Anal. Chem. 1983, 55, 1929-1933
AND/IOR). These two functions exhibit the best perfonnance based on the FOM. In order of decreasing performances based on FOM the next best metrics are Grotch, XOR, and then AND. For comparison the NAND metric was used. This is neither a similarity nor a dissimilarity metric, and its complete lack of utility is reflected in the exceedingly poor FOM. Weighted List Positions Results. The final column of Table I11 contains FOMs for the various binary metrics using l / j weighting of the list index positions. It is seen that the results present the metrics in exactly the same order as for unweighted evaluation. The weighted FOMs are slightly higher than the unweighted values. This is caused by the gradual decrease of FOM with hit-list length. In the weighted case the list index positions at the top of the hit-list are given more impact on the FOM, which makes the resulting FOM higher than that, for an unweighted evaluation. A later publication will consider optimal weighting of list index positions in more detail.
COIVCLUSION A quantitative method for the evaluation of library search systems containing any combination of spectral representation and comparison metric hai been presented. While a particular standard of comparison has been used, any desired standard could, in fact, be chosen. It has been demonstrated that this approach can be used to optimize spectral representations or to select comparison metrics. The proposed evaluation process will facilitate the design of LS systems in an objective and quantitative manner and should therefore provide an important step away from the development of such systems by trial and error. Because of the flexibility of this evaluation tool, it will serve well as one of the many techniques which are needed to study the utility of combined spectral information (e.g., mass spectrometry/FTIR) for structure elucidation of chromatographically separated components.
1921)
ACKNOWLEDGMENT The authors wish to thank Jules Abadi for developing the program used to generate the information in Table I. LITERATURE CITED (1) Grotch. S. L. Anal. &em. 1973, 45, 2. (2) Wangen, L. E.; Woodward, W. S.; Isenhour, T. L. Anal. Chem. 1971, 43. - , 1605. . - - -. (3) Woodruff, H. B.;Lowry, S. R.; Ritter, G. L.; Isenhour, T. L. Anal. Chem. 1975, 4 7 , 2027. (4) Penskl, E. C.; Padowskl, D. A.; Bouck, J. B. Anal. Chem. 1974, 4 6 , 955. (5) Kwiatkowskl, J.; Riepe, W. Anal. Chim. Acfa 1979, 112, 219. (6) Rasmussen, G. T.; Isenhour, T. L. J . Chem. I n f . Comput. Sci. 197% 19, 179. (7) Fox, R. C. Anal. Chom. 1976, 48, 717. (8) Heller, S. R.; Koniver, D. A.; Fales, H. M.; Milne, G. W. A. Anal. Chernl. 1974, 46, 947. (9) Zupan, J.; Heller. S. 13.; Milne, G. W. A.; Miller, J. A. Anal. Chlm. Acta 1978, 103, 141. (10) Erley, D. S. Appl. Spectrosc. 1971, 2 5 , 200. (11) Lowry, S. R.; Hyppler, D. A. Anal. Chem. 1981, 53, 889. (12) Kwiatkowskl, J.; Riepe, W. Fresenius' Z . Anal. Chem. 1980, 302, 300. (13) Grotch, S. L. Anal. (?hem. 1870, 42, 1214. (14) Warren, F. V.; Delaniey, M. F. Appl. Spectrosc. 1983, 3 7 , 172. (15) Lam, R. B.; Foulk, S. J.; Isenhour, T. L. Anal. Chem. 1981, 53, 1670. (16) Lam, R. B.; Wleboidl, R. C.; Isenhour, T. L. Anal. Chem. 1981, 5 3 , 889A. (17) Delaney, M. F.; Uden, P. C. Anal. Chem. 1979, 5 1 , 1242. (18) Grotch, S. L. Anal. Chem. 1971, 4 3 , 1362. (19) Grotch, S. L. Anal. Chem. 1974, 4 6 , 526. (20) Grotch, S. L. Anal. Chem. 1975, 4 7 , 1285. (21) Woodruff, H. B.; Lowry, S. R.; Ritter, G. L.; Isenhour, T. L. Anal. Chem. 1975, 4 7 , 2027. (22) Rogers, D. J.; Tanimoto. T. T. Science 1960, 132, 1115.
RECEIVED for review ]December 28,1982. Accepted June l:', 1983. Acknowledgment is gratefully made to the donors of the Petroleum Research Fund, administered by the American Chemical Society, and the National Science Foundation'b Information Science and Chemistry Division (Grant No. IST-8120255) for the financial support of this research.
Enzymatic Determination of Thiamine Pyrophosphate with a pC0, Membrane Electrode Purneshwar Seegopaul and Garry A. Rechnitz*
Department of Chemistry, Uniuersity of Delaware, Newark, Delaware 19711
Thiamine pyrophosphate (ThPP) is determined by potentiometrically measuring the initial rate of carbon dioxide formation from a reactlon sequence involving the recombination of ThPP with pyruvate decarboxylase apoenzyme to the hoioenzyme. The proposed method is hlghiy selective and permtts determination of less than 1 ng mL-' ThPP without any separation procedures or secondary reactions. I n tests of synthetic laboratory samples, the method shows good agreement wlth an enzyme-coupled spectrophotometric procedure.
Thiamine (vitamin BJ exists in blood and tissues both in the free form and as phosphate esters. Thiamine pyrophosphate (diphosphate) or cocarboxylase, ThPP, is the metabolically active coenzyme form of thiamine in a large number of enzymes catalyzing acyl group transfer reactions, for example, decarboxylation of a-keto acids and the formation of a-hydroxy carbonyl linkages (1, 2). In blood, thiamine
pyrophosphate is found mostly in the erythrocytes and, to a lesser extent, in the plasma (3). Clinically, significant reduction in dietary thiamine results in the classical deprivation syndrome, beriberi. Both enzymatic and nonenzymatic methods have been developed to determine thiamine and its phosphate esters. Oxidation of thiamine and its phosphate esters to fluorescent thiochrome derivatives forms the basis for the most widely used chemical assay procedures (4).Separation techniques coupled with fluorometric detection have been proposeld. These techniques include electrophoresis (5),paper chromtatography (6), column chromatography (7), and high-perfornnance liquid chromatography (8,9). Determination of ThPP has also been possibde with a direct current polarographic method (10). Most of these methods suffer from matrix iinterferences, time-consuming procedures, and expensive iinstrumentation. Enzymatic assays for ThPP involve the use of either the transketolase or pyruvate decarboxylase apoenzymes. Re-
0003-2700/83/0355-1929$01.50/0@ 1983 American Chemical Soclety
1930
ANALYTICAL CHEMISTRY, VOL. 55, NO. 12, OCTOBER 19
combination of the inactive apoenzyme with thiamine pyrophosphate gives the holoenzyme whose catalytic activity can then be related to the ThPP concentration, under the appropriate conditions. The transketolase reaction is coupled with a triose isomerase and a-glycerophosphate dehydrogenase-NADH system so that the rate of oxidation of NADH is spectrophotometrically measured and related to ThPP levels (11). Erythrocyte transketolase activity (RBCTK) and the associated T h P P effect give a functional evaluation of vitamin B1 adequacy (12). The pyruvate decarboxylase reaction (eq l) can be manometrically monitored by the carbon dioxide
CH3COCOO- + HzO
pyruvate decarboxylase
CHSCHO + OH-
+ COZ
(1)
liberated or by titration of the released hydroxyl ions (13). It can be coupled to an alcohol dehydrogenaseNADH system (eq 2) so that the rate of oxidation of NADH is proportional to T h P P concentration (14, 15).
CHBCHO + NADH
+ H+
alcohol .b
dehydrogenase
CZH,OH
+ NAD+ (2)
In this paper, we present a potentiometric method for the determination of thiamine pyrophosphate. Carbon dioxide produced in the pyruvate decarboxylase reaction is directly detected with a pC0, membrane electrode. It will be shown that the method is selective, sensitive, and without need for sample pretreamtent or secondary reaction sequences.
EXPERIMENTAL SECTION Apparatus. An Orion Model 95-02 carbon dioxide electrode was used to monitor carbon dioxide produced in the reaction. The potentiometric data were taken with a Corning Model 12 Research pH/mV meter and recorded on a Heath Schlumberger Model SR-255B strip chart recorder operated at a chart speed of 5 mm rnin-l and a range of 100 mV. Enzymatic reactions were performed in a 10-mL jacketed glass cell thermostated at 37 & 0.1 "C with a Haake Model FM constant temperature circulator. Enzyme extracts were separated with an IEC Model HT centrifuge kept at 4 "C to prevent enzyme deactivation. Yeast cells were subjected to ultrasonic processing with a Heat Systems Sonicator Model W220F (20 kHz) in conjunction with a standard (tapered) microtip. Spectroscopic measurements were made with a Hitachi Model 100-60 spectrophotometer. Reagents. Reagent grade chemicals were used without further purification, unless otherwise noted, and all solutions were prepared in deionized water. Saccharomyces cerevisiae (Baker's yeast) YSC-1 Type I, alcohol dehydrogenase (400 units/mg of protein) E.C. 1.1.1.1from yeast, pyruvic acid (sodium salt, Type 11), thiamine hydrochloride (vitamin B1), thiamine monophosphate, thiamine pyrophosphate (ThPP, cocarboxylase), @nicotinamide adenine dinucleotide (reduced form, @-NADH,grade I11 in 2 mg vials), and bovine albumin (fraction V, powder) were M obtained from Sigma Chemical Co. A stock solution of thiamine pyrophosphate was prepared each day by dissolving 4.7 mg in 100 mL of water. The 100- and 1000-fold dilutions of the M, stock solution provided working solutions of lo4 and respectively. A 0.512-mL portion of 0.01 M disodium hydrogen phosphate was added to the 2 mg @-NADHvial to yield a 5 mM solution. Alcohol dehydrogenase enzyme reagent was prepared by dissolving 2 mg in 1.0 mL of 50 mM sodium phosphate buffer, pH 6.8. Pyruvate decarboxylase apoenzyme, 1.3 units/mg total protein was used directly from a solution of ammonium sulfate and disodium hydrogen phosphate obtained in the preparative procedure outlined below. A stock bovine albumin solution of 100 mg mL-l was used to prepare working standards for total protein determinations. Procedures. Preparation of Pyruvate Decarboxylase Apoenzyme, E.C. 4.1.1.1, f r o m Baker's Yeast (16). A 6-g sample of Baker's yeast was suspended in 24 mL of cold 0.1 M sodium phosphate buffer, pH 7.1, for 48 h at 5 "C. The cells were removed
by centrifugation at 15OOOg for 30 rnin and mixed with 12 mL of ice-cold water. The yeast cells were then broken to release the enzyme by ultrasonic processing for 20 min in a Heat systems cold shoulder cooling cell cooled in a stirred salt-ice water bath. Unbroken cells and cell debris were removed by centrifugation at 15OOOg for 30 rnin to give the enzyme protein extract. The cell-freeextract was made 0.4 M with ammonium sulfate and the pH adjusted to 8.6 with 2.0 M cold Tris solution. After 20 rnin of stirring, the suspension was centrifuged at 15OOOg for 25 min. The sediment containing the precipitated phosphates of magnesium and other cations was discarded and solid ammonium sulfate was added to the supernatant to a concentration of 33%. The suspension was thoroughly mixed and the pH adjusted to 8.6 with 0.1 M NaOH in 2.0 M Tris solution. The precipitated proteins were removed by centrifugation at 15OOOg for 30 rnin while the supernatant contained the thiamine pyrophosphate. The protein was again suspended in 0.1 M disodium hydrogen phosphate, pH 6.0. The pH was made 8.6 by 2.0 M Tris and the solution centrifuged at 15000g for 25 min. Solid ammonium sulfate was added to the supernatant to a final concentration of 33% and the pH of the suspension corrected to 8.6 with 0.1 M NaOH in 2.0 M Tris solution. After centrifugation at 15OOOg for 30 min, the precipitated pyruvate decarboxylase apoenzyme extract was suspended in a solution of equal volumes of 0.5 M disodium hydrogen phosphate, pH 6.0, and 25% ammonium sulfate. The enzyme solution was separated into 2-mL aliquots in glass vials and stored frozen. All of the above operations were performed at 0-5 "C. Determination of Pyruvate Decarboxylase Apoenzyme Activity (15). The enzyme was preincubated for total recombination before use. A 2.1-mL sample of 0.1 M sodium phosphate buffer (pH 6.8), 0.4 mL of 0.6 M magnesium sulfate, 0.4 mL of 20 mM thiamine pyrosphosphate, and 0.1 mL of the enzyme solution were mixed and left for 15 min at 25 OC. Following the recombination to give the active holoenzyme, the activity measurements were carried out. A 1.0-mL sample of 0.3 M citrate buffer (pH 6.0), 0.1 mL of 1.0 M pyruvate, 0.1 mL of 5 mM @-NADH,and 1.6 mL of water were mixed in a 3.0-mL quartz cuvette having a 1-cm light path. The solution was brought to 25 "C and 0.1 mL of alcohol dehydrogenase solution added. The reaction was initiated by the addition of 0.1 mL of the preincubated solution of the apoenzyme. The initial rate of absorbance change per minute at 340 nm was recorded and used to compute the activity. The activity units are defined as micromoles of pyruvate decarboxylated per minute at pH 6.0 and 25 "C. The millimolar absorptivity ( 6 ) for @-NADHat 340 nm was taken as 6.02 cm2wmol-l. This assay consisted of a dual enzymatic procedure in which acetaldehyde produced in the pyruvate decarboxylase reaction formed the substrate for alcohol dehydrogenase in the second reaction. The total protein content was determined by the classical biuret method (17) with bovine albumin standards. Subsequently, the activity units per milliliter of solution were related to the protein concentration to yield the specific enzyme activity in activity units per mg of protein. One activity unit of pyruvate decarboxylase (E.C. 4.1.1.1) was defined as 1.0 wmol of pyruvate decarboxylated per minute at pH 6.0 and 25 "C. Spectrophotometric Determination of Thiamine Pyrophosphate. A mixture of 0.1 mL of apoenzyme, 0.1 mL of 0.6 M magnesium sulfate, 0.8 mL of 0.1 M sodium phosphate buffer (pH 6.8), and various aliquots of lo4 or M ThPP was preincubated at 25 "C for 15 min. A 0.1-mL portion of the above mixture was added to a 3-mL quartz cuvette having a 1-cm light path. The following reagents were then placed in the cuvette: 0.1 mL of alcohol dehydrogenase, 1.0 mL of 0.3 M citrate buffer (pH 6.0), 0.1 mL of 5 mM @-NADH,and 1.60 mL of water. The contents of the cuvette were mixed and brought to 25 "C. The reaction was initiated by the addition of 0.1 mL of 1.0 M pyruvate solution. The initial rate of absorbance change per minute at 340 nm was recorded. A calibration curve was prepared by plotting the reaction velocity against ThF'P concentration. The unknown ThPP samples were similarly analyzed and their concentrations directly obtained from this standard curve. Potentiometric Determination of Thiamine Pyrophosphate. A 0.1-mL portion of 0.6 M magnesium sulfate was pipetted into the thermostated glass cell containing a small Teflon-coated
ANALYTICAL CHEMISTRY, VOL. 55, NO. 12,OCTOBER 1983
spinbar for solution mixing. One-tenth milliliter of the apoenzyme reagent was then added to the cell, followed by various aliquots of IO4 M ThPP. The solution was made up to 1.9 mL with 0.1 M citrate buffer of pH 6.!%The contents of the cell were mixed for a few seconds and the pC0, electrode was immersed in the reaction medium. Nixing was continued for 15 min, during which time both a stable electrode base line potential and formation of the holoenzyme were achieved. One-tenth milliliter of 1.0 M pyruvate solution was then added to start the reaction. The carbon dioxide liberated was detected by the pC0, electrode and the initial rates of potential changes, mV min-l, were recorded. Blank determinations in the absence of ThPP were carried out for correction of the rate measurements. Additionally, a fixed-time kinetic approach was taken by measuring potential changes (AE) after 15 min following commencement of the reaction. The initid rates were plotted against corresponding ThPP concentrations to provide a standard calibration curve. Unknown ThPP samples were similarly processed andl their levels computed from the curve.
RESULTS AND DISCUSSION Unfractionated pyruvate decarboxylase apoenzyme prepared from yeast cells was characterized prior to its use as a reagent for ThPP assay vvith the pC02electrode. The kinetic parameters influencing the rate of enzymatic catalysk were optimized, and method validation was established through correlation studies with an enzymatic spectrophotometric method. Studies of Enzyme Preparation. Interaction of excess magnesium(I1) ions and thiamine pyrophosphate with the pyruvate decarboxylase apoenzyme (apo-PDC) resulted in the formation of a stable ternary complex (eq 3) of pyruvate decarboxylase holoenzyme (holo-PdC) The active holoI
ThPP
+
Mg2+
+.
apo-PIX
-
ThPP-Mg
\PDC/
1931
Table I. Time Study of Apoenzyme Stabilitya time (weeks) 1 2 3 4 5 6 7
av % RSD
~
ThPP concentration, ng mL-' 23.0 (taken) 14.0 (found) 23.0 (found) 23.0 22.0 23.4 22.2 23.0 22.5 22.7 ~t 0.51 +2.3%
14.0 (found) 14.4 14.0 13.0 13.4 14.2 14.0 13.9 f 0.49 t3.5%
a Conditions: 23.0 and 14.0 ng mL'' ThPP at 37 "C and pH 6.5 with 50 nnM pyruvate, 30 mM MgZ', and 2.8 units mL-' enzyme.
2.5
E
1.5
K
1.0
t 1
1
,i
/
(3) (holo-PDlC)
enzyme subsequently decarboxylated the substrate, pyruvate, to release carbon dioxide which was monitored by the pC0, electrode. The use of a large excess of Mg2+and apoenzyme made the T h P P Concentration rate limiting, thereby permitting its assay in the method. In the spectrophotometric procedures described, acetaldehyde formed was estimated by the alcohol dehydrogenase-NADH system (eq 1and 2). Due to the large excess of NADH and alcohol dehydrogenase, the decarboxylase reaction foirmed the rate-determining step and the NADH consumption was directly related to the T h P P or decarboxylase concentrations. The enzyme extract obtained in the preparative procedure showed an activity of 56 units mL-l. After total protein determination, the specific activity was calculated as 1.3units mg-' of total protein. The protein concentration was rather high due to the omission of fractionation in the enzyme preparation. This activity proved to be appropriate for sensitive determination of ThPP. The stability of the frozen enzyme was investigated over a period of 7 weeks. Enzyme preparations from a single yeast extract were divided into 2-mL portions of similar activity and stored frozen. A different portion was used once each week to assay samples of 14.0 and 23.0 ng mL-l T h P P at pH 6.5 and 37 OC. The 'ThPP concentration was computed from a standard calibration curve, and the results are given in Table I. Excellent reproducibility was obtained which demonstrated the stability of the unfrnctionated enzyme as an analytical reagent. Standard deviations of f0.51 with an RSD of :k2.3% and *0.49 with an RSD of *3.5% were calculated for T h P P samples of 23.0 and 14.0 ng mL-', respectively. The pyruvate decarboxylase apoenzyme (apo-PDC) utilized in this study consisted of a crude enzyme extract obtained without any fractionation procedures. It was previously reported that such crude pireparations gave enhanced stability and could be kept for many months without any appreciable
Enzyme
A c t i v i t y , Units/rnL
Flgure 1. Effect of enzyme actlvity on the rate of decarboxylation of 50 mM pyruvate at pH 6.5and 37 O C , using 30 mM Mg2+ and 23.0 ng mL-' ThPP.
deterioration in activity (16). On the contrary, purified apo-PDC exhibited poor stability evidenced by significant loris in activity within a few days (15). This lack of stability introduces a serious disadvantage in methods involving the use of purified apo-PDC. During the course of this work, it was found that preparation of apoenzyme from purified enzyme by gel filtration exhibited poor stability and rapid loss in activity. Recently, it was shown that the low stability of purified pyruvate decarboxylase made its use impractical for the construction of a pyruvate enzyme electrode compared to the superior performance of a tissue-based pyruvate enzyme electrode involving intact enzyme (18). It is possible, therefore, that the absence of fractionation allowed the enzyme to remain in a stable environment enhanced by the presence of other proteins. Effect of Enzyme Activity and Preincubation. Figure 1 shows the effect of enzyme activity on the T h P P assaly method with measurements made a t 37 "C and pH 6.5. With a sample of 23.0 ng mL-l ThPP, the initial rate increased linearly up to 2.5 units mL-l enzyme, followed by small increases at higher concentrations. An activity of 2.8 units mL-l apoenzyme was employed throughout the remainder of the work to ensure rate independence from enzyme concentratioin. Reconstitution of the holo-PDC is a time-dependent procem and studies were performed to establish the preincubation time necessary for optimum activity in this method. By use
1932
ANALYTICAL CHEMISTRY, VOL. 55, NO. 12,OCTOBER 1983
Table 11. Effect of Incubation Time on the Rate of Decarboxylation at pH 6.5 and 37 '(2, with a Sample of 23.0 ng mL-' ThPP with 50 mM Pyruvate, 3 0 mM Mg2+, and 2.8 Units mL" Apoenzyme incubation time, min
% activity
2.5 8.0
36 79
15
100
30 40 60
94 88 72
2.5
1
Magnesium(l1) Concentration,mM Figure 3. Effect of Mg2+ concentration on enzyme activity at pH 6.5
and 37 O C , using 23.0ng mL-' ThPP, 50 mM pyruvate, and 2.8 units mL-I enzyme.
0
20
40
60
80
100
P y r u v a t e Concentration, m M
Figure 2. Effect of substrate concentration on the reaction rate at pH 6.5 and 37 O C , with 23.0 ng mL-' ThPP and 2.8 unlts mL-' apoenzyme.
of a sample of 23.0 ng mL-l ThPP, the initial rates of decarboxylation were measured in separate experiments involving different preincubation periods for holoenzyme formation. Table I1 summarizes the percent activity obtained at 37 "C for the periods ranging from 2.5 to 60 min. Incubation times over 15-30 min gave maximum activity for ThPP assays. Longer preincubation periods resulted in decreases in the reaction rates probably due to partial denaturation of the enzyme. Effects of Pyruvate and Magnesium(I1). Studies on the effect of pyruvate concentration on enzymatic activity revealed an optimum substrate concentration of 60 mM or higher, as shown in Figure 2. A final concentration of 50 mM used in this work allowed rate independence from any substrate effect, thereby permitting the measurement of ThPP. Higher levels of pyruvate caused rather large blanks. Analysis of these results by the Lineweaver-Burk reciprocal method gave a KM constant of 24 mM. KMvalues ranging from 1to 30 mM were previously reported for the enzyme and pyruvate substrate (19, 20). Figure 3 shows the influence of magnesium(I1) ions on the rate of decarboxylation at pH 6.5 and 37 O C . A concentration of 30 mM or greater produced maximum enzymatic activity; thus, a concentration of 30 mM Mg2+was used in all experiments. The rather high concentration of Mg2+needed for optimum catalytic activity can be attributed to the use of an unfractionated enzyme extract, with possible binding of ions to other proteins. Effect of Temperature and pH. The influence of temperature on the sensitivity of the ThPP assay was studied over the range 20-45 "C by using a sample of 23.0 ng mL-l T h P P in the 2-mL reaction volume. Figure 4 shows an optimum
I
20
25
30
35
40
45
Temperature, OC Figure 4. Dependence of the reaction rate on temperature at pH 6.5 with 23.0 ng mL-' ThPP, 2.8 units mL-' enzyme, 30 mM Mg2+, and 50
mM pyruvate.
temperature at 37 "C with a linear increase in reaction rate from 24 to 35 O C . At temperatures higher than 37 O C , the rate decreased due to enzyme denaturation. All experiments were therefore carried out at 37 O C . The incubation time of 15 min is directly related to the temperature effect because a lengthy incubation period leads to enzyme protein denaturation at high temperatures. The p H profile of the decarboxylase catalyzed reaction is shown in Figure 5. At 37 "C,studies over the pH range of 5.4-7.2 gave an optimum pH of 6.4-6.5 and a pH of 6.5 was used in all experiments. pH optimum values of 5.4-6.0 were previously reported (19). In this method, however, both the enzyme recombination process and the decarboxylation reaction were performed in the same reaction medium. Both processes show different pH optima, with the holoenzyme formation needing maximum pH values of about 6.8 (16).The pH optimum obtained in this study, therefore, reflected a compromise between these two reactions and the nature of the detection technique. Additionally, the small increase in
ANALYTICAL CHEMISTRY, VOL. 55, NO. 12, OCTOBER 1 9 8 3
L l ~ l - - . l - U 54
5.8
6.2
66 PH
7-0
7.4
Figure 5. pH profile of the enzyme recombination and catalytic aci at 3 7 'C, using 50 mM pyruvate, 30 mM Mg2+, 23.0 ng mL-' Tt and 2.8 units mL-' enzyme.
-
Table 111. Precision and Relative Errors Obtained from Random Sampling of ThPP at pH 6 . 5 and 37 "C, with 50 mM Pyruvate, 30 mM Mg2+,and 2.8 Units mL-' Enzyme thiamine concn, ng mL-' --___-
re1
taken
found
error, %
RSD, %
1.20 4.60 9.20 14.0 23.0 35.0
1.25 4.6'7 9.110 14.3 22.6 35.5
t 4.2
t4.0 k 2.1 t 1.0 k0.7 t0.8 t1.9
t 1.5
-1.1 t 2.1
-1.7 t 1.4
reaction rate between pH 5.6 to 5.8 has been attributed to a pH-independent activity region noted earlier (19). Precision and Accuracy Studies. The precision and accuracy of the potentiometric method were investigated by the random analysis of Eieveral T h P P samples within the concentration range of l.f!0-35.0 ng mL-l. Each sample was analyzed six times under optimum conditions and the results were compared to a standard calibration curve to obtain their concentrations. The percentage RSD was calculated to give the precision while the mean concentration was compared to the actual concentration of T h P P to give the percentage relative error. Table I11 shows an error range of -1.1 to +4.2% and precision of k0.7 to d:4.0% for the concentration range studied. Selectivity Studies and Comparison with Spectrophotometric Methods. The selectivity of the method was examined by substituting other thiamine compounds for ThPP in the assay procedure. A 100-fold excess of thiamine or thiamine monophosphate failed to produce any noticeable interference. The method is specific for this form of thiamine, that is, thiamine pyrophosphate, since the other thiamine compounds cannot function as the coenzyme necessary for formation of the holoenzyme.
1933
Aqueous synthetic laboratory samples of ThPP were ana. lyzed by both the potentiometric and the spectrophotometric methods described earlier. The samples were freshly made and analyzed on the same day. Statistical evaluation of the results yielded a correlation coefficient of 0.99 with a slope of 0.99 and a y intercept of 0.15. These data indicated good agreement between the two methods over the concentration range of 1.20-35.0 ng mL-l ThPP. Determination of Thiamine Pyrophosphate. With the optimum conditions experimentally established, the calibration plot of ThPP concentration vs. reaction rate gave a linew range of up to 30.0 ng mL-l ThPP. Least-squares treatmenit of the results revealed a slope of 0.08 while the correlation coefficient of 0.99 showed that the data approximated a straight line. The method allowed a detection limit of less than 1.0 ng mL-l ThPP, which compared favorable with otheir methods. This value is much lower than the normal concentration range of 3-1 1pg/lOO mL of ThPP present in blood (3). The potentiometric enzymatic method, therefore, offers a simple and sensitive assay procedure for the determination of ThPP. Enzymatic specificity is coupled with the inherent selective features of the membrane electrode which directly monitors the reaction rate without any secondary reactions. Interferences from ionic or optically absorbing species are effectively eliminated in the potentiometric method. These advantages should make the method attractive for the routine analytical determinatiion of ThPP in different kinds of samples. Registry No. Thiamin pyrophosphate, 154-87-0; pyruvate decarboxylase, 9001-04 -1.
LITERATURE CITED (1) Bohinski, Robert C. "Modern Concepts in Biochemistry", 3rd ed.; Aiiyn and Bacon: Boston, MA, 1979; p 407. (2) Mlckeisen, 0.; Yamamoto, R. S. I n "Methods of Biochemicaii Analysis"; Giick, D., Ed.; Interscience: New York, 1958; Vol. 6, 19 1-257. (3) Henry, Richard, J. "Clinical Chemistry: Principles and Techniques"; Harper and Row: New York, 1965; p 720. (4) Penttinen, Hannu, K. Methods Enzymol. 1979, 6 2 , 58-59. (5) Penttinen, Hannu, K. .4cta Chem. Scand., Ser. B 1978, 32, 609-612. (6) Lewin, Lawrence, M.; Robert, W. Anal. Biochem. 1066, 16, 29-35. (7) Matsuda, T.; Cooper, Jack, R. Anal. Biochem. 1981, 117, 203-207. (8) Sanemori, H.; Ueki, H.; Kawasaki, T. Anal. Biochem. 1980, 107, 451-455. (9) Kimura, M.; FuJita.T.; Nishida, S.; Itokawa, Y. J . Chromatogr. 19801, 188, 417-419. (10) Vergara, T.; Marin, D.; Vera, J. Anal. Chim. Acta 1980, 120, 347-35 I . (11) Gubier, C. J.; Johnson, L. R.; Wittorf, J . H. Methods Enzymol. 19701, 18A, 120-125. (12) Brin, M. Methods Enzymol. 1970, f8A1 125-133. (13) Scheiienberger, A.; Hubner, G.; Lehman, H. Angew. Chem., I n t . Ed. Eng/. 1 9 6 8 ~7 , 886-887. (14) Scheiienberger, A. Angew. Chem., Int. Ed. Engl. 1967, 6 , 1024-1035. (15) Uiirich, J. Methods Enrymol. 1970, 18A, 109-115. (16) Morey, A. V.; Juni, E. J . Blol. Chem. 1968, 243, 3009-3019. (17) Kachmar, John F. In1 "Fundamentals of Clinical Chemlstry"; Tietz, NI. W., Ed.; W. 8. Saunciers: Philadelphia, PA, 1970; pp 188-191. (18) Kuriyama, S.;Arnold. M. A.; Rechnitz, G. A. J . Membr. Sci. 1983, 12, 269-278. (19) Jordan, F.; Kuo, D. J.; Monse, E. V. Anal. Biochem. 1978, 86, 298-302. (20) Barman, T. E. "Enzyme Handbook"; Springer-Veriag: New York, 1969; Voi. 11, p 701.
RECEIVED for review March 31, 1983. Accepted July 8, 1983. We gratefully acknowledge the support of the National Institutes of Health, Grant GM-25308.
