substrate specificity and ph dependence of homogeneous ... · homogeneous wheat germ acid...

ARCHIVES OF BIOCHEMISTRY AND BIOPHYSICS

Vol. 288, No. 2, August 1, pp. 634-645, 1991

Substrate Specificity and pH Dependence of Homogeneous Wheat Germ Acid Phosphatase

Robert L. Van Etten’ and Parvin P. Waymack Department of Chemistry, Purdue University, West Lafayette, Indiana 47907-1393

Received November 27, 1990, and in revised form April 17, 1991

The broad substrate specificity of a homogeneous isoenzyme of wheat germ acid phosphatase (WGAP) was extensively investigated by chromatographic, electrophoretic, NMR, and kinetic procedures. WGAP exhibited no divalent metal ion requirement and was unaffected upon incubation with EDTA or o-phenanthroline. A comparison of two catalytically homogeneous isoenzymes revealed little difference in substrate specificity. The specificity of WGAP was established by determining the Michaelis constants for a wide variety of substrates. p- Nitrophenyl phosphate, pyrophosphate, tripolyphosphate, and ATP were preferred substrates while lesser activities were seen toward sugar phosphates, trimetaphosphate, phosphoproteins, and (much less) phosphodiesters. An extensive table of K,,, and V,,, values is given. The pathway for the hydrolysis of trimetaphosphate was examined by calorimetric and “I’ NMR meth- ods and it was found that linear tripolyphosphate is not a free intermediate in the enzymatic reaction. In contrast to literature reports, homogeneous wheat germ acid phosphatase exhibits no measureable carboxylesterase activity, nor does it hydrolyze phenyl phosphonothioate esters or phytic acid at significant rates. o 1991 Academic

Press, Inc.

The biological roles of plant and animal acid phosphatases remains largely unknown. Substrate specificity studies can provide important, even essential clues for elucidating these roles. In the case of wheat germ acid phosphatase, such studies appeared particularly important not only to obtain insights about the biological role of the enzyme, but also to test some provocative proposals that had been presented earlier. Reportedly, wheat germ acid phosphatase contained three isoenzymes that were separable by DEAE-cellulose chromatography, and phosphatase and carboxylesterase activities were coincident

’ To whom correspondence should be addressed.

634

in all three chromatographic peaks (1). It thus seemed possible that a hypothesis of Dorn might apply to these wheat germ phosphatase and esterase activities (2). This hypothesis, which was derived from genetic studies, was based on the presumed multiple-polypeptide nature of acid and alkaline phosphatases, and proposed that some of the polypeptide chains were common to these two enzymes. Therefore, a multiple polypeptide structure containing esterase and phosphatase polypeptide chains represented a possible explanation for the dual activities. This was thought preferable to an explanation based upon a dual activity of one catalytic site. Related studies in which fractions that were eluted from preparative polyacrylamide gels exhibited both phosphatase and carboxylesterase activity seemed to support this hypothesis (3). This remarkable possibility deserved examination.

Previous studies of the specificity of wheat germ acid phosphatase were carried out using enzyme preparations of low specific activity and at a single substrate concentration (1, 4, 5). It was not convincingly demonstrated that the presence of activities toward a wide variety of phosphate and even carboxylic acid esters resulted from a single enzyme species. Consequently, a careful study of the substrate specificity of a homogeneous enzyme was necessary in order to test the hypothesis that the extremely broad specificity which has been reported for the enzyme is due to an individual protein rather than to the presence (as impurities) of entirely different enzymes.

It was also necessary to establish if wheat germ acid phosphatase possessed a number of phosphatase activities. Broad and overlapping specificities have been suggested to be characteristic of plant acid phosphatase isoenzymes, although this view too was based largely on studies with partially purified preparations (6). However, homogeneous sunflower seed and soybean acid phosphatases do hydrolyze a wide range of substrates, although the Michaelis constants of only a few substrates have been measured (‘7,8). Furthermore, other plant phosphatases may exhibit a broad specificity and may hydrolyze many of the same substrates as acid phosphatases. For

0003.9861/91 $3.00 Copyright 8 1991 by Academic Press, Inc.

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WHEAT GERM ACID PHOSPHATASE SUBSTRATE SPECIFICITY 635

example, an acid phosphodiesterase from tobacco leaves reportedly exhibits activity toward p-nitrophenyl phosphate, pyrophosphate, and ATP (9).

The natural role of wheat germ acid phosphatase is presumably related to some aspect of seed germination. In ungerminated seeds, the bulk of total phosphorus (storage phosphate) appears in phytin (myoinositol hexaphosphate). It seemed important to investigate phytin as a possible natural substrate for acid phosphatase isoenzymes since the enzyme might have a role in phytin mobilization. The phosphate competitive inhibition constant is sufficiently large that it might allow wheat germ acid phosphatase to be an effective hydrolytic agent despite the elevated phosphate levels that may be present during germination. However, although a phytase from wheat bran possesses a pH optimum similar to that of acid phosphatase, and also hydrolyzes cu-naphthyl phosphate (lo), careful studies of soybean acid phosphatase and phytase have demonstrated distinct enzymes (8,ll).

In the present study, the fundamental question of specificity was first approached by the isolation of a homogeneous isoenzyme, here termed WGAP2 (12). In this way, questions about the hydrolysis of a wide variety of phosphomonoesters, phosphodiesters, and polyphosphates by a single enzyme species could be approached by determining the coincidence of these activities during chromatography, after polyacrylamide gel electrophoresis and as a function of purity. The examination was then ex- tended to include accurate measurements of the Michaelis kinetic constants for a wide range of substrates. Deter- mination of the Michaelis constants for different substrates provides information of significance in interpreting the potential biological role of the enzyme. In addition, the behavior of V,,,,, often provides information about the mechanism of action of such enzymes. Finally, the pH dependence of catalysis was examined by determining the pH variation of the Michaelis constants for both p-nitrophenyl phosphate and P-glyceryl phosphate.

EXPERIMENTAL PROCEDURES

Phosphatase activity was measured at 25°C and pH 4.6 using 5 mM

p-nitrophenyl phosphate as a substrate. Substrate solution (2 ml) contained 0.11 M sodium acetate buffer and 0.01% Triton X-100. Enzyme dilutions were made to give a final absorbance reading of 0.3 to 1.0 after quenching the reaction with 1 ml of 1 N NaOH. Carboxylesterase activity was measured using 1.0 mM p-nitrophenyl acetate as a substrate in 3.00 ml of 0.11 M, pH 4.6, acetate buffer. The increase in absorbance at 325 nm (f = 7.7 X lOa Mm’ cm’) was monitored directly using a Gilford 2000 spectrophotometer. After electrophoresis at pH 4.4 (13), carboxylesterase activity staining was carried out using a-naphthyl acetate as a substrate (14).

Phytase activity was measured with assay mixture containing 1.0 mM

sodium phytate (myoinositol hexaphosphate, Na,,C,H,O,,P,. 33H20; Sigma grade V) in pH 5.0 0.1 M acetate buffer. After the addition of 50 aI of enzyme solution to 2.0 ml of the substrate solution, the reaction

’ Abbreviations used: WGAP, a homogeneous isoenzyme of wheat germ acid phosphatase.

was incubated for various times at 37°C. The reaction was quenched by the addition of 0.50 ml 3% ammonium molybdate in 0.5 M pH 4.0 acetate buffer followed by the addition of 0.20 ml 1% ascorbic acid. The absorbance at 700 nm was read after 30-45 min.

Polyacrylamide gel electrophoresis was carried out at pH 4.4 and pH 8.0 (13). After electrophoresis the gels were incubated in 5 mM substrate solution made up in 0.1 M acetate buffer at pH 5.0. After a time sufficient to allow sufficient inorganic phosphate release, the incubation mixture was replaced by the developing reagent which quenched the enzymatic reaction (15). The white or light yellow precipitate which formed indicated the position of enzymatically produced phosphate and was stable to several days of storage in 7% acetic acid. In a typical experiment, l- 5 units of enzyme were applied to a gel. After electrophoresis, incubation of the gel with the substrate solution for 2-30 min was followed by treatment with the Sugino reagent for approximately 5 min.

Phosphate assays were conducted in two ways, depending on the sensitivity required. In one (16), the basis for phosphate quantitation is the absorbance of the reduced form of phosphomolybdic acid which is formed with ascorbic acid at pH 4 (where the nonenzymatic hydrolysis of labile phosphate esters is usually minimized). The procedure was slightly modified. In a typical analysis, 2.0 ml of a substrate solution was incubated with enzyme and the reaction quenched by the addition of 0.50 ml of 1.5% ammonium molybdate in 2.79 MpH 4.0 acetate buffer. Reduction to the chromophore was achieved by the addition of 0.20 ml of 1% ascorbic acid and after full color development (15-60 min), the absorbance was read at 700 nm (t = 4200 M-i cm-‘). Incorporated into this procedure are the following modifications of the original procedure (16): (a) The use of trichloroacetic acid as an activity quench (which contributes to nonenzymatic hydrolysis) is avoided since the addition of the molybdate reagent (a potent competitive inhibitor) stops the enzymatic reaction. (b) The 2.79 M pH 4.0 buffer has sufficient buffer capacity to bring the mixture to the prerequisite pH 4.0 under most assay conditions. High pH buffers and concentrated substrate solutions require compensation by lowering the pH of this buffer to give pH 4.0 in the final mixture. (c) The molybdate concentration of 1.5% was found sufficient for up to 5 mM phosphate esters which did not contain significant inorganic phosphate contaminant and for up to 1 mM pyrophosphate or ATP. Higher concentrations of substrate required larger volumes or higher concentrations (up to 6%) of ammonium molybdate reagent. (d) The total volume of the final mixture for which the absorbance was determined was usually 2.75 or 3.25 ml (1.00 ml molybdate added), compared to 20 ml for the original procedure. This gives a significant improvement in sensitivity by avoiding excessive dilution. The effective molar extinction coefficient was defined as molar extinction of product X (sample volume)/(reading volume) in order to evaluate modifications and for comparison to standard procedures. The effective molar extinction coefficients of this modified assay and the assay described in the following section were 3050 and 14,600 Me’ cm-‘, respectively.

An alternative method was developed based on the fact that an examination of the absorbance spectra of the previous pH 4 assay products revealed the presence of an intense absorbance peak at 325 nm. This wavelength was found to be reliable in the determination of inorganic phosphate if a high energy monochromator was employed in the constant slit mode. The procedure was the same as that described in the previous section except that 0.1% rather than 1% ascorbic acid was added. The lower ascorbic acid concentration required a longer time for development of 325 nm absorbance but the uv absorbance of 1% ascorbic acid created unmanageable absorbance in the blank. The product formed was linear with concentration and gave a molar extinction coefficient of 2.01 X 10” Mm’ cm-‘. The time required for maximum 325 nm absorbance was long- est at the highest substrate concentration employed, 5 mM. The standard procedure was to add a known amount of phosphate standard to the highest substrate concentration employed and follow the absorbance with time in order to determine how much time was required to reach maximum absorbance. A limitation of the 325.nm method is that it cannot be used with aromatic substrates that absorb at 325 nm, or with substrates where a higher concentration of molybdate reagent is required (e.g., 1 mM pyrophosphate).

636 VAN ETTEN AND WAYMACK

Several substrate specificity studies were carried out in conjunction with protein preparations at different stages of purification. A single activity peak was obtained upon G-75 Sephadex chromatography (12). The coincidence of this peak with that of catalytic activity toward various substrates is a necessary indication that WGAP alone is acting upon the substrates. The elution profile from a 5.0 X 90 cm G-75 Sephadex column was assayed for activity toward p-nitrophenyl phosphate, pyrophosphate, 5’-ATP, 5’-AMP, and glucose 6phosphatase, at pH 4.6, ATPase activity with 1.0 mM ATP was determined by the novel Lowry modification which measures 325 nm absorbance. The activities toward 5.0 mM sodium salts of pyrophosphate and glucose 6-phosphate were measured as 700 nm absorbance by the standard Lowry type assay. Activity toward 20 mM 5’-adenosine monophosphate was also measured by this Lowry assay with the concentration of the molybdate reagent increased to 3%. The elution profile from a SP-Sephadex ion exchange column was similarly assayed for acid phosphatase activity. The sample was a 50-60% ammonium sulfate fraction. The activities toward nitro- phenylphosphate, pyrophosphate, ATP, and glucose 6-phosphate were assayed essentially as described above. Phosphate ion competitive inhibition constants were measured using both p-nitrophenyl phosphate and glucose 6-phosphate as substrates. The phosphate which was released was measured by a Lowry type assay (with 6% molybdate reagent) mea- suring 700 nm absorbance, using the constant slit mode of the Gilford 2000 spectrophotometer to suppress the large blank absorbances.

Diesterase activity toward bis-p-nitrophenyl phosphate was found in purified fractions, although it was initially at such low levels that the presence of a p-nitrophenyl phosphate contaminant in bis-p-nitrophenyl phosphate preparations or a diesterase contaminant in the enzyme needed to be considered. Since bis-p-nitrophenyl phosphate was found to be a better substrate at pH 8.5 than at pH 5, and this seemed to indicate a contaminatingphosphodiesterase, the assay of an ion exchange column eluate was undertaken to determine if the phosphodiesterase and phosphomonoesterase activities were coincident. A sample containing several wheat germ acid phosphatase isoenzymes was applied to a SP-Sephadex column at pH 5.2. The sample was a methanol precipitate fraction (12) with a specific activity of about 8 U/mg. The low specific activity and mixture of isoenzymes in this sample made it a good test for detecting a phosphodiesterase contaminant that was not coincident with WGAP isoenzymes. Activity toward bis-p-nitrophenyl phosphate was detected by the addition of aliquots from each column fraction to 2.20 ml of 10 mM substrate in 0.05 M pH 8.5 barbital buffer. The increase in 400 nm absorbance was followed directly at 25°C on a Beckman ACTA V spectrophotometer. The activity toward p-nitrophenyl phenylphosphonate was measured by a similar assay with 10 mM substrate.

In order to estimate the Michaelis constants of other substrates having widely varied V,,, and K, values, it was necessary to employ several inorganic phosphate assays and assay modifications. The substrate concentrations (minimum of five) used in these studies were chosen to avoid substrate inhibition in some cases and to enable measurement of turnover under initial velocity conditions. The data was evaluated graphically to detect substrate inhibition or nonlinearity and the Michaelis constants were evaluated by computer fitting to the rectangular hyperbola (17). Substrate solutions were prepared in 0.1 M pH 5.0 acetate buffer and assays were carried out at 25.0 + O.l”C. The constants forp-nitrophenyl phosphate were determined as previously described. The Michaelis constants of the substrates phenyl phosphate andp-chlorophenyl phosphate were determined in 0.1 M pH 4.6 acetate buffer by a similar method.

The V,,, and K,,, of bis-p-nitrophenyl phosphate, thymidine-5’-phosphate p-nitrophenyl ester and p-nitrophenyl phenylphosphonate were determined at pH 7.0 in 3,3dimethylglutarate buffer and at pH 8.5 in barbital buffer. With p-nitrophenyl phenylphosphonate at pH 5.0, a 2.00-ml incubation mixture with 1.00 ml of 1 M Na2C08 quench was employed. The stoichiometry of inorganic phosphate andp-nitrophenol release at pH 5.0 and pH 8.5 was determined with 10 mM bis-p-nitrophenyl phosphate as a substrate. After less than 2% total hydrolysis, the 5.0-ml reaction mixture was quenched with 0.5 ml of 3% ammonium

molybdate in 0.5 M, pH 4.0, acetate. The 400-nm absorbance of 2.00-ml portions of the quenched mixture was determined after the addition of 1.00 ml of 1 M NazCO,. The inorganic phosphate content of identical 2.00-ml portions was determined by the Lowry assay.

Stock solutions of p-nitrophenyl phenylphosphonate (25 mM) were prepared in pH 8.50 0.05 M sodium barbital buffer with 0.1 M NaCl added and the pH was adjusted with a small amount of concentrated HCl. The lower substrate concentrations were made by diluting stock substrate with the buffer. Enzyme was added to 2.40-ml portions of the substrate at 25.O”C and the absorbance at 400 nm recorded directly using a full-scale absorbance of 0.1 or 0.2 on a Beckman Acta V spectrophotometer. The rate was measured from the initial linear part of the recorder plot. The substrate range 0.5-20.0 mM was investigated.

Studies of the hydrolysis of trimetaphosphate hydrolysis using srP NMR were conducted with 20 mM trimetaphosphate in a 0.05 M pH 4.0 acetate buffer in 90% DzO. Solutions (20 mM) of pyrophosphate and tripolyphosphate were prepared in 0.05 M pH 4.0 acetate buffer in 90% D20. 31P NMR spectra were obtained with a Varian XL-100 NMR spec- trometer operating at 40.507 MHz for 31P, locked on the *H resonance of the solvent (90% DzO). Chemical shifts were measured relative to 85% phosphoric acid at the probe ambient temperature of 25-30°C.

The phosphoproteins ovalbumin and phosvitin were also tested as substrates for WGAP. Ovalbumin (5 mg/ml) and phosvitin (1 mg/ml) were incubated with WGAP at 25°C in 0.1 M pH 5.0 acetate buffer. The reaction was quenched with an equal volume of 20% trichloracetic acid containing 10% ascorbic acid and 5% sodium lauryl sulfate to prevent protein precipitation. Then 0.20 ml of 6.25% ammonium molybdate and 0.50 ml of 2% acetic acid containing 6% sodium citrate and 2% sodium arsenite were added to each 1.00 ml of the quenched mixture (18). After 30 min the absorbance was determined at 700 nm versus an identical blank mode with enzyme added after the quench.

The effect of WGAP treatment on ovalbumin was also examined by polyacrylamide gel electrophoresis. A 2.0 mg/ml ovalbumin sample in pH 5.4 acetate buffer was divided into two portions. After overnight treatment of one of the samples with 1.5 units of WGAP, the two samples were subjected to electrophoresis at pH 8.0. Samples of untreated ovalbumin, treated ovalbumin, and a mixture were stained for protein and the electrophoretic patterns compared.

The pH dependence of the Michaelis constants was determined at 25°C over the pH range 2.59-9.20 in 0.05 M buffer with added 0.1 M NaCl. Below pH 7, a discontinuous assay was employed with enzyme added to a substrate volume of 2.00 ml which was thermostated at 25’C. The reaction was quenched with either 1.00 or 0.40 ml of 1.0 M NaOH and the absorbance at 400 nm was measured. Usually, seven substrate concentrations (minimum of five) were employed and the maximum extent of hydrolysis at the lowest substrate concentration employed in the calculations was 8%. In a few cases, a combination of small K,,, values (-0.1 mM) and substrate inhibition at low pH limited the range of substrate concentrations that could be employed to values of Km and above, with the upper limit being set by substrate inhibition.

At pH 7.0 and above the rates were determined by adding enzyme to 2.40 ml of substrate solution and the absorbance versus time was followed at 400 nm. All absorbance measurements were corrected for p-nitro- phenolate ionization using pK,, = 7.00 for p-nitrophenol. The slope of the initial linear portion of the curve was measured. Correction was made for the inactivation during assay at pH 8 and above. The values of the Michaelis-Menten parameters and the associated error limits were calculated by the use of the program HYPER (17).

RESULTS AND DISCUSSION

Crude samples of wheat germ acid phosphatase were subjected to gel electrophoresis followed by phosphatase and carboxylesterase activity-staining. The gels showed multiple bands of esterase and phosphatase activity with some of them clearly resolved and some of them appar-


ently coincident. Electrophoretic and chromatographic results indicated that most of the carboxylesterase activity in wheat germ was due to esterases that were of lower molecular weight than acid phosphatase. The elution profile of a 2.5 X 90 cm column of Sephadex G-75 showed three peaks of carboxylesterase activity. Two large overlapping peaks accounted for about 95% of the esterase activity and had molecular weights lower than acid phosphatase. A single phosphatase activity peak which had an elution volume similar to that of the third small carboxylesterase activity peak was found. Some of the carboxylesterase isoenzymes were electrophoretically similar in behavior to some of the phosphomonoesterase isoenzymes. However, as WGAP was purified, the increasing ratio of phosphatase to carboxylesterase-specific activities quickly showed that the two activities were due to separate enzymes. Enzyme fractions from various stages of purification (12) were assayed for phosphatase and esterase activity. The phosphatase to carboxylesterase activity ratio steadily increased with increasing phosphatase specific activity (Table I). Esterase activity became undetectable in the very highly purified samples. Activity staining for esterase with enzyme having a (phosphatase) specific activity of 230 U/mg revealed no esterase activity band. These results clearly establish that the report of concomitant carboxylesterase and phosphatase activities in wheat germ phosphatase proteins (1) was erroneous. The present electrophoretic studies suggest that the large number of esterase activities present in wheat germ led earlier work- ers to the incorrect assumption of dual activities. The esterase activity with a molecular weight similar to that of WGAP which was observed in the G-75 Sephadex elution profile is probably the same wheat germ esterase which was reported to have a molecular weight of 51,000 (18). The separation of these activities, together with the finding that WGAP contains a single polypeptide chain

(la), indicates that the original hypothesis of Dorn (2) is not applicable here.

Studies of phytase activity showed that the relatively low phytase activity in the initial extract decreased sig- nificantly as the acid phosphatase was purified (Table I). The homogeneous acid phosphatase isoenzyme WGAP has no significant activity toward phytic acid. This conclusion is consistent with recent studies on soybean phosphatase and phytase (8, ll), and suggests that a reex- amination of contrasting results with rice ear, apricot kernel, and rice bran acid phosphatases is in order (6,20, 21). Conceivably, a single phytase can catalyze the se- quential breakdown from hexaphosphate to monophosphate or inositol. In one case, at least 12 phosphate-containing intermediates have been detected (22). Although WGAP is not a primary phytase, it could conceivably be active in a cooperative role involving some of the large number of possible intermediate hydrolysis products. The diversity and practical unavailability of the individual potential intermediates precluded their being tested as substrates.

Both the partially purified enzyme and a homogeneous isoenzyme exhibited catalytic activity toward a wide variety of substrates. The activity profile obtained upon elution of partially purified enzyme from an SP-Sephadex column (step 7 of Ref. (12)) shows that for each substrate there are two activity peaks, corresponding to the elution position of two isoenzymes of wheat germ acid phosphatase (Fig. 1). The activity ratios measured in these experiments indicate no significant differences in the specificities of the two isoenzymes. The activity profiles obtained with the phosphodiester bis-p-nitrophenyl phosphate and with p-nitrophenyl phenylphosphonate (data not shown) are also consistent with the hydrolysis of all of the substrates by the two acid phosphatase isoenzymes.

TABLE I

Acid Phosphatase, Carboxylesterase, and Phytase Activities in Wheat Germ Acid Phosphatase Preparations

Phosphatase activity” (firno min-’ mg-‘)

0.46 1.8

33 93

230 451

0.078 (initial extract) 27 70

308

Carboxylesterase activity* Ratio of phosphatase (pm01 mini mg-‘) to carboxylesterase

0.0147 31 0.0196 91 0.0557 590 0.00761 1.22 x lo4

d >107 d >lol

Phytase activity’ (pm01 mini rng-‘)

0.013 1.13 0.64 0.37

Ratio of phosphatase to phytase

6 24

109 832

D Measured using p-nitrophenyl phosphate as substrate at pH 4.6. * Measured usingp-nitrophenyl acetate as substrate at pH 4.6. ’ Measured using sodium phytate at pH 5.0. d Not distinguishable from nonenzymatic hydrolysis.


20

t

5 P-PI 0 PNPP l ATP ‘3 Glucose-6-P

15 oa-

c - 5 = 0.6 F IO 5

2 F 0 04- u

0.5 0.2

0 i

FRACTION NUMBER

FIG. 1. Multiple phosphate ester assay of a mixture of isoenzymes during SP-Sephadex chromatography. The assay procedures described in the text were used to assay for activity toward inorganic pyrophosphate (0); p-nitrophenyl phosphate (Cl); 5’.adenosine triphosphate (0); and glucose 6-phosphate (a). The activity units for each substrate were normalized for clarity in plotting.

The coincidence of catalytic activities toward numerous phosphate ester substrates during gel filtration chromatography is shown in Fig. 2. The position of the maximum activity was identical with every substrate and only a single peak was found for each substrate. Thus, if several enzymes are responsible for the activities, they would all have to possess identical elution volumes in this G-75 elution experiment. Purification of one isoenzyme (WGAP) to homogeneity (12) made it possible to perform experiments in which a single, highly purified isoenzyme was involved. When an SP-Sephadex chromatographic elution profile was assayed (data not shown), the ratios of p-nitrophenyl phosphatase, ATPase, and pyrophosphatase activities were constant across the activity peak, indicating that a single enzyme species is catalyzing the hydrolysis of all three substrates. Thus, WGAP possesses pyrophosphatase and ATPase activities in addition to phosphomonoesterase activity.

Heat treatment was used to partially inactivate WGAP and the activity toward p-nitrophenyl phosphate, bis-p- nitrophenyl phosphate, and p-nitrophenyl phenylphosphonate was measured in order to determine if the activities were separable. The results (Fig. 3) are consistent with common activity toward these substrates. The value of the heat denaturation test is emphasized by the results of a similar study of an apparently homogeneous fungal acid phosphatase. Although that preparation possessed almost equal activity toward p-nitrophenyl and bis-p-nitrophenyl phosphates, heat treatment resulted in a more rapid loss of the activity toward bis-p-nitrophenyl phosphate, thus establishing the inhomogeneity of the sample (23).

Another way to show that the same acid phosphatase enzyme is catalyzing the hydrolysis of many substrates is to demonstrate the coincidence of activity toward these substrates after polyacrylamide gel electrophoresis. This was done with each of the phosphate monoesters listed

l P-PI 0 PNPP 0 ATP 6 Glucose-6-P l 5’-AMP

-----s IO 20

FRACTION NUMBER

FIG. 2. Multiple phosphate ester assay of an acid phosphatase isoenzyme mixture accompanying G-75 Sephadex chromatography. The various assay procedures described in the text were used to assay for activity toward inorganic pyrophosphate (01; p-nitrophenyl phosphate (0); 5’-adenosine triphosphate (0); glucose-6-phosphate (A); and 5’. adenosine monophosphate (m). The activity units for each substrate were normalized for clarity in plotting.

in Table II by detecting the appearance of the phosphate ion hydrolysis product using an in situ phosphate-precipitation method (15). The results showed that in each case the samples containing pure WGAP gave a single band of white precipitate using the phosphate precipitation method (15) and a corresponding single activity band using the cY-naphthyl phosphate activity stain. Pyridox- amine-5’-phosphate, which Verjee (5) reported was not a substrate for wheat germ acid phosphatase, was in fact found to be a substrate (see also Table II).

If a single enzyme is responsible for the hydrolysis of the wide variety of substrates that are acted upon by WGAP preparations, then a mixture of substrates should compete for the active site and result in a reaction rate that is intermediate to that given by the substrates sep- arately. When the product of only one substrate is observed (e.g.,p-nitrophenol fromp-nitrophenyl phosphate), then the other substrate generally behaves as a compet-

0

:>,

40 50 60 70 80

Temperature (” C )

FIG. 3. Thermal inactivation of the isoenzyme WGAP. The enzyme was partially inactivated by heating and then assayed for activity toward p-nitrophenyl phosphate (O), bis-p-nitrophenyl phosphate (0) and p- nitrophenyl phenylphosphonate (A).


TABLE II

Michaelis Kinetic Constants for the Hydrolysis of Phosphate Esters by Wheat Germ Acid Phosphatase at pH 5.0

Substrate K,” v a max (mM) (pm01 mini’ mg-‘)

p-Nitrophenyl phosphate 0.091 I 0.010 605 + 22 cY-Naphthyl phosphate 0.29 + 0.03 325 i 9 @Glycerophosphate 11.3 z!z 1.1 529 * 22 Pyrophosphate 0.027 f 0.004 463 + 9 Tripolyphosphate 0.18 i 0.02 720 f 36 5’.Adenosine triphosphate 0.054 f 0.007 329 f 9 B’Adenosine monophosphate 2.67 + 0.53 254 + 27 5’.Adenosine diphosphate 0.31 + 0.01 218 f 4 5’-Adenosine monophosphate 4.10 + 0.54 49 + 4 D(-)-3-Phosphoglycerate 0.45 + 0.03 187 f 4 Glucose B-phosphate 2.86 + 0.25 151 f 9 Ribose 5-phosphate 2.98 f 0.38 93 f 4 Glucose l-phosphate 28.0 f 1.7 42 f 1 Fructose 6-phosphate 1.04 f 0.22 22 + 2 Pyridoxal5’-phosphate 2.61 z!z 0.08 142 + 4 Pyridoxamine 5’-phosphate 4.71 f 0.83 19 k 2 0-Phospho-L-serine 6.25 f 0.77 124 + 9 Choline O-phosphate 25.1 k 2.2 8 rk 0.4 Ethanolamine phosphate 31.9 f 3.2 138 + 9

o Kinetic constants and standard error were determined using the computer program HYPER (17).

itive inhibitor. To test this, solutions of p-nitrophenyl phosphate were prepared in 0.1 M, pH 4.6, acetate buffer by serial dilution of a concentrated stock solution. Various concentrations of pyrophosphate, adenosine triphosphate, D(-)-3-phosphoglycerate and 5’-AMP were added to the p-nitrophenyl phosphate solutions and the assays carried out at 25°C. Each of these substrates was found to be a competitive inhibitor. Figure 4 shows several examples of the competitive inhibition that was observed.

A further proof that two substrates are acted upon by the same enzyme is the identity of inhibition constants for the product common to the two substrates. Using glucose 6-phosphate and nitrophenyl phosphate as substrates at pH 5.0, competitive inhibition constants for phosphate were determined. The Ki value that was determined using glucose 6-phosphate was 1.3 mM, in excellent agreement with that determined using p-nitrophenyl phosphate as a substrate (1.2 mM). This is also consistent with action by the same enzyme.

In literature studies of substrate specificity, a frequent approach is to measure turnover using a single, fixed substrate concentration. In only a few investigations of plant acid phosphatases have Michaelis constants been determined, and then with a very limited number of substrates. Previous studies of the substrate specificity of wheat germ acid phosphatase were carried out at fixed substrate concentrations (4,5). Such studies are much less informative than measurements of K,,, and V,,,, so more nearly de- finitive results were sought in the present work. The sub-

strate specificity studies were also performed with highly purified isoenzymes that were chromatographically and electrophoretically homogeneous with respect to activity. The results of these studies are summarized in Table II.

It is evident that in terms of Km and V,,,, the highly negatively charged substrates, pyrophosphate, ATP, and tripolyphosphate are comparable to p-nitrophenyl phosphate as “good” substrates. A comparison of the substrates tripolyphosphate, ATP, ADP, AMP, and ribose 5-phosphate is interesting in that V,,,,, decreases with de- creasing negative (phosphate) charge while the removal of the adenosine moiety somewhat reversed the trend in ribose 5-phosphate. The sugar phosphates in general seem to be fair to poor substrates. Several substrates were not investigated beyond the use of a single (high) substrate concentration. Although not investigated in detail, fructose 1,6-diphosphate, DL-a-glyceryl phosphate and phos- phoenol pyruvate are all good substrates, while trimetaphosphate is a moderate substrate (V,,, N 10% that of nitrophenyl phosphate). An important result of these studies is that WGAP seems to be active toward almost any phosphate monoester, although the V,,, values with different substrates may vary widely.

Most of the plant acid phosphatase literature is con- cerned with seedling and leaf sources of the enzyme. Pu- rifications of loo-fold are typical, and although some 700- to 1000-fold purifications have been described, few of them present any evidence for the catalytic purity of the preparations. The overall pattern of WGAP specificity can be compared to that of homogeneous preparations from sweet potato (24), kidney bean (25), barley root (26), soybean (S), sunflower (7), and a partially purified (lOOO-fold) preparation from lupine seedlings (27). Several of those studies were carried out at a single substrate concentration but most show an overall similarity to WGAP in that ATP, ADP, and/or pyrophosphate were also very good substrates. With WGAP, ,&glycerophosphate has a high V,,, that is comparable to that of p-nitrophenyl phosphate. Nevertheless, due to its high Km, it would be a poor substrate if compared under the fixed, nonsaturating substrate conditions employed in some earlier studies. A

01234567

I/CSl

FIG. 4. Competitive inhibition of the WGAP-catalyzed hydrolysis of p-nitrophenyl phosphate by added phosphate esters at pH 5.0.


striking similarity to WGAP is that 3’-AMP is a good substrate while $-AMP is a relatively poor substrate for these enzymes. In general, sweet potato and lupine seedling acid phosphatases appear to be more active than WGAP toward the common sugar phosphates. Because ATPase activity was reportedly absent in the potato enzyme (28), it is possible that the high levels of ATPase activity shown by wheat germ, lupine seedling, soybean, and sweet potato acid phosphatases is not a general property of plant acid phosphatases, but it might be prudent to reexamine the potato enzyme.

The finding of pyrophosphatase activity in many partially purified seed, seedling, and leaf acid phosphatases led to attempts to separate the activities. Studies of these partially purified acid phosphatase preparations (200-fold from pea leaves (29), 700-fold from soybean meal (30), and IOO-fold from dwarf bean seedling (31), led to the conclusion that the activities were inseparable. On the basis of early studies, it was tentatively concluded that a high acid pyrophosphatase activity with no Mg+’ requirement was probably a general property of acid phosphatases (6). The presence of pyrophosphatase activity in homogeneous sweet potato, soybean, sunflower seed, and wheat germ acid phosphatase supports this conclusion.

The specificity of WGAP is distinctly different from mammalian acid phosphatases since ATP and pyrophosphate are either not substrates or are extremely poor substrates for those enzymes. These results also contrast with the total lack of specificity which was found with homogeneous yeast acid phosphatase (32). That enzyme possessed a constant activity (within a factor of 2) for a variety of monoesters, nucleotides, sugar phosphates, pyrophosphate, and ATP. A broader comparison of the present results with those in the literature on substrate specificity is difficult since fixed substrate concentrations were generally used in other studies.

The absence of bis-p-nitrophenyl phosphate phosphodiesterase activity in purified bovine liver (36), spleen (37), and porcine uterine (38) acid phosphatases are typical of the mammalian enzymes. Yeast acid phosphatases also seem to lack this phosphodiesterase activity (32). Essen- tially dual activities towardp-nitrophenyl phosphate and bis-p-nitrophenyl phosphate have been found in two purified fungal acid phosphatases (39,40) while a third contains low levels of activity (41). The dual activity of two purified bacterial acid phosphohydrolases has been established (42, 43). Three highly purified plant acid phosphatases are available for phosphodiesterase comparison. The sweet potato enzyme contains no diphenyl phosphate hydrolase activity (24), while the sunflower seed enzyme possesses a low activity (7). In contrast, soybean acid phosphatase is active against bis-p-nitrophenyl phosphate (8). The acid phosphatases from potatoes and lupine seedlings have been partially purified and exhibit no significant diesterase activity (27, 28).

The present studies at pH 5 indicate that it is not possible to separate the diesterase and monoesterase activities of WGAP. To confirm this, we selected examples of several substrate types and measured the K,,, and V,,, values at several pH values (Table III). Both of the diester substrates exhibit a higher maximal velocity at pH 8.5 than at pH 7.0. The maximal velocity with p-nitrophenyl phenylphosphonate showed little variation with pH. Be- cause molybdate ion inhibited both bis-p-nitrophenyl phosphate and p-nitrophenyl phosphate activities, it is likely that WGAP is hydrolyzing both substrates. While bis-p-nitrophenyl phosphate has a V,,, which is about 16% of that of p-nitrophenyl phosphate at pH 8.5, it is

TABLE III

The variability in V,,, indicated by the WGAP studies at pH 5.0 (Table II) is not consistent with a rate-limiting dephosphorylation of a common phosphoenzyme intermediate. This has been discussed in terms of the pH dependence of the apparent rate-limiting step (33). This behavior also differs from the results obtained with prostatic acid phosphatase, where a constant V,,, was found with a wide variety of substrates (34), and with a cyto- plasmic phosphotyrosyl phosphatase, where burst titra- tion kinetics have been convincingly demonstrated (35).

Hydrolysis of p-Nitrophenyl Phenylphosphonate and Several Phosphodiesters at pH 5.0, 7.0, and 8.5

K,” v a max Substrate PH (mM1 (mnol mini mg-‘)

p-Nitrophenyl 5.0 5.7 f 0.95 0.99 f 0.08 phenylphosphonate 7.0 1.9 + 0.09 1.07 f 0.09

8.5’ 6. 0.63 t 0.06 bis-p-Nitrophenyl phosphate’ 5.0 nd 0.22 f 0.02

7.0 9.6 I? 1.49 0.41 f 0.04 8.5’ 10. 5.4 * 0.5

Thymidine&phosphate- 5.0 nd 0.15 f 0.02 p-nitrophenyl esterd 7.0 0.53 0.07 f 0.01

8.5b 10. 1.52 p-Nitrophenyl phosphate 5.0 0.09 f 0.01 609 f 22

7.0 0.76 + 0.02 569 f 9 8.5 1.08 + 0.06 33 2 4

The low specific activity of the enzyme preparations and the fixed substrate concentration conditions employed in an earlier study (5) appear sufficient to explain the differences with the present substrate specificity results. Thus, an inadequate fixed substrate concentration is the reason that pyridoxamine 5’-phosphate (with a relatively large Km and small V,,,, Table II) was found by Verjee not to be a substrate (5). This emphasizes the need to measure K,,, and V,,, values in such studies. “2mM.

’ Kinetic constants and standard error were determined using the computer program HYPER (17).

b Estimated graphically. ’ 10 mM.


SCHEME I

estimated to be less than 0.1% at pH 5.0. The stoichiometry of p-nitrophenol release and of inorganic phosphate release was measured to be 1.9:1 at both pH 5.0 and 8.5, consistent with the release of two moles of p- nitrophenol and a mole of inorganic phosphate from each mole of bis-p-nitrophenyl phosphate. In the experiment at pH 8.5, the bis-p-nitrophenyl phosphate concentration (10 mM) remained near the K, of bis-p-nitrophenyl phosphate while the maximum concentration of p-nitrophenyl phosphate which could have been produced as a product during this experiment was about 0.2 mM (which is about one-fifth the K,,, of p-nitrophenyl phosphate); Thus, the subsequent rebinding and hydrolysis of a p-nitrophenyl phosphate intermediate after release from the enzyme would not be favored, suggesting that the p-nitrophenyl phosphate intermediate may not be released before the second cleavage step occurs to yield phosphate and the second mole of p-nitrophenol. A similar result was found with a fungal enzyme that was purified as an acid phosphatase and renamed as a phosphodiesterase-phosphomonoesterase (44). Thymidine 5’-phosphatep-nitrophenyl ester was the only nucleotide diester tested with WGAP and it appears to be a relatively poor substrate (Ta- ble III).

It is apparent that varying degrees of phosphodiesterase activity may be considered to be a characteristic feature of many nonmammalian acid phosphatases. The apparent ability of WGAP to act as a relatively poor “alkaline phosphodiesterase” appears to be unique among the acid phosphatase enzymes reported to date, although a detailed study might reveal similar kinetics for the soybean enzyme (8). This interesting property probably has limited biological significance, but it does carry implications with respect to the mechanism of the enzyme. The apparent ability to cleave two P-O bonds before product release, which is indicated for bis-p-nitrophenyl phosphate as a substrate, is consistent with a similar action by WGAP on trimetaphosphate, as will be described subsequently.

The use of p-nitrophenyl phenylphosphonate (Scheme I) provided information on several points. In Fig. 5 the pronounced biphasic nature of the Lineweaver-Burk plot that is evident is more extreme than that encountered at certain pH values with p-nitrophenyl phosphate (12). Nonlinear plots with p-nitrophenyl phenylphosphonate could be due to a diesterase contaminant. One might ex- pect that p-nitrophenyl phenylphosphonate would be a substrate for some possible diesterase contaminants since it has been proposed as a specific substrate for certain phosphodiesterases (45). Estimation of the maximal ve-

FIG. 5. Lineweaver-Burk plot of the WGAP-catalyzed hydrolysis of p-nitrophenyl phenylphosphonate at pH 8.5.

locity using the high substrate concentration points gave values that were only l-3% of V,,, measured with p-nitrophenyl phosphate under comparable conditions. How- ever, there is a concomitant loss of activity toward p-nitrophenyl phosphate and p-nitrophenyl phenylphosphonate during heat denaturation (Fig. 3). Thus, it appears that WGAP is capable of hydrolyzing both substrates, although with considerably less activity toward the phenylphosphonate.

Activity of WGAP toward p-nitrophenyl phenylphosphonothioate (Scheme II) was investigated because some enzymatic activity toward this substrate is present in crude WGAP preparations. It was reported that acid phosphatases (including a commercial preparation of the wheat germ enzyme) exhibit similar maximal velocities with p-nitrophenyl phosphate and 0-p-nitrophenyl thio- phosphate as substrates (46). This substrate was of par- ticular interest because it could be obtained in an optically active form, with the chirality at phosphorus. Initial experiments with the phenylphosphonothioate indicated that it was a poor substrate at pH 5, but that some catalysis could be observed at pH 8. To determine if WGAP was responsible for activity toward this substrate, the activity and the effect of added EDTA was measured as a function of WGAP purity. Another method employed was to measure the activity toward p-nitrophenyl phosphate and p-nitrophenyl phenylphosphonothioate across the elution profile of a preparative G-75 Sephadex chromatographic run. For these studies the standard assay for p- nitrophenyl phenylphosphonothioate activity was a direct continuous assay with 3 mM substrate in pH 8.0 barbital buffer. We found that relative activity of highly purified samples of WGAP toward p-nitrophenyl phenylphos-

SCHEME II


phonothioate was less than 1% of that observed in the initial extract of wheat germ. The activity toward p-ni-

trophenyl phenylphosphonothioate was inhibited by EDTA to a greater extent in low specific activity preparations than in the pure enzyme. This, along with the decrease in total p-nitrophenyl phenylphosphonothioate activity, indicated that most of the p-nitrophenyl phenylphosphonothioate activity was being removed during purification.

The identity of the enzyme responsible for the phosphonothioate activity deserves further study. Assay of the elution profile during a preparative G-75 Sephadex chromatography run revealed single, approximately coincident activity peaks. However, comparison of the activity ratios suggested that thep-nitrophenyl phenylphosphonothioate activity had a slightly higher (5-10 kDa) molecular weight. Further evidence of the presence of a p-nitrophenyl phenylphosphonothioate-hydrolyzing impurity was the lack of effect of 1.0 mM tungstate ion, a potent inhibitor of the phosphatase (47), on the hydrolysis ofp-nitrophenyl phenylphosphonothioate by a crude enzyme sample.

The ability of highly purified WGAP to hydrolyze trimetaphosphate completely to inorganic phosphate is tac- itly assumed by the use of the phosphate assay to detect activity toward trimetaphosphate. However, there was a possibility that the presence of impurities such as tripolyphosphate or pyrophosphate in the trimetaphosphate substrate could also explain the apparent activity toward trimetaphosphate, since both of those ions are good substrates for WGAP (Table II). Furthermore, the ring opening process required during hydrolysis of trimetaphosphate could produce tripolyphosphate and pyrophosphate as intermediate products. These questions about substrate contamination and intermediate product accumulation were examined by using 31P NMR spectroscopy. The trimetaphosphate spectra in 99% DzO and buffered in 0.05 M pH 4.0 acetate in 90% D80 were identical and consisted of a single line at -21.5 ppm (Fig. 6). This is expected from the presence of three equivalent phospho groups in the six-membered trimetaphosphate ring and indicates a high degree of purity of the sample as well as the stability of the ring at pH 4 during the time course of these experiments. The spectrum of inorganic pyrophosphate under the same conditions at pH 4 yielded a single line at -10.6 ppm while (linear) tripolyphosphate gave a doublet (-9.94 and -10.4 ppm) and a triplet (-22.2, -22.7, and -23.2 ppm) as characteristic spectral features of these compounds (Fig. 6). Trimetaphosphate did not contain detectable amounts of these ions.

It was important to demonstrate that WGAP activity toward trimetaphosphate was not due to action on pyrophosphate or tripolyphosphate, since these substrates could be present in low to moderate amounts, but might not be detectable by NMR. The disappearance of the trimetaphosphate line at -21.5 ppm was observed after the addition of about 8 units of WGAP to 3.10 ml of 20 mM

-IOppm15

us L - -- e-J 0 -5 -10 -IS -20 -25

I wm

Lc 0 -5 -10 -15 -20 -25

PPm

IO b -i lpii5 4”;:

0 -5 -10 -15 -20 -25 wm

FIG. 6. Identification by 31P NMR spectroscopy of intermediates in the WGAP-catalyzed hydrolysis of trimetaphosphate. NMR spectra were taken as described in the text. (A) Spectrum of trimetaphosphate before addition of enzyme to the sample. (B, C) Spectra taken 20 and 90 min after addition of a catalytic amount of WGAP. Spectrum D is of inorganic pyrophosphate, while spectrum E is linear tripolyphosphate.

trimetaphosphate at pH 4. Within 20 min a large inorganic phosphate peak (0 ppm) had appeared. In addition to a diminished trimetaphosphate peak at -21.5 ppm, the spectrum contained a small new peak at -10.7 ppm that was identified as pyrophosphate (PPi). The relative peak heights, trimetaphosphate:Pi:PPi after 20, 90, and 180 min incubation with the enzyme (Fig. 6) were: l.OO:l.ZO: 0:30,1.00:3.91:0.43, and l.O0:7,53:nd, respectively. (In the 3-hr spectrum, the pyrophosphate peak was barely distinguishable from baseline noise.)

The decrease in the trimetaphosphate peak, with a concomitant increase in the inorganic phosphate peak, provided direct evidence that the WGAP preparation is active toward trimetaphosphate and that it apparently cleaves two P-O bonds before release of products. There was no evidence for accumulation of tripolyphosphate as an intermediate product.

The possibility that tripolyphosphate is released and rebound before subsequent hydrolysis was considered in more detail. The K, (0.18 mM) and high V,,, for tripolyphosphate (Table II) indicate that it is one of the best substrates for WGAP. It is reasonable to postulate that the complex that would exist prior to a possible release of tripolyphosphate would favor immediate rebinding in a catalytic mode. If tripolyphosphate is released, then it would have to compete with trimetaphosphate for binding


to the active site. Experiments using trimetaphosphate as an inhibitor of p-nitrophenyl phosphate hydrolysis indicated a fairly small binding constant for trimetaphosphate (e.g., 3 mM trimetaphosphate resulted in a 50% inhibition of 0.16 mM p-nitrophenyl phosphate hydrolysis at pH 4.6). This result would indicate that 20 mM trimetaphosphate would effectively compete for the active site and cause tripolyphosphate accumulation if it was released into solution rather than further hydrolyzed before release. To test this possibility, a much more limited (about 10%) hydrolysis of 20 mM trimetaphosphate was allowed to occur before a 31P NMR spectrum was obtained. This spectrum displayed clearly identifiable inorganic phosphate and inorganic pyrophosphate peaks but showed no evidence of tripolyphosphate. The evidence indicates that two P-O bonds of trimetaphosphate are cleaved before product release.

WGAP was found to hydrolyze phosphoprotein substrates. Ovalbumin and phosvitin (both phosphoserine esters) are substrates for WGAP with rates of dephosphorylation of 0.014 and 0.023 mol of Pi min-’ rng-‘, respectively. The action of WGAP toward phosvitin seems low considering the very high phosphate content of phosvitin, but an early study (48) with a crude wheat kernel phosphatase preparation indicated that only phosphoserine residues peripheral to the predominant heavily phosphorylated segments were susceptible.

Electrophoresis of ovalbumin revealed the presence of two bands (Fig. 7) which were presumed to be the Ai (2 phosphates, hence, electrophoretically more mobile) and AZ (1 phosphate) forms found by Perlmann (49). These were converted to the less mobile dephosphorylated form A3 by treatment with WGAP (Fig. 7C). (A second new band of low intensity which is just slightly more mobile than the dephosphorylated band also appeared; its nature is unknown.) The activity of WGAP toward ovalbumin, with which total dephosphorylation is observed, differs from the reported action of prostatic acid phosphatase upon ovalbumin, where only one of two phosphates is removed (49). The phosphoprotein phosphatase activity of homogeneous WGAP is in contrast to the removal of phosphoprotein phosphatase activity (measured using casein as substrate) that was observed during the course of a lOOO-fold purification of an acid phosphatase from soybean meal (30). A recent study of thylakoid membrane acid phosphatase and protein phosphatase of wheat concluded that the acid phosphatase present in the membrane plays a minor role, at best, in the dephosphorylation of thlakoid membrane phosphoproteins. Based on its reported kinetic properties (50), that acid phosphatase seems distinct from the present one.

It is interesting that, despite the very similar mecha- nisms suggested for WGAP (51) and prostatic acid phosphatase (52), pyrophosphate and ATP are very poor substrates for the latter enzyme. In other respects the specificities of WGAP and prostatic acid phosphatase re-

A B C D

FIG. 7. Electrophoresis of ovalbumin modified by WGAP. Electro- phoresis was performed at pH 8.0 as described in the text. A (5 fig) and B (20 gg) are untreated ovalbumin; C (20 wg) is WGAP-treated; D (10 fig) is a 1:l mixture of treated and untreated ovalbumin. A,, A,, and A, denote 2, 1, and 0 phosphates attached.

semble those of other acid phosphatases with respect to their ability to hydrolyze many types of P-O bonds, albeit at widely varying efficiencies; this suggests a similar catalytic mechanism for all such acid phosphatases. This similarity was explored in further detail by studying the detailed pH dependence (53-55).

The pH dependencies of the Michaelis constants for the hydrolysis of p-nitrophenyl phosphate and /3-glyceryl phosphate as catalyzed by homogeneous WGAP are given in Tables IV and V, respectively. Presented graphically, these results would resemble those obtained for human prostatic acid phosphatase (52). The variation of log V,,,,, versus pH corresponds to a broad plateau from pH 4.5 to 7, with inflections at the extremes of pH which result in nearly unit changes in slope, consistent with the ionization of a single species at each extreme. The prostatic enzyme maintains maximal activity for about 0.6 pH unit higher at alkaline pH. The main difference is a low pH inflection exhibited by WGAP, whereas the prostatic enzyme shows no decrease in V,,,,, until acid denaturation occurs at low pH. The general forms are consistent with the partici- pation of a phosphohistidine covalent intermediate re- sulting from a nucleophilic displacement on the substrate (52). The observed differences may be partially accounted for by the fact that rate-limiting dephosphorylation is found at pH 5 for prostatic acid phosphatase (34), while for WGAP such a dephosphorylation appears to become rate limiting only at high pH (33). For WGAP, a graphical estimation of the apparent pK,‘s given by the inflections yielded values of 3.7-3.8 and 7.2-7.3 for the groups ion- izing at low pH and high pH, respectively. Similar values were obtained with P-glycerophosphate as substrate (Ta- ble V).


Buffer PH K,, (mM) V,. (pm01 min-’ rug-‘)

Glycine 2.59 0.083 + 0.02 35 f 2 2.91 0.11 * 0.01 93 +4 3.22 0.096 f 0.004 169 + 9 3.46 0.10 f 0.01 249 ic 9

Acetate 3.79 0.11 f 0.01 343 + 13 3.99 0.099 f 0.005 446 f 9 4.19 0.10 -t 0.01 489 f 13 4.40 0.11 + 0.01 498 f 9 4.60 0.10 + 0.01 534 + 13 4.79 0.10 f 0.01 498-c 22 5.00 0.09 2 0.01 605 + 22 5.24 0.13 f 0.01 627 2 9 5.53 0.15 f 0.02 632 ?z 22

Dimethyl glutarate 5.50 0.18 + 0.01 792 + 9 5.75 0.17 f 0.01 627 f18 6.02 0.20 f 0.01 702 f 9 6.25 0.21 f 0.01 556+ 9 6.48 0.32 + 0.01 672 k9 6.76 0.50 f 0.02 498 f 9 6.98 0.76 f 0.02 564 f 9 6.02 0.62 3~ 0.18 387 f 62 6.25 0.69 k 0.02 427 f 9 6.95 0.36 f 0.02 520f 9 7.22 0.59 -+ 0.02 369 + 9 7.50 0.81 f 0.01 284-c 5 8.01 0.99 f 0.07 106 f 2 8.15 1.11 f 0.04 68 f 2 8.50 1.08 f 0.06 33 + 4 8.76 1.01 + 0.14 12 * 4 8.97 0.83 k 0.11 9.0 f 0.3 9.20 0.99 + 0.10 5.3 + 0.2

Sucinate Maleate Barbital

TABLE IV

pH Dependence of the Michaelis Constants for the Hydrolysis of p-Nitrophenyl Phosphate by Homogeneous

Wheat Germ Acid Phosphatase

The biphasic reciprocal plots encountered with WGAP at high pH may affect the estimates of V,, and K,,, (12). The near linearity in the low substrate range of the reciprocal plot always gives a lower apparent V,,,,, and smaller apparent K,,, and results in log V,, slopes between -1 and -2 at high pH for WGAP. In Fig. 1 of Ref. (56) a slope of -1.4 in the log V,,, plot is evident (although the author represented the slope as -l.O), which leads one to suspect that V,,, determinations for potato acid phosphatase were subject to similar biphasic behavior at high pH.

The “pH dependence” of the turnover of enzymes such as acid phosphatases is often measured at a single fixed substrate concentration. The shape of the curve and the pH optimum that results depends on the substrate concentration and reflects not only changes in maximal velocity but also changes in K,,, with pH. When the pH dependence of K,,, is known, a high fixed substrate concentration may be employed as a strategy to approximate maximal velocity. However, the broad plateau that is obtained with WGAP in the pH range 4.5-7 for the depen-

dence of V,,, on pH would probably not be observed at a fixed high substrate concentration due to substrate inhibition (which occurs at low pH) and non-hlichaelis- Menten kinetics and inactivation (which occur at high pH). Thus, not only is the use of a single substrate concentration inadequate, but in addition a determination of kinetic constants must be carried out over a wide substrate range in order to detect kinetic anomalies. For example, the use of the same substrate concentration range in the high pH range as employed in the low pH range would result in anomalously low V,,, and K,,, values due to the biphasic character of the plot used for constant determination.

The small lo-fold variation in K,,, observed for the WGAP-catalyzed hydrolysis of p-nitrophenyl phosphate is in strong contrast to that of the potato acid phosphatase (56) or prostatic acid phosphatase (57), where changes in Km of 200-fold and 300-fold, respectively, were observed from about pH 5 to pH 8. However, the 3.6-fold change in K,,, that is exhibited by potato acid phosphatase with P-glycerophosphate as a substrate is similar to that for WGAP.

Taken together, these experiments make a strong case for the ability of a single WGAP enzyme to catalyze the hydrolysis of a wide variety of phosphate monoesters, polyphosphates and diesters. They also show that the enzyme does not possess carboxylesterase activity. Evidence for a single active enzyme species was provided by gel electrophoresis and by competitive inhibition experiments, as well as direct kinetic measurements. The diesterase activity of WGAP was lost concomitantly with p- nitrophenyl phosphatase activity during heat treatment.

In conclusion, several provocative hypotheses that arose in connection with early experiments on wheat germ acid phosphatase have been disproved. The enzyme does not possess significant phytase or carboxylesterase activity. Although initial extracts exhibited a high activity toward phosphonothioate esters, this activity effectively disap-

TABLEV

pH Dependence of the Michaelis Constants for the Hydrolysis of P-Glyceryl Phosphate by Homogenous

Wheat Germ Acid Phosphatase

Buffer PH Km (mM) V,,, (pm01 min-’ mg-‘1

Glycine 2.90 7.1 + 0.8 46 + 2 Acetate 3.50 14.4 f 2.7 182+ 31

3.90 13.8 + 2.1 311+ 27 4.50 10.5 f 1.4 418+ 31 5.00 11.3 -c 1.1 529+ 22

3,3-Dimethyl glutarate 5.75 12.3 3~ 0.9 512 k 13 6.26 15.1 2 1.4 444 f 18

Barbital 7.30 20.1 f 1.6 222 +13 7.80 44.1 k 3.8 58 AI 4 8.25 32.3 + 1.8 30+1


peared during purification of the acid phosphatase. On the other hand, the pure enzyme exhibits a broad activity against a remarkable range of phosphomonoesters, diesters, and polyphosphates. Interesting kinetic behavior is seen in the hydrolysis of trimetaphosphate, where the two P-O bonds are cleaved before product release. These and other aspects, including the action of the enzyme on phosphoprotein substrates and a determination of the protein primary structure, all merit further study. How- ever, the purification of large amounts of this enzyme remains a problem.

ACKNOWLEDGMENTS

This research was supported by NIH Research Grant GM 27003. We thank Professor Stephen Benkovic for a gift of p-nitrophenyl phenylphosphonothioate.

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substrate specificity and ph dependence of homogeneous ... · homogeneous wheat germ acid...

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