Soil Science Society of America Journal 67:139-145 (2003)
© 2003 Soil Science Society of America
DIVISION S-3—SOIL BIOLOGY & BIOCHEMISTRY
Myrosinase Activity in Soil
Ahmad I. Al-Turkib and
Warren A. Dick*,a
a School of Natural Resources, The Ohio State University, 1680 Madison Ave., Wooster, OH 44691-4096
b King Saud University Qasim Branch, Buraidah Soil and Water Department, P.O. Box 1482, Saudi Arabia
* Corresponding author (dick.5{at}osu.edu)
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ABSTRACT
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Myrosinase (thioglucoside glucohydrolase; EC 3.2.3.1) is an enzyme that hydrolyzes glucosinolates to D-glucose and allelochemicals that have biological potential to suppress weed seed germination in soil. This enzyme, found in some microorganisms and released to soils via root exudation and decomposition, can be assayed by adding sinigrin (2-propenyl-glucosinolate) to soil as a substrate. We describe a simple and rapid method to assay myrosinase activity in soils. In this method, 1 g of soil is treated with toluene (0.2 mL) and incubated at 37°C with 2.8 mL of a buffered solution (pH 7) of sinigrin (20 mM final concentration) for 4 h. Glucose released upon sinigrin hydrolysis is extracted and its concentration is measured spectrophotometrically. Tests showed that recovery of glucose was quantitative if toluene was included in the assay mixture. Myrosinase activity in five soils studied ranged from 71 to 338 µg glucose g-1 soil 4 h-1. The rate of sinigrin hydrolysis increased with temperature from 10 to 40°C. The activation energy of myrosinase in four soils ranged from 40.3 to 52.8 kJ mol-1. The Vmax values for sinigrin hydrolysis calculated by the three linear transformations of the Michaelis–Menten equation ranged from 76 to 518 (avg. 275) µg glucose g-1 soil 4 h-1 and the apparent Km values for myrosinase ranged from 5.3 to 12.9 (avg. 8.1) mM. The method developed in this study is accurate, fast, and simple.
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INTRODUCTION
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MYROSINASE is found in all glucosinolate-containing plants such as Brassica species and possibly in some bacteria and fungi (Rask et al., 2000). Glucosinolates are sugar anionic thioesters containing ß-thioglucoside-type bonds (Brown and Morra, 1996). Myrosinase is normally segregated from glucosinolates in plant tissues, but when plant cells are damaged or decomposed, myrosinase is released and catalyzes the hydrolysis of glucosinolates. The products of glucosinolate hydrolysis include glucose, sulfate, and a number of active allelochemicals such as isothiocyanates, nitriles, thiocyanates, cyanides, and others depending on substrates and reaction conditions used (Gil and Macleod, 1980). These allelochemicals have been found to inhibit weed seed germination and some pathogens in soil (Angus et al., 1994; Brown and Morra, 1997). Recently, several studies have proposed the use of glucosinolate-containing plants as a cover crop to reduce the use of synthetic pesticides (Martin et al., 1990; Boydston and Hang, 1995; Yenish et al., 1996).
Glucosinolates by themselves are not biologically active but must be enzymatically hydrolyzed by myrosinase to the allelochemicals capable of suppressing weed seeds and pathogens (Brabban and Edwards, 1995). Myrosinase is, thus, the key factor for allelochemical expression derived from glucosinolates and hence its study in soil is of interest.
Myrosinase is thought to be released to soils cropped with Brassica or the related Sinapis via root exudation, disruption and decomposition (Borek et al., 1996). Soils cropped with Brassica, therefore, are expected to have enhanced myrosinase activity. Currently, no assay has been developed to measure myrosinase activity in soil although Borek et al. (1996) have studied this enzyme in soil extracts. Extraction of enzymes from soil, however, is a demanding procedure and is often incomplete (Tabatabai and Fu, 1992). Moreover, the conditions used by Borek et al. (1996) to extract myrosinase possibly altered its activity (Ludikhuyze et al., 1999). The procedure to measure myrosinase activity in soil extracts also involved the use of a gas chromatograph coupled to a mass spectrometry, which is not commonly available in many laboratories.
Glucose is a predominant product of glucosinolate hydrolysis and hence a change in its concentration can be used as a direct indication of myrosinase activity. Several studies have measured glucose released from glucosinolates to quantify the activity of myrosinase purified from plant tissues (Wilkinson et al., 1984; Jwanny et al., 1995; Ludikhuyze et al., 1999). Sinigrin, a glucosinolate extracted from horseradish (Armoracia rusticana), has been used in these studies routinely as a substrate for myrosinase and produces D-glucose and allelochemicals.
We propose the use of glucose to measure myrosinase activity in soil. Glucose is extremely water-soluble and can be readily extracted from soil and analyzed (Frey et al., 1999). Different methods have been used to determine glucose concentrations in soil including nonenzymatic (Deng and Tabatabai, 1994) and enzymatic methods. Nonenzymatic methods, however, are time demanding and some require the addition of concentrated acids that heat the soil and thus likely cause chemical hydrolysis of carbohydrates to glucose in soil that will not be distinguished from glucose released from sinigrin hydrolysis. Nonenzymatic methods may also be affected by the presence of metals in soil (Deng and Tabatabai, 1994). An enzymatic approach to glucose measurements, however, is accurate, fast and simple (Raba and Mottola, 1995). The reagents necessary for enzymatic glucose measurements are commercially available as package kits.
The purpose of this work was to develop an assay to measure myrosinase activity in soils previously cropped with Brassica or Sinapis species. The assay is based on measuring the concentration of glucose released upon the enzymatic hydrolysis of sinigrin in soils.
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MATERIALS AND METHODS
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Soil Preparation
Soils used (Table 1) were selected to obtain a range of texture, pH, and organic matter. Before use, soils were grown with Sinapis alba (cv. Ida Gold) for 2 mo in a greenhouse pot experiment to establish myrosinase activity in soils. Plants were removed from pots and excess soil on roots was removed by shaking. Soil adhering to roots was then collected as rhizosphere soil to be used in this study. A portion of each soil was maintained at field moisture condition and the remainder was left on the laboratory bench for 48 h at laboratory temperature (25–28°C) to air-dry and passed through a 2-mm screen. Recognizable root residues were carefully removed from soil samples. Soil samples were stored in glass jars at 4°C for subsequent myrosinase activity measurements. A portion of each soil was saved for chemical and physical analyses.
Reagents
N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid (TES) buffer (Sigma Chemical Co., St. Louis, MO) was chosen for use in this work. Buffer solution (0.1 M) was prepared by dissolving 22.92 g of TES in approximately 1 L of doubled deionized water, the pH was adjusted to the desired level by titration with NaOH (1 M), and then the final volume was brought to 1 L. The glucosinolate used as a substrate was sinigrin (Sigma Chemical Co., St. Louis, MO). Toluene was obtained as a Fisher certified reagent (Fisher Scientific Co., Chicago, IL). D-Glucose was used to construct a standard curve and to study glucose recovery from soil. A glucose diagnostic kit (Trinder, Catalog Number 315-100) (Sigma Chemical Co., St. Louis, MO) was used to measure the amount of glucose released upon enzymatic hydrolysis of sinigrin. The glucose diagnostic kit was reconstituted according to the manufacturer's specification. The reagent was stored at 4°C and was stable for 3 mo.
Standard Procedure
Place 1 g of soil in a 50-mL plastic centrifuge tube, add 0.2 mL of toluene, 2.3 mL of TES buffer (0.1 M, pH 7), and 0.5 mL of sinigrin prepared in 0.1 M TES buffer (pH 7) to obtain a final concentration of 20 mM. Swirl the tube for a few seconds to mix the contents. Stopper the tube and incubate it at 37°C. After 4 h, centrifuge the soil suspension at 8000 x g for 10 min and filter the supernatant through a 0.45 µm MF-Millipore membrane filter (Millipore Corp., Bedford, MA) into a test tube. During the centrifugation period, pipette 2 mL of reagents from the diagnostic glucose kit to a labeled test tube and allow it to warm to assay temperature. Add 1 mL of the filtered supernatant to the labeled test tube containing the 2 mL of reagents from the diagnostic kit and mix by gentle inversion. Incubate the tube at room temperature (20 to 25°C) for exactly 20 min and then add immediately 25 µL of AgNO3 (1 M) to stop activity of all enzymes. Measure the absorbance of the pink color of the quinoneimine complex formed with a spectrophotometer adjusted to a wavelength of 505 nm. Calculate the concentration of glucose by referring values of absorbance to a standard curve. To establish a standard curve, prepare a glucose stock solution of 1000 µg glucose mL-1. Then pipette 0, 50, 100, 200, 400, and 600 µL of glucose stock solution to a series of labeled test tubes and add doubled deionized water to obtain a total volume of 3 mL in each tube. Pipette 1 mL from each tube to 2 mL of reagents from the diagnostic kit and proceed as described for analysis of glucose in the soil filtrate.
Proper controls must be performed in each series of analysis to measure glucose background in soil. To perform controls, add 2.3 mL of TES buffer (pH 7) and 0.2 mL of toluene to 1 g of soil and incubate for 4 h. After incubation, add 0.5 mL of sinigrin to obtain a final concentration of 20 mM and proceed as described above for the standard procedure.
Glucose Recovery
The efficiency of glucose recovery from soil was tested by adding 3 mL of varying glucose concentrations prepared in TES (pH 7) and containing 50 to 1000 µg glucose to 1 g of different soils. Toluene (0.2 mL) was added to each sample and controls (without toluene) were performed to examine the effect of toluene on glucose recovery. Samples were incubated at 37°C for 4 h. Glucose was extracted and measured as described in the standard procedure.
Assay Conditions
Myrosinase activity in field-moist vs. air-dry soil.
Using the standard assay procedure, myrosinase activity was measured in field-moist soil immediately after sampling and in air-dried soil. Myrosinase activity in both soils was calculated based on oven-dry weight.
Effect of pH.
To study the effect of pH on myrosinase activity in soils, TES buffer was adjusted to range from pH 5.5 to 9.0. Buffer (2.3 mL) adjusted to various pH values, toluene (0.2 mL), and sinigrin (0.5 mL), prepared in the buffer at the same pH and in an amount needed to obtain a final concentration of 20 mM, were added to 1 g of soil. Soils were incubated for 4 h and myrosinase activity was measured as described in the standard procedure.
Effect of soil amount.
Myrosinase activity was measured in different amounts of soils (0.5, 1.0, 1.5 g soil) using the standard procedure.
Effect of incubation time.
Myrosinase activity was measured using the standard assay procedure but incubated for a time course ranging from 1 to 8 h.
Thermodynamic and Kinetic Determinations
Thermodynamic determinations.
The effect of temperature on myrosinase activity was examined by equilibrating soil, buffered with TES (pH 7) and treated with toluene as described in the standard assay, at various temperatures (10–70°C) for 10 min. The reaction was then initiated by the addition of sinigrin (final concentration 20 mM) and incubation temperatures were maintained at 10 to 70°C for 4 h. Myrosinase activity was measured as described in the standard procedure. The activation energy values for the enzymatic hydrolysis reactions of sinigrin in soils were determined based on the linear form of the Arrhenius equation:
where k is the rate constant for the reaction, A is the frequency factor, Ea is the activation energy, R is the gas constant, and T is the temperature in the Kelvin scale (Segel, 1975).
Ea values were determined by plotting the log of myrosinase activity, expressed as micrograms of glucose per gram of soil per 4 h (µg glucose g-1 soil 4 h-1), versus T-1. The activity values can be used as they are directly proportional to the rate constant (k). Assay temperatures used were 10, 20, 30, and 40°C, and were converted to the Kelvin scale prior to plotting.
Kinetic determination.
The Km and Vmax values of myrosinase in soil were determined by measuring myrosinase activity (µg glucose g-1 soil 4 h-1) using a suitable range of sinigrin concentrations (5–50 mM). Enzyme activities were plotted against the corresponding sinigrin concentrations according to the three linear transformations of the Michaelis–Menten equation: Lineweaver–Burk plot (1/v vs. 1/S), Eadie–Hofstee plot (v vs. v/S), and Hanes–Woolf plot (S/v vs. S) (Hofstee, 1952).
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RESULTS AND DISCUSSION
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Assessment of myrosinase activity is based on measuring glucose released when soil is incubated with the substrate sinigrin. The extraction efficiency of glucose from soil, the activity of enzyme in air-dried versus fresh soil, and the effect of buffer pH, amount of soil, time of incubation, and temperature were all evaluated to optimize and characterize myrosinase activity in soil. Unless indicated otherwise, all results presented were obtained using the standard assay.
Extraction and Estimation of Glucose
Glucose is water-soluble and can be readily extracted from soil with water. Extraction efficiencies were always 100 ± 0.9% when glucose was added to soil and then immediately extracted (data not shown). However, when soils amended with different glucose concentrations were incubated for 4 h, extraction efficiency was greatly reduced to as low as 5.8% as a result of microbial utilization (Table 2). As such, it was necessary to use a biostatic reagent to suppress microbial utilization of glucose. Toluene was used for this purpose and it was found to increase glucose recovery to a range of 91.9 to 99.7% (Table 2). Tests showed that toluene had no effect on the analytical procedure used to measure glucose concentrations or on myrosinase activity. Therefore, toluene (0.2 mL) was added as a part of the standard myrosinase activity assay.
Concentrations of glucose extracted from soil were measured using the glucose diagnostic kit. Tests indicated that more than 80% of color intensity was formed in the first 5 min and the color development was essentially complete after 10 min. A standard period of 20 min was chosen for color development and silver nitrate was then immediately added after this period to inactivate any further reaction that may occur and affect the color intensity. We found addition of silver nitrate makes the color stable for more than 1 h after color development was completed.
Fresh vs. Air-Dried Soil
Enzyme activities in soils are often affected by air-drying (Tabatabai, 1994). We tested myrosinase activity in field-moist versus air-dried soils. Use of air-dried soils is desirable because they are much easier to handle and store. For all soils used in this work, myrosinase activities were slightly higher in air-dried versus field-moist soils (Table 3). Therefore, myrosinase can be assayed in air-dried soil without loosing activity. It is important that soils are not dried in an oven prior to myrosinase activity measurements, as this will inactivate a large amount of the total activity.
Buffer and pH
Several buffers were tested in this study including phosphate, TES buffer, tris(hydroxymethyl)aminomethane (THAM), and 4-morpholine-ethanesulfonic acid (MES). Phosphate buffer, and to some extent THAM buffer, caused extraction of dark-colored organic material from soil that interfered with the spectrophotometric measurements of glucose. The MES seemed to be a weak buffer and was not able to maintain soil pH close to the desired pH. TES, however, was able to maintain soil pH very close to the desired pH (i.e., ±0.08 pH units) and did not extract any color from soil. Tests also indicated that TES did not interfere with the analysis of glucose when using the diagnostic kit.
The effect of buffer pH on activity of myrosinase is shown in Fig. 1
. Results indicated that myrosinase exhibited the highest activity at a buffer pH of 7.0 and greatly decreased when pH was higher or lower than this point in all soils used in this work. Ohtsuru and Kawatani (1979), Iori et al. (1996), and Sharma and Garg (1996) obtained similar results when studying myrosinase purified from Brassica species. However, other studies reported different pH optima for myrosinase. Yen and Wei (1993) reported the optimum pH of myrosinase purified from cabbages (Brassica oleracea) was 8.0. Bjorkman and Janson (1972) showed the optimum pH values for the white mustard and rapeseed myrosinases ranged from pH 4.5 to 4.9.
Temperature and Energy of Activation
Figure 2
shows the temperature dependency of myrosinase activity in soils. Myrosinase activity increased with increasing temperature up to 40°C, and started to decline above 40°C. These results are indicative of an enzyme-catalyzed reaction and are similar to those obtained by Sharma and Garg (1996) for myrosinase purified from Brassica juncea. Springett and Adams (1989) showed that optimum temperature for myrosinase extracted from Brassica oleracea was 50°C and the inactivation temperature was approximately 60°C.
The Arrhenius plots of myrosinase activity values in soils studied, used to calculate Ea, were linear between 10 and 40°C (Fig. 3) . Slopes obtained from the Arrhenius plots are almost identical and thus Ea values for myrosinase in these soils were similar. Activation energy of reactions catalyzed by myrosinase in the four soils, expressed in kilojoules per mole (kJ mol-1), ranged from 40.3 for the Wooster soil to 52.8 for the Luray soil.
Amount of Soil and Incubation Time
The amount of glucose formed increased linearly with increasing amounts of soil (Fig. 4)
. The linear relationship obtained is evidence that the method developed reflects concentration of myrosinase in soil. The fact that no activity was observed when soil was not present also indicates that the glucose extracted from soil was due to enzymatic action and not to substrate degradation.
The effect of incubation time on myrosinase activity is shown in Fig. 5
. Myrosinase activity was linear with time for 8 h in all soils used in this work, showing that the assay of soil myrosinase activity by this method is not affected by microbial assimilation of enzymatic reaction products. This linearity indicates also that glucose formation was a zero-order reaction for at least 8 h. However, further incubation caused the reaction rate to deviate from linearity suggesting either that the reaction became substrate limiting, products inhibiting the reaction catalyzed by myrosinase were accumulating, or the product was being utilized by microorganisms. To minimize errors that may occur because of these potential problems, a 4-h incubation period was selected. This both reduces potential problems and ensures there is enough time to produce detectable glucose levels in soils with low myrosinase activity.
Substrate Concentration and Kinetic Constants
When assaying enzymes, the substrate concentration must be sufficient to maintain zero-order kinetics, thus achieving a reaction rate proportional to enzyme concentration (Segel, 1975). Zero-order reaction kinetics were achieved at a concentration of 20 mM sinigrin for all soils used in this study. This concentration is equivalent to 8.3 mg of sinigrin per milliliter of soil solution. Borek et al. (1996) found that 10 mg of sinigrin per milliliter of soil extract was required to ensure that the reaction was not substrate limited when measuring myrosinase activity in soil extracts.
Determinations of the Michaelis–Menten kinetic constant (Km) and the maximum rate of reaction (Vmax) are of interest for several reasons. The apparent Km is comprised of a number of rate constants and can be used to rapidly predict the substrate concentration at which the reaction rate is independent of the substrate concentration. Vmax is a good indicator of enzyme concentration per volume or mass unit and, thus, its numerical value provides a means of comparing enzyme content in different soils. Km and Vmax are constant for a specific enzyme and enzyme concentration but they may vary independently of each other under different conditions (Dick and Tabatabai, 1984). The Km and Vmax values of myrosinase activity in soils were determined using the three possible linear transformations of the Michaelis–Menten equation (Fig. 6)
. The straight lines are those calculated by regression analysis with R2 > 0.90 for all soils. All results obtained followed the three linear transformations of the Michaelis–Menten equation.
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Fig. 6. The three possible linear transformation plots of the Michaelis–Menten equation for myrosinase activity in soils. The reaction velocity (v) is expressed as micrograms of glucose per gram of soil per 4 h (µg glucose g-1 soil 4 h-1) and the substrate concentration [S] is in molarity (M).
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The Km and Vmax values calculated from linear plots for Michaelis–Menten equation are shown in Table 4. The apparent Km values of sinigrin for myrosinase in four soils ranged from 5.3 to 12.9 (avg. 8.1) mM and the apparent Vmax values ranged from 76 to 518 (avg. 275) µg glucose g-1 soil 4 h-1. The average Km value for sinigrin obtained in this study is similar to that obtained by Leoni et al. (2000) who found that Km for myrosinase immobilized on granular nylon was 6 mM. However, these Km values are about ten times greater than that shown for myrosinase purified from Brassica species, which ranged from 0.16 to 0.51 mM (Palmieri et al., 1982; Shikita et al., 1999). The higher Km values of myrosinase activity in soil compared with that for myrosinase purified from Brassica is presumably because of the sorption of sinigrin by soil constituents causing the effective concentration of substrate reaching the enzyme to be lower. It is also possible that some sort of competitive inhibition of myrosinase may occur in soils. The Km values for several enzymes have been shown to be higher in soil compared with purified enzymes (Tabatabai and Bremner, 1971; Tabatabai and Singh, 1979; Dick and Tabatabai, 1978).
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Table 4. Km and Vmax values of myrosinase in soils calculated from the three linear transformations of the Michaelis–Menten equation.
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Precision
The precision of the method for the five soils that have widely different characteristics is shown in Table 5. The reaction rate catalyzed by myrosinase ranged from 71 in the Spinks soil to 338 µg glucose g-1 soil 4 h-1 in the Wooster soil and the standard deviation of the activity values ranged from 4.2 to 13.0 (avg. 8.3) µg glucose g-1 soil 4 h-1. The coefficient of variation for the myrosinase assay in these five soils ranged from 2.4 to 7.3%. Since the main source of myrosinase in soils we studied was derived from plant roots, it was important to remove all recognizable root remnants from soils sampled from the plant rhizosphere and maximize the homogeneity of the enzyme in the soil sample by vigorous mixing. This reduced the variability between replicates.
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CONCLUSION
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Myrosinase is present in soil where there is growth of Brassica and Sinapis plants. Hydrolysis of glucosinolate (sinigrin) produces detectable amounts of D-glucose that can be colorimetrically measured and used as a quantitative measure of myrosinase activity in soil. The optimum condition for myrosinase assay in soils is achieved at pH 7, 37°C, and a final sinigrin concentration of 20 mM. The average apparent Km value for myrosinase activity in soil is 8.1 mM. Our method is accurate, fast, and can be used in most laboratories.
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ACKNOWLEDGMENTS
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This research was supported in part by the Royal Embassy of Saudi Arabia-Cultural Mission, Washington, DC, and by state and federal funds appropriated to The Ohio State University.
Received for publication February 4, 2002.
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ABSTRACT
INTRODUCTION
