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Organophosphorus pesticides have been used in large quantities throughout the world since the first introduction of a synthetic insecticide, parathion, for use in crop protection in 1944 (16). Problems of contamination resulting from surplus pesticides and wastewater from pesticide factories have become obvious. The transformation of pesticides in the environment results from physicochemical reactions as well as from the activity of cellular or extracellular components of the biota (microorganisms, plants, and animals), but the principal biological pathway is microbial degradation (7): microorganisms can metabolize various pesticides both in soil and in water. The earlier metabolic studies on pesticides helped to develop a new approach to the detoxification of pesticides using cell-free enzymes from adapted microorganisms to resolve problems related to whole-cell metabolism of pesticides. Munnecke purified an enzyme from a mixed bacterial culture grown on parathion and found that it hydrolyzed parathion and related organophosphorus insecticides (13). Mulbry and Karns purified three unique parathion hydrolases from gram-negative bacterial isolates. These enzymes with different characteristics were exploited in different types of waste disposal (12). Although there have been a number of reports on the metabolism of parathion by enzyme preparations from bacteria or by mixed-culture bacteria (12-14), there have been no successful cell-free studies with fungi capable of degrading organophosphorus insecticides. To our knowledge, this is the first dimethoate-degrading enzyme purified to homogeneity from a fungus. Characterization and comparison of the degrading enzymes from different microorganisms may help us to understand fully how these enzymatic activities may evolve and how organophosphorus insecticides are degraded in the ecosystem.

Microorganism and enzyme production. All experiments were performed in triplicate, and results, where appropriate, are presented as means. Aspergillus niger ZHY256 was isolated from sewage and the soil of cotton fields where pesticides were heavily used and maintained on potato dextrose agar. For enzyme production, the fungus was grown on basal salt medium (0.2% NaNO₃, 0.05% KCl, 0.05% MgSO₄ · 7H₂O, 0.02% BaCl₂, 0.01% MnSO₄, 0.005% CaCl₂, pH 6.8, supplemented with dimethoate at a concentration of 0.25% [wt/vol]). Dimethoate-degrading enzyme activity was detected at the late log phase and reached the maximum level 5 days after the start of cultivation. It was found that Ba²⁺ greatly stimulated the enzyme production with an optimum concentration of 1 mM in the medium. The addition of Ba²⁺ to the cell extract cultured without Ba²⁺ did not accomplish any acceleration of the enzyme activity, showing that Ba²⁺ is needed to produce the enzyme or to stabilize the enzyme produced. The effects of various carbon sources on dimethoate-degrading enzyme production by A. niger ZHY256 were investigated (Table 1). The microorganism grew on every substrate tested, although the mycelial mass yield was about four times lower on dimethoate- or malathion-containing media than on dextrose- or sucrose-containing media. Differences in growth yield were, however, not related to differences in total or relative enzyme production. The greatest amount of dimethoate-degrading enzyme activity was from cell extract grown in dimethoate-containing basal salt medium.

Enzyme purification. All experiments described below were carried out between 0 and 4°C unless otherwise specified. A. niger ZHY256 was incubated at 30°C for 5 days in 500-ml Erlenmeyer flasks containing 150 ml of medium on a rotary shaker at 150 rpm and harvested by centrifugation at 12,000 × g for 20 min. The collected mycelia were washed twice with cold 50 mM Tris-HCl buffer (pH 7.0) and disrupted (8 ml of buffer/g of wet mycelia) in a vibration homogenizer (Vibrogen vi4; Edmund, Tuebingen, Germany) with glass beads (0.1-mm diameter). After standing at 4°C overnight, the suspension was centrifuged (12,000 × g for 20 min at 4°C) to remove the unbroken cells and cellular debris. The supernatant was filtered through a 0.22-µm-pore-size membrane (Millipore), the filtrate being the crude extract of the enzyme. The protein of the crude extract was precipitated overnight with 80% (NH₄)₂SO₄. The resulting precipitate was collected by centrifugation at 12,000 × g for 20 min, dissolved in the smallest possible volume of 50 mM Tris-HCl buffer (pH 7.0), dialyzed 1,000-fold against the same buffer, and concentrated by ultrafiltration. The concentrated enzyme solution was loaded onto a Sephadex G-100 column (1.8 by 100 cm) preequilibrated with 50 mM Tris-HCl buffer (pH 7.0). The column was washed at a flow rate of 24 ml/h with 400 ml of the same buffer, and 5-ml fractions were collected. The molecular weight of the native enzyme was determined by the method of Andrew by using blue dextran and molecular weight markers from Sigma as standards (2). Proteins were eluted in fractions 18 to 80, whereas the enzyme was confined to fractions 26 to 38. The fractions with high specific activity were then pooled and concentrated for further purification. Fractions with dimethoate-degrading enzyme activity were loaded on a DEAE-Sepharose CL-6B ion-exchange column (1.2 by 30 cm) preequilibrated with 50 mM Tris-HCl buffer (pH 7.0). The column was washed at a flow rate of 20 ml/h with 500 ml of the same buffer, and proteins were eluted with a linear gradient of NaCl from 0 to 1.0 M. Five-milliliter fractions were collected. This enzyme solution was the purified dimethoate-degrading enzyme preparation used for subsequent characteristic studies. The data for the purification are summarized in Table 2. The enzyme was purified 36.1-fold to a specific activity of 227.6 U/mg of protein from the mycelia with a yield of 33.48%.

Enzyme assay. Dimethoate-degrading enzyme activity was assayed with 50 mM dimethoate as a substrate. The reaction was carried out at 50°C in a final volume of 1 ml of 50 mM Tris-HCl buffer (pH 7.0) and was stopped by adding 1 ml of 2 M trichloroacetic acid, and the amounts of all organophosphorus insecticides tested in the experiments were determined by a gas chromatograph (model PE Sigma 2000) with a glass column (1-m length, 2-mm inner diameter) packed with 5% OV-17 on Chromosorb WDMCS, operated at 220°C, with the detector at 250°C and the injector at 230°C (3, 5, 18). One unit of activity was defined as the amount of enzyme that catalyzes the degradation of 1 µmol of dimethoate per min under these conditions. The protein concentration was determined according to the method of Lowry et al., with bovine serum albumin as the standard (11).

Enzyme characterization. The purified enzyme gave a single band in sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) by the method of Laemmli (8). This indicated that the purified sample was electrophoretically homogeneous under the dissociating conditions (Fig. 1). The molecular mass of the purified enzyme was estimated to be approximately 66 kDa by gel filtration on a calibrated column of Sephadex G-100, which was quite close to the value determined for parathion hydrolase of strain SC (12). The value was confirmed by SDS-PAGE (Fig. 1). Hence, it is assumed that the native dimethoate-degrading enzyme is a monomer.

View larger version (28K): [in this window] [in a new window]   FIG. 1.   SDS-PAGE of the purified dimethoate-degrading enzyme from A. niger ZHY256. Lane 1, marker proteins (from top to bottom) phosphorylase b (Mr, 94,000), bovine serum albumin (Mr, 67,000), ovalbumin (Mr, 43,000), carbonic anhydrase (Mr, 30,000), and -lactalbumin (Mr, 14,400), respectively; lane 2, purified enzyme. The gel was stained for protein with Coomassie brilliant blue R-250 and destained in methanol-acetic acid-water (7:6:47).

View larger version (13K): [in this window] [in a new window]   FIG. 2.   Effects of temperature and pH on activity () and stability () of the purified enzyme.

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