtained from the University of Michigan's Student
4
- log
3 [Histidlne]
2
, rnol/L
Figure 2. Typical Calibration curves for bacterial (A)and enzyme extract based (a)histidine selective biomembrane electrodes.
Health Services Facility. Fluorometric histidine measurements were made by adapting the serum histidine method described by Arbrose et al. (12) to urine samples.
RESULTS AND DISCUSSION
Our attempt to utilize the histidine decarboxylating activity of Lactobacillus 30a to develop an L-histidine specific electrode was based on earlier biochemical studies of several investigators. Rodwell (19) first reported the decarboxylation of several amino acids by this bacterial strain. He observed C02 production from lysine, arginine, and histidine, however, histidine activity was 15-20 times greater than that for the other substrates. Snell-and co-workers (14,20) purified and studied the catalytic properties and structure of histidine decarboxylase from Lactobacillus 30a. They demonstrated that this enzyme is unique in that it is the only known decarboxylase which does not require PLP as a cofactor for catalytic activity. With this in mind, we felt it might be possible to use intact Lactobacillus 30a cells to prepare a histidine-sensitive electrode and that we could ultimately gain selectivity over the other amino acids simply by not adding PLP to the assay buffer. We, therefore, assembled several bacterial electrodes by entrapping the intact cells between a dialysis membrane and the surface of a C02-sensing electrode. Figure 2 shows a typical calibration for such an electrode. Response slopes were usually 35-40 mV/decade with linear response between 6 X lo4 to 7 X mol/L. Response times to reach equilibrium steady-state potentials were extremely long, 15-20 min even at high histidine concentrations. Complete washout to typical base line potentials representative of low C 0 2 levels was impossible apparently due to the bacterial cells partially clogging the gas membrane surface and/or continued resting metabolic C02production by the cells. This problem has been encountered with earlier bacterial electrodes (5,7)and, along with the relatively low catalytic activity of the intact cells toward histidine, did not allow us to prepare electrodes with acceptable response characteristics. Attempts to induce more catalytic activity in the intact cells by increasing the concentration of histidine used in the growth medium failed. Extraction and concentration of the histidine decarboxylating activity from the cells appeared to be the most promising approach to making a practical histidine electrode. Figure 2 shows a typical calibration curve for a histidine electrode prepared with a cross-linked form of the concentrated enzyme extract. Linear response is obtained over a concentration range of 3 x lo-* to 1 X mol/L. The practical detection limit for this sensor is 1.0 X low4mol/L. Although response
ANALYTICAL CHEMISTRY, VOL. 54, NO. 2, FEBRUARY 1982
3.7 x 1 r 3 00.
1.8 16-3 5
20
.
1.0 x 10-4 baseline
2
10
18 day
26
34
I
Figure 3. Stability study of absolute potentials observed with new histidine electrode at varying hlstidine concentrations, moi/L.
below this level was readily observed, this response was not reproducible due to the poor base line recovery characteristics of the electrode. The working pH chosen, pH 4.80, was based on the optimum conditions reported by Snell (14) for the isolated enzyme activity. Ammonium citrate buffer rather than ammonium-acetate (used by Snell) was used because the COz electrode has some response to the acetic acid-acetate buffer system (21). Under these conditions, response times to reach steady-state potentials were relatively rapid; e.g., 10 min at concentrations 4 X lo4 mol/L. While adequate for measurement purposes, these response times are not as fast as desired and in fact are limited by the poor response and recovery times of the present pH electrode-based COz sensor. Current work on a polymer membrane electrode-based COz sensor with dramatically improved response times could overcome this existing limitation (22).
No major attempt to study the pH optimum for activity of the immobilized enzyme was made. This was because, from preliminary studies, it once again became apparent that the response times and sensitivity of the histidine electrode were predominantly limited by the innate properties of the pCOz base sensor and not by the catalytic activity of the immobilized enzyme. This was evidenced by experiments in which increasing concentrations of enzyme extract were immobilized. After a point, additional enzyme activity does not yield any enhanced response properties. Moreover, the optimum response for the pCOz sensor is reported to be at pH 4.50 (21) which is close to the value we used throughout these studies. Long-term stability of the enzyme extract-based electrode was excellent. Figure 3 illustrates the stability of the electrode over a 41-day period, The absolute calibration potentials and slope did not significantly change even though the electrode was stored in buffer at room temperature. Mter 30 days, some loss in the catalytic activity was observed as seen by the premature curvature of the calibration plots at high histidine concentration and slightly longer response times. In certain instances, the lifetime of some of the electrodes prepared was limited by the lifetime of the silicone rubber gas membrane itself which, with the enzyme immobilized on it, tended to degrade after 20-30 days of constant use. Reproducibility within a given day’s calibration was also excellent. We normally found calibration and unknown potentials to be reproducible to f l mV over the course of the entire day. At low histidine concentrations, this precision can only be attained if between measurements, the electrode is allowed to wash all the way back to base line potential values (usually 15-20 min). This again is limited by the recovery time of the current COz sensor.
219
Table I. Summary of Selectivity Data for New Histidine Selective Biomembrane Electrode A E , ~ ~ V A E , ~ ~ V compou nda with PLPc (4x mol/L) t 91.9 t 91.9 histidine t 3.7 t 3.1 1-methylhistidine t 2.4 3-methylhistidine -0.4 imidazole lactate t 25.0 imidazole acetated t 0.1 imidazole pyruvate t 3.4 N-acetylhistidine t 11.0 t 5.0 urocanic acid t 0.1 0.0 arginine 0.0 - 0.4 glycine -0.6 ornithine -0.2 -0.2 lysine -0.4 glutamic acid -0.6 hydroxyproline - 0.3 -0.5 phenylalanine -0.5 glutamine -0.6 asparagine -0.3 -0.3 tryptophan -0.3 threonine -0.3 methionine -0.5 -0.3 tyrosine - 0.5 -0.3 proline 0.0 urea a Where appropriate, only biologically active L form of amino acid or derivative was tested. Potential change relative to base line potential (no histidine) after 5 min. PLP present at 1.0 X mol/L. Predominantly nonenzymatic response (see text). Selectivity of the histidine electrode was examined on two fronts. First, the electrode was evaluated for response to a wide range of naturally occurring amino acids, imidazole derivatives, and other biomolecules by directly adding such compounds to the working buffer. Absolute potential changes from base line values were recorded for each compound added at 4 X mol/L. Table I summarizes the results of that study. For each compound tested, the electrode was also checked for response toward histidine in the presence of the compound. In no instance was histidine response inhibited by any of these molecules. In addition, since the enzyme preparation used was only a crude extract of the bacterial cells, we felt other decarboxylases, particularly lysine and arginine (as reported by Rodwell (19)),could also be present in the immobilized biocatalytic layer. However, these activities would only be noticed if PLP was present in the test solution. Thus, we also checked the selectivity of the electrode in the presence of PLP toward known PLP dependent decarboxylase substrates. The results of those studies are also listed in Table I. It is evident that either in the absence or presence of PLP, the electrode displays remarkable selectivity for histidine, Lysine and arginine response was not observed even with PLP probably because the relative activity of their decarboxylases in the cross-linked membrane was minimal. Urocanic acid and imidazole-acetate are the two principal interferences noticed, although it was subsequently determined that imidazole-acetate response was not due to decarboxylating activity but rather to the innate response of the COz sensor to certain organic acids which can diffuse through the silicone rubber membrane (21,23). The urocanic acid response was definitely due to catalytic activity. Fortunately, for practical purposes, both of these compounds normally exist in physiological fluids a t levels 2 orders of magnitude less than histidine (11). Very slight activities observed toward 1- and 3-methylhistidine and N-acetylhistidine may actually be due to trace histidine impurities in these compounds. Overall, the selectivity observed appears to be adequate for real sample measurements.
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ANALYTICAL CHEMISTRY, VOL. 54, NO. 2, FEBRUARY 1982
Table 11. Summary of Histidine Analytical Recovery Experiments on Spiked Urine Sample sample 1
2 3 4
amt added," amt found,b mmal mmol % recovery 0.0250 0.0500 0.1000 0.2000
av
0.0266 0.0566 0.1010 0.1972
106.4 113.2 101.0 98.6 104.8
a Standard additions made to a urine sample initially containing 1.83 X mol/L histidine based on two determinations by potentiometric method. Average of two determinations.
Table 111. Results of a Comparison Study for Potentiometric and Fluorometric Determinations of Histidine in Urine Samples sample no.
fluorometric," potentiometric," mmol/L mmol/L
0.57 0.60 0.91 0.83 0.48 0.52 0.55 0.40 1.01 1.47 0.38 0.37 0.09 0.09b 0.82 1.13 0.59 0.99 0.87 1.05 0.84 1.05 0.81 0.66 0.66 0.64 0.80 0.84 0.43 0.61 y = 1.28~ - 0.05 r = 0.91 Estimated value a Average of two determinations. from nonlinear portion of calibration curve. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 Equation of Line:
To demonstrate real sample application, we undertook histidine recovery studies on a spiked urine sample. Table I1 summarizes the results of that study. Excellent recovery was observed indicating that the normal constituents of urine do not interfere in any way with the detection of histidine. Consequently, aqueous standards can be used for calibration purposes. Although the COPcontent of urine is normally quite low (24),we found that it was a good idea to degas each sample for about 10 min with N2 before making electrode measurementa. This simple step completely eliminates any unlikely, yet potential COz interference. We further wanted to demonstrate that this new histidine electrode functions effectively as a urinary histidine sensor by comparing potentiometric analytical results obtained on urine samples to an o-phthaldehyde reaction fluorometric method (12).Table I11 summarizes the results obtained for 15 nonidentifiable pooled human urine samples. Fairly good correlation was observed on most samples; however, some
positive bias was apparent with the electrode method. Values obtained for both methods were predominantly within the normal range expected for urine specimens (approximately 130-2100 pmol/L (25)).The lower values obtained for the fluorometric method on certain samples may be due to the presence of some type of quenching agent in these samples. In any event, it is clear that the electrode functions reasonably well in urine samples and yields values which are in close agreement with an accepted method. In summary, a practical, highly selective histidine biomembrane electrode has been developed which has response properties desirable for real sample histidine determinations. Through the use of a simple enzyme extract of bacterial cells, it has once again been demonstrated that highly purified and costly enzyme reagenta are not necessarily required to prepare useful selective biomembrane electrodes. Hopefully, this new histidine electrode will find wide use as a simple alternative to current histidine assay methods.
LITERATURE CITED Fung, K. W.; Sung, G.; Kuan, S. S.; Guilbault, G. G. Anal. Chem. 1979, 51, 2319-2324. Arnold, M. A.; Rechnltz, 0. A. Anal. Chem. 1980, 52, 1170-1174. Rechnitz, G. A.; Arnold, M. A.; Meyerhoff, M. E. Nature (London) 1979, 278. 446. Kobos, d. K.; Ramsey, T. A. Anal. Chlm. Acta 1980, 121, 111-118. Kobos, R. K. I n "Ion-Selective Electrodes in Analytical Chemlstry, Voi. 11"; Freiser, H., Ed.; Plenum Press: New York, 1980; Chapter 1. Walters, R. R.; Johnson, P. A.; Buck, R. P. Anal. Chem. 1980, 52, 1684-1 890. Walters, R. R.; Moriarty. B. E.; Buck, R. P. Anal. Chem. 1980, 52, 1880-1684. Whlte, W. C. Ph.D. Dlssertatlon, The University of New Orleans, New Orleans, LA, 1977. Ghadlmi, H.; Partington, M. W.; Hunter, A. New Engl. J. M e d . 1961, 265, 221-224. Auerbach, V. H.; DiGeorge, A. M.; Baldridge, R. C.; Tourteilotte, C. D.; Brlgham, M. P. J. Pedlst. 1962, 60, 487-497. Wadman, S. K.; DeBree, P. K.; Van Der Heiden, C.; Van Sprang, F. J. Clln. Chlm. Acta 1971, 31, 215-224. Ambrose, J. A.; Crimm. A.; Burton, J.; Pauilin, K.; Ross, C. Clin Chem. (Wlnsfon-Salem, N.C.)I W S , 15, 381-366. Gerber, D. A. Anal. Blochem. 1970, 34, 500-504. Chang, G. W.; Snell, E. E. Blochemisfry 1968, 7 , 2005-2012. Rosenthaler, J.; Guirard, B. M.; Chang, G. W.; Snell, E. E. Proc. Natl. Acad. Scl. 1965, 54, 152-158. Toneili, D.; Budinl, R.; Gattavecchia, E.; Glrotti, S. Anal. Biochem. 1981. .- - ., 1. 1. 1. , 189-194 . - - .- . . Rechnitz, G. A.; Riechel, T. L.; Kobos, R. K.; Meyerhoff, M. E. Science 1978. 199. 440. Mascinl, M.'; Gullbault, G. G. Anal. Chem. 1977, 49, 795-798. Rodwell, A. W. J. Gen. Mlcroblol. 1953, 8 , 224-232. Chang, G. W.; Snell, E. E. Biochemlsfry 1068, 7 , 2012-2020. "Instruction Manual for Carbon Dioxlde Electrode"; Orion Research, Inc.: Cambridge, MA, 1978. Greenberg, J.; Meyerhoff, M. E., Department of Chemistry, Unlversity of Mlchlgan, unpublished results, 1981. Kobos, R. K.; Meyerhoff, M. E., Parks, S., Department of Chemistry, University of Michigan, unpublished results, 1981. Tietz, N. 8.; Siggaard-Andersen, 0. I n "Fundamentals of Clinical Chemistry"; Tletz, N. W., Ed.; W. B. Saunders: Philadelphia. PA, 1976 Chapter 18. Ibbott, F. A. I n "Clinical Chemistry, Princlples and Technics"; Henry, R. J., Canno, D. C., Winkelman, J. W., Eds.; Harper and Row: Hagerstown, MD, 1974; Chapter 18.
RECEIVED for review August 31,1981. Accepted October 20, 1981. Acknowledgment is made to the National Institutes of Health for support of this research (GM-2882-01).
