Environ. Sci. Technol. 1997, 31, 2392-2398
Enzymatic Coupling of the Herbicide Bentazon with Humus Monomers and Characterization of Reaction Products JANG-EOK KIM,† ERROL FERNANDES, AND JEAN-MARC BOLLAG* Laboratory of Soil Biochemistry, Center for Bioremediation and Detoxification, 129 Land and Water Building, The Pennsylvania State University, University Park, Pennsylvania 16802
To elucidate the binding mechanism of the herbicide bentazon (3-isopropyl-1H-2,1,3-benzothiadiazine-4(3H)-one 2,2dioxide) with humic monomers in the presence of an oxidative enzyme, the reaction of bentazon with catechol, caffeic acid, protocatechuic acid, and syringaldehyde was investigated. In the presence of a laccase from the fungus Polyporus pinsitus, catechol was the most reactive humic monomer; bentazon with catechol in the presence of the laccase was completely transformed in 30 min at pH 4.0. The reactivity of bentazon decreased with increasing pH, but reactivity of bentazon decreased with increasing pH, but complete transformation of bentazon could be achieved even at high pH if the concentration of catechol was increased. When bentazon was incubated with humic acid (extract of leonardite) in the presence of the laccase, a reaction of the two substrates was observed, as indicated by bentazon disappearance. Two metabolites that result from the reaction of bentazon with catechol were isolated by TLC and HPLC and identified by mass spectrometry and NMR spectroscopy. A product with a molecular weight of 348 was characterized by 1-D, 2-D 1H-, and 13C-NMR spectroscopy and identified as a dimer composed of one catechol and one bentazon molecule. A second reaction product with a molecular weight of 586 appeared to be a trimer, consisting of one catechol molecule and two bentazon molecules. The analyses also showed that catechol was bound to the protonated nitrogen of the heterocyclic ring and not to a carbon of the aromatic ring of bentazon; this incorporation results from nucleophilic addition of the o-quinone to the nitrogen.
Introduction The fate of pesticides and other xenobiotics in the environment is important because they create a potential biohazard, accumulating in soil and possibly contaminating groundwater. Most of the pesticides that are sprayed in the plantsoil agricultural system enter the soil and become bound to soil constituents by physical or chemical forces of varying strength (1, 2). Therefore, the elucidation of the linkage to soil constituents is important for assessing the fate of pesticides in the environment. * Corresponding author: phone: (814)863-0843, fax: (814)8657836; E-mail:
[email protected]. † Present address: Department of Agricultural Chemistry, Kyungpook National University, Taegu, 702-701, Korea.
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Kim et al. (3) showed that the herbicide bentazon can be transformed by oxidative coupling reactions in the presence of humic substances. These results indicate that bentazon in soil can be incorporated within humic substances by oxidative enzymes. Since incorporation generally renders the xenobiotics unavailable for plants and animals (4, 5), it can be considered a detoxification process (6-8). This method of detoxification through incorporation can be considered an alternative method of pollution control. However, it is difficult to characterize the detoxification mechanism because humic substances are complex and heterogeneous, as are the reaction products generated (9, 10). In addition, interaction among the various compounds and their breakdown products increases the complexity of the soil environment. To circumvent these problems in the present study, we designed model experiments in which bentazon was incubated with simple humus constituents. In order to elucidate the mechanism of this coupling, model experiments were designed in which xenobiotics were incubated with simple humus constituents that would define the types of chemical bonds that occur. To date, studies on the fate of bentazon in the soil environment have shown that it can be quickly transformed through hydroxylation at the 6 or 8 position on the phenyl ring and through the formation of 2-amino-N-isopropyl benzamide (AIBA) under aerobic conditions (15, 16). Therefore, the primary path of transformation lies in the incorporation of the three reactive metabolites, 6-hydroxy bentazon, 8-hydroxy bentazon, and AIBA, into the organic soil matrix. However, the nature of the bonding between bentazon and humic substances still remains unclear. This investigation was initiated to elucidate the reactivity, binding mechanism, and chemical structure of reaction products arising from incubation of the herbicide bentazon with humic monomers in the presence of the oxidative enzyme, namely, laccase. An additional objective was to add to present knowledge concerning the binding mechanisms in reactions catalyzed by oxidoreductases (3, 11-14).
Materials and Methods Chemicals. Unlabeled and 14C-labeled bentazon were obtained from BASF AG (Germany). Radiolabeled bentazon was uniformly labeled in the aromatic ring with a specific radioactivity of 5.21 MBq/mg. Catechol was purchased from Fisher Scientific Co. (Fair Lawn, NJ), caffeic acid was from Aldrich Chemical Co. (Milwaukee, WI), protocatechuic acid was from Sigma Chemnical Co. (St. Louis, MO), and syringaldehyde was from Fluka AG (Buchs, Switzerland). Humic acid was extracted from leonardite provided by Humus Products of America (Richmond, TX). The extraction procedure was based on the method of Stevenson (17). The laccase from the fungus Polyporus pinsitus was obtained from Novo Nordisk Bioindustrials, Inc. (Danbury, CT), and its activity was determined spectrophotometrically by using 2,6dimethoxyphenol as a substrate. One unit was defined as the amount that caused a change in optical density of 1.0/ min at 468 nm. Enzymatic Reaction Conditions. Unless otherwise specified, enzymatic reactions were conducted at 25 °C for 4 h in 5 mL of 0.1 M citrate-phosphate buffer containing 4 units/ mL laccase with 1 mM bentazon and 1 mM humic monomer. The enzymatic reaction was stopped by adjusting the reaction mixture to pH 2.0 with acetic acid; subsequently, the reaction mixtures were centrifuged at 12000g for 10 min to remove the precipitates. The supernatants were filtered through a 0.45-
S0013-936X(96)01016-4 CCC: $14.00
1997 American Chemical Society
TABLE 1. Transformation of Bentazon by a Laccase in Presence of Humus Monomers as a Function of pHa bentazon transformed in % monomers
pH 3.0
pH 4.0
pH 5.0
pH 6.0
pH 7.0
catechol caffeic acid protocatechuic acid syringaldehyde
100 76 14 38
100 59 40 38
82 29 12 31
17 7 3 2
5 5 4 2
a
Incubation time, 4h.
FIGURE 2. Effect of catechol concentration on the transformation of bentazon by a laccase of Polyporus pinsitus at pH 4 and pH 6 (incubation time, 4 h).
FIGURE 1. Time course for transformation of bentazon at pH 4 and pH 6 by a laccase of Polyporus pinsitus in the presence of catechol. µm nylon membrane, and the residual bentazon was quantified by HPLC. Boiled enzyme was used in the control sample. For product isolation, the reaction conditions were the same as described above, but the reaction volume was scaled up to 100 mL and included 65 000 dpm/mL of [14C]bentazon. After incubation for 4 h, the reaction mixture was extracted three times with an equal volume of methylene chloride. The extract was dehydrated by anhydrous sodium sulfate and evaporated to 1-2 mL by rotary evaporation at 35 °C and then dried under a stream of N2. Radiocounting. To determine the extraction efficiency of the solvent, radiocounting was performed using an Analytic Beta Trac 6895 scintillation counter (Elk Grove, IL), with Ecoscint A (National Diagnostics, Manville, NJ) as the scintillation cocktail. Thin-Layer Chromatography (TLC). The residues obtained were re-dissolved in methanol and separated by TLC using silica gel 60 F-254 plates (EM SCIENCE, Germany) with a mobile phase mixture of chloroform:ethyl acetate:methanol (50:25:25, v/v/v). The separated compounds were examined for radioactivity with a Bioscan Imaging 200 scanner (Bioscan, Washington, DC). Radioactive bands determined by the scanner were scraped from the TLC plate and extracted with methanol. After centrifugation of isolated samples, they were filtered through a Sep-Pak C18 cartridge (Millipore Corp., Milford, MA), and the eluent was further purified by HPLC. High-Performance Liquid Chromatography (HPLC). The disappearance of bentazon and the formation of reaction products were monitored by a Waters Associates HPLC system (Milford, MA) equipped with a Model 6000A and a Model 501 solvent delivery system using a 4.0 mm × 25 cm reversephase column (Nucleosil C18, 5 µm, Macherey Nagel) with UV detection at 254 nm. The mobile phase set at a flow rate of 0.8 mL/min was a mixture of an aqueous component [A] (1% acetic acid in Milli-Q water) and [B] methanol. Isocratic conditions were used for the analysis of bentazon with a mobile phase ratio of 50/50 (A/B). The reaction products were isolated by a gradient elution in which the initial mobile-
FIGURE 3. Effect of humic acid concentration on the transformation of bentazon in the presence of a laccase of Polyporus pinsitus (incubation time, 4 h). phase composition of 70/30 (A/B) was changed to 35/65 (A/ B) over 18 min. This ratio was then maintained for 5 min. Next, the solvent ratio was taken from 35/65 (A/B) to 0/100 (A/B) in 7 min, and this final composition was held for 10 min. The column was equilibrated at initial conditions for 10 min prior to each injection. The eluted peaks were collected, the combined fractions were diluted with distilled water, and the resulting solution was extracted twice with methylene chloride. The solvent extract was dehydrated and evaporated as described above. Mass Spectrometry. Molecular weights of the reaction products obtained following the oxidation of the bentazon and humic monomers were determined by electron ionization (70 eV) mass analysis. A Kratos MS 9/50 double-focusing mass spectrometer was used for the analysis, and sample introduction was by direct insertion probe with the source temperature between 250 and 350 °C. NMR Spectroscopy. 1H- and 13C-NMR spectroscopy were used to elucidate the structure of the reaction products. 1-D and 2-D experiments were carried out using high-resolution NMR spectrometers operating in the quadrature mode. All of the 1-D NMR experiments were performed on a Bruker AM-500 spectrometer except for the 13C analysis of the bentazon standard, for which a Bruker WM-360 spectrometer was used. The 13C-NMR measurements were made with continuous WALTZ decoupling (18) of the protons.
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FIGURE 4. HPLC analysis of the reaction products of bentazon and catechol after incubation with a laccase of Polyporus pinsitus. Two-dimensional proton-proton shift correlation spectroscopy (double quantum filtered COSY, Bruker package) was employed to observe the coupling of neighboring protons. For the analysis of standard bentazon and the products, a 2-D C-H correlation experimentsvia double quantum coherence using BIRD sequence in inverse mode and 13C decoupling during acquisition using GARP-1 (19)swas carried out in each case to observe the connectivities between protons and carbons. For all the 2-D experiments, a Bruker AM-300 spectrometer was employed. Regular NMR tubes (5 mm o.d.) were used for the samples. Deuterated methanol was used to dissolve the samples and provide a 2H lock for NMR; except in the case of the 13C study of bentazon, deuterated chloroform was used as the lock solvent. To simplify the discussion of structure elucidation by NMR spectroscopy and to facilitate reference to specific carbons and protons, a different (non-IUPAC) numbering scheme was applied for the structures of bentazon and reaction product A.
Results The transformation of bentazon by a laccase of Polyporus pinsitus in the presence of various humic monomers at various pH values was investigated (Table 1). Bentazon was completely transformed by a laccase in the presence of a catechol at pH 3.0 and pH 4.0, but by increasing the pH to 7, the transformation rate was reduced. Similar reductions in transformation with increasing pH were observed for caffeic acid, protocatechuic acid, and syringaldehyde. Catechol was the most reactive monomer for the transformation of bentazon in the presence of the laccase at pH 4.0, followed by caffeic acid, protocatechuic acid, and syringaldehyde, in decreasing order of transformation. The time course for the reaction of bentazon with catechol by the laccase at two different pH values is shown in Figure 1. The transformation of bentazon was complete in 30 min of incubation at pH 4.0, but had reached only 17% after 4 h at pH 6.0, with no additional transformation thereafter. Figure 2 shows the effect of catechol concentration on the transformation of bentazon by the laccase at pH 4.0 and pH 6.0. At pH 4.0, the removal of bentazon was optimized at a ratio of 0.6 mM/1 mM catechol/bentazon. At pH 6.0, the reaction did not proceed until the ratio was greater than 0.8 mM/1 mM; bentazon was completely removed at a catechol/ bentazon ratio of 5 mM/1 mM.
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The transformation of bentazon by the laccase in the presence of humic acid is shown in Figure 3. Bentazon removal was enhanced as the concentration of humic acid was increased and reached 26% at a humic acid concentration of 2%. To isolate products, a reaction mixture containing [14C]bentazon was extracted by methylene chloride. The extraction efficiency of radioactivity by the solvent was confirmed as 99% as determined by liquid scintillation counting. Through the use of a radioscanner for detection of components separated by TLC, a strongly radioactive band of products from the reaction of bentazon, catechol, and the laccase was detected at rf 0.83. This band was subsequently analyzed by HPLC and was found to contain three components: unreacted bentazon (retention time, 18 min), product A (retention time, 22 min), and product B (retention time, 30 min) (Figure 4). Mass spectrometry analysis of products A and B gave molecular weights of 348 and 586, respectively. 1H- and 13C-NMR Spectroscopy of Bentazon. The aromatic region of the NMR spectrum of bentazon (6.4-8.2 ppm; Figure 5, I) shows four protons of equal intensity (after integration) in the form of a doublet-triplet-triplet-doublet sequence. The 2-D COSY spectrum for the bentazon standard clearly indicated the correlations between these downfield protons (data not shown). The doublet at 8.07 ppm (position 3) is coupled to the proton at 7.30 ppm (position 4), which in turn is coupled to the triplet at 7.62 ppm (position 5); finally, the proton at 7.62 ppm (position 5) is coupled to the doublet at 7.10 ppm (position 6). These proton peaks are split into multiplets due to ortho coupling with neighboring nuclei in the same system, and the values of the coupling constants (i.e., distance between the splittings, JH-H) seen in the spectra are characteristic of the same. Expansion of the downfield proton spectrum revealed further smaller splittings of the peaks, which are attributed to meta couplings between the protons. The upfield aliphatic region of the proton spectrum (0-6 ppm) showed two multiplets, which were seen directly coupled to each other in the 2-D COSY spectrum (data not shown). These peaks are assigned to the methyl protons (position 9; 1.55 ppm) and methine protons (position 8; 4.94 ppm) of the N-isopropyl group. Table 2 lists the observed peaks in the proton spectrum and their corresponding assignments. The NMR spectrum of bentazon (in CDCl3) at 90.56 MHz for 13C was obtained using a 4-ms pulse width (corresponding to a 45° 13C pulse), an acquisition time of 0.75 s, and a relaxation delay of 4 s. Table 3 shows the unambiguous
TABLE 2. Comparison of 1H Chemical Shifts of Bentazon and Compound A (ppm from TMS) 9
O 3 4 5 6
bentazon shifts (ppm) 8.07 (doublet) 7.62 (triplet) 7.30 (triplet) 7.10 (doublet)
4.94 (multiplet) 1.55 (doublet) a
2
1
8
N SO2
9
N H bentazon 7
compd A shifts (ppm)
assignment (position)
8.14 (doublet) 7.60 (triplet) 7.39 (triplet) 7.02 (doublet) 6.80 (doublet) 6.64 (m-coupling only) 6.58 (doublet m-coupling) 4.95 (multiplet) 1.47 (doublet)
03 05 04 06 12 15 11 08 09a
Methyl protons.
TABLE 3. Comparison of 13C Chemical Shifts of Bentazon and Compound A (ppm from TMS) bentazon shifts (ppm, in CDCl3)
compd A shifts (ppm, in CD3OD)
assignment (position)
162.2 (q)a
164.2 (q) 147.8 (q) 147.3 (q) 143.3 (q) 131.3 136.0 130.6 126.9 123.8 123.6 (q) 121.1 116.6 116.3 50.6 20.8
01 13 14 10 07 05 03 04 06 02 11 15 12 08 09
135.8 (q) 134.7 130.6 126.1 121.0
FIGURE 5. Proton (1H) NMR data of bentazon (I), product A (II), and product B (III) (downfield region). assignments that have been made for the carbons in bentazon. Assignments of the protonated carbons were corroborated by the 2-D C-H correlation NMR analysis (data not shown). Characterization of Product A. 1H-NMR Spectroscopy. The proton spectrum of product A (MW ) 348) was obtained using NMR conditions similar to those employed for bentazon. The plot of the downfield region (Figure 5, II) clearly shows the retention in product A of the bentazon aromatic moiety arrangement. The sequence of doublet-triplet-tripletdoublet observed for bentazon is seen here from 8.1 ppm to about 7.0 ppm. Also, the downfield region of the COSY spectrum for product A (Figure 6) reflects the same correlation between these protons as was seen in the COSY spectrum of bentazon, implying that the aromatic ring of bentazon remains unaltered after reaction with catechol. If the catechol moiety was attached to a carbon of the aromatic ring of bentazon (i.e., via a C-C or a C-O linkage), the proton spectra of bentazon and product A would clearly differ. Therefore, it must be concluded that catechol attaches at a non-aromatic site on the bentazon molecule. Additional proton peaks of approximately equal intensity seen in the product A 1H spectrum from 6.5 to 6.8 ppm (Figure 5, II) correspond to the three protons of the catechol ring after its coupling to bentazon. The protons at 6.80 ppm (position 12) and 6.58 ppm (position 11) are ortho coupled to each other, as confirmed by the COSY spectrum (Figure 6). In an expanded plot of the regular 1H spectrum, the peaks at 6.64 ppm (position 15) and at 6.58 ppm (position 11) showed additionally the small splittings indicative of meta coupling to each other.
49.2 20.7 a
q indicates quarternary carbons.
The chemistry of phenols from previous studies (20) indicates that binding of these moieties to nucleophilic sites occurs at a position para or ortho to the hydroxyl functionality. In this case, the structure of reaction product A supported by NMR spectroscopy and mass spectrometry is one in which the secondary nitrogen of bentazon binds to catechol via a position para to an OH group of catechol (Figure 7). Table 2 lists a comparison of the proton chemical shifts (in ppm from TMS) for bentazon and for product A. 13C-NMR Spectroscopy. The 13C-NMR spectrum of product A was obtained at 125.76 MHz with a 4-ms 13C pulse width and a 5-s relaxation delay. Table 3 lists the 13C chemical shifts (in ppm from TMS) for bentazon and product A, and Figure 7 presents the proposed structure for product A. The data indicate that the carbonyl group of the bentazon moiety (position 1, Figure 7) is retained and that a protonated carbon of the catechol moiety is lost. The quaternary carbon at 143.3 ppm is assigned to a binding site on the catechol unit (position 10) by which it links to bentazon to form product A. The peaks at 147.3 and 147.8 ppm for product A correspond to the carbons directly attached to the hydroxyl groups (positions 13 and 14 in Figure 7) of the catechol molecule. The small difference in the chemical shift of these carbons can be explained by the substitution of catechol para to one of the hydroxyl groups that has resulted in making the two carbon atoms attached to the two hydroxyl groups non-
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FIGURE 6. Two-dimensional proton correlation spectrum of product A (downfield region).
FIGURE 7. Proposed pathway of the reaction between bentazon and catechol generated by a laccase of Polyporus pinsitus. equivalent. The 13C-NMR findings corroborate the results of the proton NMR studies and support the structure proposed for product A. Characterization of Product B. 1H-and 13C-NMR Spectroscopy. The proton spectrum of product B (MW ) 586) was obtained using conditions identical to those used for bentazon. The downfield region of the 1H-NMR plot (Figure 5, III) reveals an aromatic proton shift sequence similar to that observed for bentazon and product A, again implying that the aromatic ring on bentazon remains unaffected by the reaction. The chemical shifts of the doublet-triplet-tripletdoublet (of equal intensity) arising due to these protons are 8.09, 7.61, 7.33, and 7.09 ppm, respectively. An additional peak of approximately the same intensity is seen at 6.90 ppm
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and arises from catechol ring protons. This peak was not seen in the COSY spectrum , implying that it must originate from protons that are para to each other, or if the protons are ortho to each other, they must be equivalent because they have the same chemical shift. Exact assignments could not be made for all carbons of product B because of its very low concentration, rendering a 2-D C-H correlation experiment impractical. Although there are small impurities present, the regular 13C-NMR spectrum of this product clearly showed the disappearance of two protonated carbons of catechol, implying substitution at two sites on catechol. Also, both the quaternary carbons attached to the hydroxyl groups exhibited the same chemical shift (149.4 ppm), unlike those for product A (147.3 and 147.8 ppm), which strongly suggests that the substitution on catechol is symmetric. In addition, since the value of this chemical shift is close to that for product A, the location of the binding site on catechol is likely to be similar to that for product A, i.e., more likely to be para to the hydroxyl groups than ortho. The only structure that fits the NMR spectroscopy and mass spectrometry results is one in which a catechol unit binds to two bentazon units, and the binding occurs para to the hydroxyl groups of catechol (i.e., producing a symmetrically substituted catechol), as shown in Figure 7. Construction of a model of product B indicates that the two bentazon residues of this product cannot be co-planar with the catechol ring. Two conformations are possible that are cis and trans with respect to the disposition of the SO2 groups. Because of the highly polar nature of the SO2 groups, the trans arrangement is overwhelmingly favored. This conformation possesses overall chirality and since the model also indicates that the barrier to interconversion of the
enantiomer of the trans conformation will be high, the two methyl groups of each isopropyl unit are diastereotopic and should exhibit chemical shift nonequivalence. This is observed in both the 1-D and 2-D COSY proton spectra. Close examination of the methyl protons in the 1H-NMR spectrum showed them to be complex multiplets ranging from about 1.2 to 1.6 ppm. The COSY spectrum of this region indicated two slightly inequivalent sets of methyl protons separated by about 0.05 ppm. These observations and the formation of the monosubstituted derivative A, provide strong support for the proposed structure of product B shown in Figure 7.
Discussion Our previous research demonstrated that the herbicide bentazon can be incorporated into humic monomers in the presence of oxidative enzymes (3) and that in the absence of humic monomers no transformation of bentazon occurs. We studied here the incorporation of bentazon into humic substances in the presence of an oxidative enzyme, a laccase of Polyporus pinsitus, and identified the major reaction products in order to elucidate the cross-coupling mechanism. The extent of bentazon removal by the laccase in the presence of humic monomers depended on the chemical structure and concentration of the monomer, the duration of the reaction, and the pH of the reaction mixture. For example, although bentazon was transformed more efficiently in the presence of catechol at pH 4, it was transformed to some extent at pH 6 given a sufficiently long incubation time and removed completely at pH 6 given a sufficiently high catechol concentration. These findings are supported by the results of Sarkar et al. (21), Dec and Bollag (8), and Thurston (22). Sarkar et al. (21) and Dec and Bollag (8) also showed that the removal of phenols by oxidative enzymes was affected by the chemical structure and concentration of the substrates, pH, and reaction time. Thurston (22) showed that the activity of most, but not all, laccases is affected by the chemical structure of the substrate. Substrate oxidation by laccase is a one-electron reaction generating a free radical, and the intermediates generated are typically unstable and can undergo chemical oxidation. If a dihydroxy phenolic compound such as catechol is used as co-substrate, oxidation by a catalyst yields initially a semiquinone radical anion that is further oxidized to oquinone. The quinone thus obtained is very susceptible to a reaction with nucleophilic compounds such as those containing NH2 groups. Martin et al. (23) have also shown that enzymatic oxidation of phenolic compounds in the presence of amino acids, peptides, and proteins yields nitrogenous polymers, and their linkage occurs through nucleophilic addition. In the present study, two products (A and B) were isolated after reaction of bentazon and catechol in the presence of the laccase. Product A with a molecular weight of 348 was found to be a dimer composed of one bentazon molecule (MW 240) and one catechol molecule (MW 110), while product B (MW 586) appeared to be a trimer consisting of two bentazon molecules and one catechol molecule. The coupling of a bentazon molecule with a catechol molecule to produce product A results in the removal of two hydrogens. In such a situation, three possible structures could be anticipated: (1) binding of a carbon of o-quinone (generated from catechol) to a carbon in the aromatic ring of bentazon (carbon-carbon coupling); (2) binding of the oxygen from the semiquinone radical anion to a carbon in the aromatic ring of bentazon (carbon-oxygen coupling); and (3) binding of a carbon of o-quinone to the protonated nitrogen of the heterocyclic ring of bentazon (carbon-nitrogen coupling). 1H-NMR and 13C-NMR spectroscopy indicated that binding via the third of these alternatives occurred. In the case of
product A, a dimer was produced by the binding of the protonated nitrogen of one bentazon molecule to a carbon of the o-quinone molecule generated from oxidized catechol (Figure 7). Similarly, in the case of product B, a trimer was formed by the binding of the protonated nitrogens of two bentazon molecules each to a carbon of an o-quinone molecule. The reaction of bentazon with catechol in the presence of a laccase is not a radical reaction, but proceeds via nucleophilic addition to a quinone structure. Therefore, it appears that the herbicide bentazon can be incorporated into soil organic matter by nucleophilic addition to quinone-like substances during the humification process. From previous studies (25), it is known that many pesticides are hydrolyzed or otherwise transformed to phenols or aromatic amines, and these intermediates are frequently incorporated into humic materials by oxidative coupling reactions. If the pesticide intermediates are substituted phenols or anilines, their reaction with oxidative catalysts proceeds easily and quickly produces free radicals for binding. The final chemical linkages formed are stable. In previous work (3), the phenolic intermediates of bentazon, 6-hydroxyand 8-hydroxy bentazon, were transformed rapidly and completely by oxidative enzymes in the absence of a cosubstrate. The present study demonstrates that bentazon, a relatively inert chemical, can also be incorporated into humic substances in the presence of a laccase.
Acknowledgments Funding for this research project was provided by the Office of Research and Development, Environmental Protection Agency (EPA; Grant R-823847). The EPA does not necessarily endorse any commercial products used in the study, and the conclusions represent the views of the authors and do not necessarily represent the opinions, policies, or recommendations of the EPA. We would like to thank Dr. L. M. Jackman for his help in interpreting the NMR data. We also like to thank Novo Nordisk Bioindustrials, Inc. (Danbury, CT), for providing the laccase from the fungus Polyporus pinsitus.
Literature Cited (1) Koskinen, W. C.; Harper, S. S. In Pesticides in the Soil Environment: Processes, Impacts, and Modeling; Cheng, H. H., Ed.; Soil Science Society of America, Inc.: Madison, WI, 1990; pp 51-73. (2) Stevenson, F. J. Humus Chemistry; Genesis, Composition, Reactions, 2nd ed.); John Wiley & Sons, Inc.: New York, 1994; pp 453-471. (3) Kim, J.-E.; Wang, J. C.-J.; Bollag, J.-M. Submitted for publication to Biodegradation. (4) Lee, J. K.; Fu ¨ hr, F.; Mittelstaedt, W. Chemosphere 1988, 17, 441450. (5) Ebert, D. Ph.D. Thesis, Natural Science Department, RuprechtKarls-University, Heidelberg, 1992. (6) Bollag, J.-M.; Myers, C. J.; Minard, R. D. Sci. Total Environ. 1992, 123/124, 205-217. (7) Nannipieri, P.; Bollag, J.-M. J. Environ. Qual. 1991, 20, 510-517. (8) Dec, J.; Bollag, J.-M. Arch. Environ. Contam. Toxicol. 1990, 19, 543-550. (9) Ruggiero, P.; Dec, J.; Bollag, J.-M. In Soil Biochemistry, Vol. 9; Stotzky, G., Bollag, J.-M., Eds.; Marcel Dekker, Inc.: New York, 1996; pp 79-122. (10) Sparks, D. L. Environmental Soil Chemistry; Academic Press: New York, 1994; pp 53-79. (11) Roper, J. C.; Sarkar, J. M.; Dec, J.; Bollag. J.-M. Water Res. 1995, 29, 2720-2724. (12) Dec, J.; Bollag, J.-M. Environ. Sci. Technol. 1994, 28, 484-490. (13) Tatsumi, K.; Freyer, A.; Minard, R. D.; Bollag, J.-M. Soil Biol. Biochem. 1994, 26, 735-742. (14) Simmons, K. E.; Minard, R. D.; Freyer, A. J.; Bollag, J.-M. Int. J. Environ. Anal. Chem. 1986, 26, 209-227. (15) Huber, R.; Otto, S. Rev. Environ. Contam. Toxicol. 1994, 137, 111-134. (16) Otto, S.; Beutel, P.; Drescher, N.; Huber, R. Advances in Pesticide Science IUPAC and Pergamon Press: Oxford, 1978; pp 551-556.
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(17) Stevenson, F. J. Humus Chemistry; Genesis, Composition, Reactions, 2nd ed.); John Wiley & Sons, Inc.: New York, 1994; pp 24-56. (18) Shaka, A. J.; Keeler, J.; Freeman, R. J. Magn. Reson. 1983, 53, 313-340. (19) Shaka, A. J.; Barker, P. B.; Freeman, R. J. Magn. Reson. 1985, 64, 547-552. (20) Sjoblad, R. D.; Bollag, J.-M. In Soil Biochemistry, Vol. 5; Paul, E. A., Ladd, J. N., Eds.; Marcel Dekker: New York, 1981; pp 113152. (21) Sarkar, J. M.; Malcolm, R. L.; Bollag, J.-M. Soil Sci. Soc. Am. J. 1988, 52, 688-694. (22) Thurston, C. F. Microbiology 1994, 140, 19-26. (23) Martin, J. P.; Haider, K.; Bondietti, E. In Proceedings of the International Meeting on Humic Substances, Nieuwersluis; Pudoc: Wageningen, 1975; pp 171-186.
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(24) Nonhebel, D. C.; Walton, J. C. Free-radical chemistry; structure and mechanism; Cambridge University Press: London, 1974; pp 330-340. (25) Bollag, J.-M. In Pesticide Transformation Products; Somasundaram, L., Coats, J. R., Eds.; American Chemical Society: Washington, DC, 1991; pp 122-132.
Received for review December 9, 1996. Revised manuscript received April 11, 1997. Accepted April 21, 1997.X ES961016L
X
Abstract published in Advance ACS Abstracts, June 15, 1997.
