Assay of acetohydroxyacid synthase

ANALYTICAL

BIOCHEMISTRY

171, 173-179 (1988)

Assay of Acetohydroxyacid BIJAY K. SINGH,’ American

Cyanamid

MARK Company,

A. STIDHAM,

Synthase AND DALE

P.O. Box 400, Princeton,

L. SHANER

New Jersey

08540

Received October 2, 1987 Acetohydroxyacid synthase (AHAS), also known as acetolactate synthase, has received attention recently because of the finding that it is the site of action of several new herbicides. The most commonly used assay for detecting the enzyme is spectrophotometric involving an indirect detection of the product acetolactate. The assay involves the conversion of the end product acetolactate to acetoin and the detection of acetoin via the formation of a creatine and naphthol complex. There is considerable variability in the literature as to the details ofthis assay.We have investigated a number of factors involved in detecting AHAS in crude ammonium sulfate precipitates using this spectrophotometric method. Substrate and cofactor saturation levels, pH optimum, and temperature optimum have been determined. We have also optimized a number of factors involved in the generation and the detection of acetoin from acetolactate. The results ofthese experiments can serve as a reference for new investigators in the study ofAHAS. o 1988 Academic KEY

Ress, Inc. WORDS:

imidazolinones;

acetohydroxyacid synthase; acetolactate synthase; amino acid biosynthesis; sulfonylureas.

Acetohydroxyacid synthase (also known as acetolactate synthase, EC 4.1.3.18) is the first enzyme unique to the biosynthesis of the branched chain amino acids valine, leucine, and isoleucine (1,2). This enzyme is under feedback regulation by these amino acids in plants (3,4). AHAS catalyzes the following reactions in this biosynthetic pathway: 2 Pyruvate + Acetolactate Pyruvate + a-ketobutyrate

+ CO2

+

Acetohydroxybutyrate

+ CO2 .

This enzyme uses thiamine pyrophosphate (TPP) as the co-enzyme in these condensation reactions. An intermediate formed between pyruvate and the TPP C-2 carbanion undergoes decarboxylation to the stabilized anion of hydroxyethyl-TPP, which in turn ’ To whom correspondence should be addressed. * Abbreviations used: AHAS, acetohydroxyacid synthase; TPP, thiamine pyrophosphate; BMS, Black Mexican Sweet. 173

acts as a nucleophile on the 2-keto group of a second molecule of pyruvate or cYketobutyrate, releasing TPP and acetolactate or acetohydroxybutyrate (5). Flavin adenine dinucleotide (FAD) is also a co-factor which stabilizes the enzyme and causes a moderate increase in the activity, though the role of FAD is obscure since no net oxidation or reduction occurs in the reaction. Magnesium chloride is required for the assay, presumably for the binding of TPP to the enzyme. Production of acetolactate during this reaction is measured spectrophotometrically after conversion of acetolactate to acetoin (6). In recent years, AHAS has received a great deal of attention with the discovery that this enzyme is the site of action of two different new classes of herbicides: imidazolinones and sulfonylureas (7- 10). This renewed interest in this enzyme has resulted in the appearance of numerous papers in the literature on this subject. However, a systematic study on the assay of this enzyme has never been published, and each group has adopted 0003-2697188 $3.00 Copyright 0 I988 by Academic Pres, Inc. All rights of reproduction in any form reserved.

174

SINGH,

STIDHAM,

its own assay protocol. Several of these protocols seem to have been adopted without much time spent on the standardization of the assay itself, resulting in an underestimation of the product and a lengthy assay. In this report, we present data on each and every aspect of this assay from which we have generated a protocol which is not only very sensitive, but also less time-consuming. Any saving in time is of importance during enzyme purification, especially in this particular case where the enzyme is very labile (7,11-13). MATERIALS

AND

METHODS

Black Mexican Sweet (BMS) cells. Embryo-derived cell suspension cultures of Zea mays var. Black Mexican Sweet, were obtained from Molecular Genetics Inc. (Minnetonka, MN) and cultured on MS salts ( 14,15) with 2% (w/v) sucrose, 0.5 mg/ml thiamine, 0.15 mg/ml L-asparagine, and 2 mg/liter 2,4-D. Cells were harvested on Day 7, the age normally used for subculturing. Enzyme extraction. For the extraction of AHAS, 10 g of cells was powdered in liquid nitrogen and then homogenized in 100 mM potassium phosphate buffer (pH 7.5) containing 10 mM pyruvate, 5 mM MgClz ,5 mM EDTA, 100 PM FAD, 1 mM valine, 1 mM leucine, 10% glycerol, and 10 mM cysteine. The homogenate was filtered through a nylon cloth (53-pm mesh) and centrifuged at 25,000g for 20 min. The supernatant fraction was brought to 50% saturation with respect to (NH&SO4 and allowed to stand for 20-30 min on ice. It was then centrifuged at 25,000g for 20 min and the supematant was discarded. The ammonium sulfate pellet was dissolved in 50 IkIM potassium phosphate buffer (pH 7.5) containing 1 mM EDTA and 100 mM NaCl and used for the assay procedures. Ammonium sulfate (0.2-20%) and NaCl(O.0 1- 1 M) have no inhibitory effect on the AHAS activity; however, desalting of enzyme is recommended to accurately control the salt content of the assay mixture.

AND

SHANER

AHAS assay. The AHAS assay described here is a modification of the assay procedure described previously (3). AHAS activity was measured by estimation of the product, acetolactate, after conversion by decarboxylation in the presence of acid to acetoin. Standard reaction mixtures contained the enzyme in 50 XIIM potassium phosphate buffer (pH 7.0) containing 100 mM sodium pyruvate, 10 mM MgC12, 1 mM thiamine pyrophosphate, and 10 PM FAD. This mixture was incubated at 37°C for 1 h after which time the reaction was stopped with the addition of H2S04 to make a final concentration of 0.85% H2S04 in the tube. The reaction product was allowed to decarboxylate at 60°C for 15 min. The acetoin formed was determined by incubating with creatine (0.17%) and 1-naphthol ( 1.7% in 4 N NaOH) by the method of Westerfeld (6). Maximum color was observed by incubation at 60°C for 15 min and then further incubation at room temperature for 15 min. The absorption of color complex was measured at 520 nm. Appropriate checks of direct acetoin formation during the enzyme assay were made. Each assay was run at least in duplicate and the experiments were repeated a minimum of two to five times. RESULTS

AND

DISCUSSION

Temperature In order to find the temperature optimum for AHAS, the assay was performed under the standard conditions as discussed under Materials and Methods at different temperatures. AHAS activity increased with the increase in temperature (Fig. 1) with the maximum activity between 46 and 50°C. However, at higher temperatures (40°C and above), there was a reduction in the sensitivity of AHAS to leucine and valine. Although this reduction in sensitivity to leucine and valine was small, the observation was highly reproducible. Therefore, AHAS was assayed at 37°C as a routine protocol even though

ACETOHYDROXYACID

SYNTHASE

175

ASSAY

a

6.

25

30

35

2

40

TEMPERATURE

45

50

5

6

7

6

(‘C)

Ftc. 1. Formation of acetolactate by AHAS from BMS cells as a function of temperature. The enzyme was assayed under the standard reaction conditions as described under Materials and Methods except that the temperature of assay was varied as indicated. Control (O), leucine + valine (A), imazapyr (m).

the activity was significantly lower than the maximum observed. In the literature, there is a great variability in the temperature at which AHAS has been assayed. Some of the temperatures at which AHAS has been assayed include 22 (16), 25 (17), 30 (2,10,18), 37 (19,20), and40”C (21). Differences in the temperature optima for enzymes from different sources is expected, but such a wide range is unlikely. Results presented in Fig. 1 show that AHAS from BMS cells has the maximum activity at 46-50°C nearly 50% greater than the activity at 37°C. These temperatures may not be good for some kinetic studies where the property of the enzyme is altered due to high temperatures, e.g., sensitivity to leucine and valine. However, in cases where quantitative data is not required (e.g., location of enzyme activity after chromatography), the enzyme may be assayed at higher temperatures for a shorter period of time.

9

PH FIG. 2. pH optimum of AHAS from BMS cells. AHAS was assayed under the standard reaction conditions as described under Materials and Methods except that the pH of assaywas varied as indicated. Phosphate (A), Tris (m), 4-morpholine ethane sulfonic acid (0).

enzyme (data not shown). Therefore, phosphate was chosen for the standard protocol. It is recognized that pH optima for an enzyme vary from one source to the other; therefore, an optimization of the pH for assay of AHAS is recommended for each new species.

Substrate and Co-factor Requirement Results presented in Fig. 3 show that the pyruvate saturation curve is hyperbolic. The K, for pyruvate is 5 mM and the enzyme is

6 . 5-

l -

w 0

0

1

0

10

20

30

40

50

PI

PH AHAS was assayed at several different pH values using three different buffer systems. As shown in Fig. 2, there is a broad pH op tima for AHAS between pH 6 and 7. There was no difference in the buffer system used, but phosphate had a stabilizing effect on the

Oh 0

50

100 PYRUVATE

150

200

(mu)

FtG. 3. Substrate saturation curve for pyruvate with AHAS from BMS ceils. AHAS assay was conducted under the standard reaction conditions as described under Materials and Methods except that the concentration of pyruvate was varied as indicated. The inset is a [S]/v versus [S] plot (Hanes Woolf plot) of the data.

176

SINGH, STIDHAM,

1 .o

1.5

2.0

2.5

TPP (mM)

AND SHANER

0

2

4

6 MO2

6

10

(mM)

FIG. 4. Saturation curve for TPP with AHAS from BMS cells. AHAS assay was conducted under the standard reaction conditions as described under Materials and Methods except that the concentration of TPP was varied as indicated. The inset is a [S]/v versus [S] plot of the data.

FIG. 5. Saturation curve for MgQ with AHAS from BMS cells. AHAS assay was conducted under the standard reaction conditions as described under Materials and Methods except that the concentration of MgC12 was varied as indicated. The inset is a [S]/v versus [S] plot of the data.

saturated at 100 mM pyruvate. Similar results have been reported for enzyme from barley (3) in the range of pyruvate concentration examined here. However, at low concentrations of pyruvate, a sigmoid curve of pyruvate saturation was observed in the previous study. The point of interest here is that the standard protocol for the AHAS assay in the same and other works (1517) have included only 20 mM pyruvate. At such a concentration of pyruvate, the enzyme would not be saturated. A significant underestimation of the total enzyme activity would result. Assay of the enzyme at unsaturating concentrations of substrate may also lead to misleading kinetic constants for the co-factors and the inhibitors of this enzyme. In the absence of TPP and MgCl,, there was very little AHAS activity (Figs. 4 and 5). The saturation curve for both co-factors is hyperbolic and the enzyme is saturated at 0.5 and 1 mM of TPP and MgC12, respectively. On the other hand, there appears to be no absolute requirement for FAD by the enzyme in a relatively crude preparation (Fig. 6). However, there is about 35-40% enhancement of AHAS activity with 5 PM or higher concentrations of FAD. Lack of response to low concentrations of FAD in the

preparation used here may be due to residual FAD bound to the enzyme from high concentrations of FAD (100 FM) used during the extraction procedure (7). It would be interesting to examine the FAD requirement in a highly purified preparation of AHAS. A similar requirement of FAD by AHAS from microbial sources has also been reported (22,23). Since the reaction catalyzed by AHAS involves no net oxidation or reduction, the role of flavin in this process is still obscure.

FlG. 6. Saturation curve for FAD with AHAS from BMS cells. AHAS assay was conducted under the standard reaction conditions as described under Materials and Methods except that the concentration of FAD was varied as indicated. The inset is a [S]/v versus [S] plot of the data.

ACETOHYDROXYACID

SYNTHASE

177

ASSAY

Conversion of Acetolactate to Acetoin 0.6

Decarboxylation of acetolactate to acetoin should be a standard procedure during this assay, but even this step varies in the literature from 5 min at 80°C (9) to 60 min at 40°C (17). Our results presented in Fig. 7 show that heating in acid at 60°C requires 15 min for complete decarboxylation of acetolactate. Acidic pH stimulates decarboxylation of acetolactate. Therefore, we examined the acid concentration required for decarboxylation of aCetOla&tte. The rt?SUltS presented here (Fig. 8) show that there is no effect of acid concentration (above 0.625%) on the decarboxylation of acetolactate to acetoin. The lowest acid concentrations used here reduced the pH of the mixture to 2.5. Higher acid concentrations cause precipitation of I-naphthol. Therefore, a minimum acid concentration was used for the decarboxylation of acetolactate.

Detection of Acetoin The standard procedure for the detection of acetoin is by the method of Westerfeld (6). In this procedure, mixing of acetoin with creatine and 1-naphthol gives a pink-colored complex. The absorption spectrum of this complex is shown in Fig. 9. The spectrum shows a broad maxima at 520-530 nm. ,

6,

$ 3 g o’4 2 E 0.2 9 0.0

0

-1 1

2

3 H2S04

4

5

(%)

FIG.8. Acid requirement for decarboxylation of acetolactate to acetoin. At the end of incubation of AHAS with substrate and co-factors, the reaction mixture was acidified with H2S04 at the indicated final concentration. Acidified mixtures were heated at 60°C for 15 min and then acetoin was determined by the method described under Materials and Methods. Without enzyme

(a), with enzyme(A).

The dependence of acetoin color development on creatine and I-naphthol was examined in separate experiments. Data presented in Figs. 10 and 11 show that both of these components are absolutely required for color development. There was no dependence of color development on creatine concentration in the range examined here (0.02 to 0.34%). On the other hand, color development responded to the increasing concentrations of I-naphthol with maximum color at a final concentration of 1.33 to 1.67%. After acetoin was mixed with creatine and I-naphthol, the time allowed for color devel-

1 -0

5

10

15

20

25

30

TIME (min)

FIG. 7. Decarboxylation of acetolactate to acetoin. After acidification with H2S04 the mixture was heated at 60°C for the indicated amount of time after which acetoin was determined by the method described under Materials and Methods.

0.0 -

400

500 WAVELENGTH

600 (nM)

FIG.9. Absorption spectrum of the acetoin/creatine/ I-naphthol complex.

178

SINGH, STIDHAM,

AND SHANER

I

0.0 0.1

0.0

0.2 CREATINE

0.3

15

(%)

opment has been reported to be different in different publications. In the original publication (6), a color development time of 60 min at room temperature was reported. Since then, several variations have appeared in the literature. We found that heating accelerated color development in agreement with another study (12). In our experiments, maximum color at room temperature is observed between 45 and 60 min (Fig. 12). On the other hand, heating the acetoin/lnaphthol/creatine mixture at 60°C for 15 min and then allowing it to stand at room temperature gives maximum color in an-

1.6 1.5 1.2 0.9 0.6 0.3 0.0 0.3

0.6

0.9 NAPHTHOL

45

60

TIME (min)

FIG. 10. Color development of the acetoin/creatine/lnaphthol complex at varying concentrations of creatine and acetoin as indicated. I-Naphthol was used at a final concentration of 1.7%. Acetoin concentrations used were 41.6 (O), 83.3 (A), and 166.7 (m) pM.

0.0

30

1.2

1.5

(%)

FIG. 11. Color development of the acetoin/creatine/ lnaphthol complex at varying concentrations of naphthol and acetoin as indicated. Creatine was used at a final concentration of 0.17%. Acetoin concentrations used were 41.6 (O), 83.3 (A), and 333.3 (m) pM.

FIG. 12. Color development of the acetoin/creatine/ lnaphthol complex at room temperature (0) or after heating at 60°C for 15 min (A) and then keeping the mixture at room temperature. The time indicated in the figure is from the time of mixing acetoin with creatine and I-naphthol.

other 15 min (total color development time, 30 min). The maximum absorbance observed in the heating treatment is similar or higher than the maximum color obtained in the unheated control. In conclusion, the data presented here represent the first set of information ever published on the different aspects of the assay of AHAS. The standard protocol for AHAS assay should be as described under Materials and Methods. However, an optimization of the pH, temperature, substrate, and co-factor saturations is recommended for each source of the enzyme. Results presented here not only elaborate the different steps of this assay, but also show a significant improvement in the assay. The minor details examined here have improved the sensitivity of the assay as well as minimized the time required for the assay. In addition, explanation of the different aspects of this assay would help researchers in taking short cuts where quantitative data is not required. REFERENCES 1. Umbarger, H. E. (1978) Annu. Rev. Biochem. 47, 533-606. 2. Umbarger, H. E. (1983) in Amino Acids: Biosynthesis and Genetic Regulation (Hermann, K. M., and Somerville, R. L., Eds.), pp. 245-266, A& d&n-Wesley, London.

ACETOHYDROXYACID 3. Miflin, B. J. (1971) Arch. Biochem. Biophys. 146, 542-550. 4. Miflin, B. J., and Cave, P. R. (1972) J. Exp. Bof. 23, 511-516. 5. Walsh, C. (1979) Enzymatic Reaction Mechanisms. Freeman, San Francisco. 6. Westerfeld, W. W. (1945) .I. Biol. Chem. 161, 495-502. 7. Muhitch, M. J., Shaner, D. L., and Stidham, M. A. (1987) Plant Physiol. 83,45 l-456. 8. Shaner, D. L., Anderson, P. C., and Stidham, M. A. (1984) Plant Physiol. 76, 545-546. 9. LaRossa, R. A., and Schloss, J. V. (1984) J. Biol. Chem. 259,8753-8757. 10. Ray, T. B. (1984) Plant Physiol. 75, 827-83 1. 1 I. Magee, P. T., and DeRobichon-Szulmajster, H. (1968) Eur. J. Biochem. 3,507-5 11. 12. Glatzer, L., Eakin, E., and Wagner, R. P. (1972) J. Bacterial. 112,453-464. 13. Takenaka, S., and Kuwana, H. (1972) J. Biochem. 72, 1139-l 145.

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14. Murashige, T., and Skoog, F. (1962) Physiol. Plant. l&473-497. 15. Anderson, P. C., and Hibberd, K. A. ( 1985) Weed Sci. 33,479-483. 16. Goulden, S. A., and Chattaway, F. W. ( 1969) J. Gen. Microbial. 59, 11 l- 118. 17. Oda, Y., Nakano, Y., and Kitaoka, S. (1982) J. Gen. Microbial. 128, I 2 1 1- 12 16. 18. Ryan, E. D., and Kohlhaw, G. B. (I 974) J. Bucteriol. 120,631-637. 19. Rubin, B., and Casida, J. E. (I 985) Weed Sci. 33, 462-468. 20. Satyanarayana, T., and Radhakrishnan, A. N. (1983) Biochim. Biophys. Acfa. 77, 121-132. 2 1. Wittenbach, V. A., and Erbes, D. L. (1986) Plant Physiol. (Suppl.) 80, 350. 22. Schloss, J. V., Van Dyk, D. E., Vasta, J. F., and Kutny, R. M. (1985) Biochemistry 24, 4952-4959. 23. Stormer, F. C., and Umbarger, H. E. (1964) Biothem. Biophys. Res. Commun. 17, 587-592.