- Email: [email protected]
Biodegradation of the herbicide bromoxynil and its plant cell wall bound residues in an agricultural soil
PESTICIDE Biochemistry & Physiology
Pesticide Biochemistry and Physiology 78 (2004) 49–57 www.elsevier.com/locate/ypest
Biodegradation of the herbicide bromoxynil and its plant cell wall bound residues in an agricultural soil P. Rosenbrock, J.C. Munch, I. Scheunert, and U. D€ orfler* GSF-National Research Center for Environment and Health, Institute of Soil Ecology, DE-85764 Neuherberg, Germany Received 2 May 2003; accepted 9 September 2003
Abstract The mineralization and formation of metabolites and nonextractable residues of the herbicide [14 C]bromoxyniloctanoate ([14 C]3,5-dibromo-4-octanoylbenzonitrile) and the corresponding agent substance [14 C]bromoxynil ([14 C]3,5dibromo-4-hydroxybenzonitrile) was investigated in a soil from an agricultural site in a model experiment. The mineralization of maize cell wall bound bromoxynil residues was also investigated in the agricultural soil material. The mineralization of [14 C]bromoxynil and [14 C]bromoxyniloctanoate in soil within 60 days amounted up to 42 and 49%, respectively. After the experiments, 52% of the originally applied [14 C]bromoxynil and 44% of the [14 C]bromoxyniloctanoate formed nonextractable residues in soil. Plant cell wall bound [14 C]bromoxynil residues were also mineralized to an extent of about 21% within 70 days; the main portion of 76% persisted as nonextractable residues in the soil. In bacterial enrichment cultures and in soil two polar metabolites were observed; one of it could be identified as 3,5-dibromo-4-hydroxybenzoate and the other could be described tentatively as 3,5-dibromo-4-hydroxybenzamide. Ó 2003 Elsevier Inc. All rights reserved. Keywords: Bromoxynil; Bromoxyniloctanoate; Cell wall bound residues; Mineralization; Soil; Microbial enrichment cultures
1. Introduction Bromoxynil (3,5-dibromo-4-hydroxybenzonitrile) is a widely used postemergence herbicide for the control of broad leaved weeds in cereal crops. Since atrazine was banned in Germany, bromoxynil is an important substitute for this herbicide. Biodegradation experiments showed a rapid disappearance of bromoxynil [1] and a high mineralization of bromoxyniloctanoate [2] in soil. In a * Corresponding author. Fax: +49-89-3187-3376. E-mail address: doerfl[email protected] (U. D€ orfler).
biofilm reactor Agrobacterium radiobacter was able to degrade bromoxynil within a short time [3]. In soil and in cultures of Pseudomonas putida 13XF, various metabolites resulted from stepwise hydrolysis of the cyano group (formation of the corresponding benzamide and benzoic acid), from partial debromination, and from hydroxyl methylation [4]. Vokounova et al. [5] and Gabriel et al. [6] also found Pseudomonas putida 13XF to be able to convert bromoxynil to 3,5-dibromo-4-hydroxybenzamide and 3,5-dibromo-4-hydroxybenzoic acid in the presence of a carbon and energy source, indicating that this strain possesses nitrile
0048-3575/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/j.pestbp.2003.09.002
50
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
hydratase activity (formation of the benzamide) as well as amidase activity (formation of the benzoic acid). A. radiobacter 8/4 however lacks of the amidase activity, since 3,5-dibromo-4-hydroxybenzamide was found as the only metabolite from bromoxynil degradation [6]. Klebsiella pneumoniae subsp. ozaenae uses bromoxynil as nitrogen source by direct formation of 3,5-dibromo-4-hydroxybenzoic acid and NHþ 4 with the enzyme nitrilase [7]. Smith and Cullimore [8] observed a bromoxynil degradation to 3,5-dibromo-4-hydroxybenzamide and 3,5-dibromo-4-hydroxybenzoic acid and a further nonidentified metabolite by a strain of Flexibacterium. In contrast, a Flavobacterium strain degrades bromoxynil by cleavage of the nitrile group and formation of cyanide and 2,6dibromohydrochinone [9]. The responsible enzyme is a hydroxylase capable to transform pentachlorophenol. In model aquifers two main pathways of bromoxynil metabolism were found: hydrolysis of the nitrile group and replacement of bromine by chlorine when chloride was present in the matrix [10]. Abiotic degradation of bromoxynil by photolysis is also reported [11,12]. The postemergence application of the herbicide results in the uptake into the cultivated plants. The susceptibility of a plant species decreases with increasing rates of bromoxynil metabolism [13]. In the last years, gene engineered plants were constructed, which are resistant against oxynil herbicides, by insertion of the nitrilase gene into various dicotyledonous species, such as tobacco, tomato, cotton, oilseed rape, carrot, potato, and eggplant [14,15]. In metabolism studies of bromoxynil in plants, the metabolites bromohydrochinone and 2,6dibromohydrochinone and the formation of glucosidic conjugates and nonextractable residues (NER) could be observed in Hordeum vulgare and Stellaria media [13]. After harvest, the herbicide residues which are adsorbed or absorbed by the plant are introduced, together with the plant residues, into the soil. Regarding this process, there are two possible ways on which a xenobiotic can enter soil—as the free substance and as a plant adsorbed or incorporated residue. The plant incorporated residues
can be present as extractable or nonextractable residues. Information on the fate of nonextractable ‘‘ plant bound’’ residues introduced into soil is very limited. In the present work, the mineralization of plant cell wall bound residues of bromoxynil in an agricultural soil was studied and compared with the mineralization of free applied bromoxynil and bromoxyniloctanoate. The cell wall bound residues were produced by applying the postemergence herbicide bromoxynil directly to the leaves of maize plants and by isolating the fraction of nonextractable residues after 6 days of plant growth.
2. Materials and methods 2.1. Investigated soil An agricultural soil (Dystric Eutrudept, 0– 30 cm) from the FAM Research Station Scheyern was used for biodegradation experiments. The Research Station is situated in the tertiary hills, 40 km north of Munich, at an altitude of 445– 498 m above sea level at the western end of the hops-growing ‘‘ Hallertau region.’’ After sampling, the soil material was sieved (<2 mm) and stored at 4 °C. Prior to the start of the experiments, the soil samples were equilibrated for 14 days at room temperature (20 °C) at a water content of about 25% of maximum water holding capacity (WHC). The soil characteristics determined, were as follows: 43% sand, 43% silt, 14% clay, 0.16% total nitrogen, 2.78% organic carbon, pH (CaCl2 ) 6.6. 2.2. Materials Uniformly 14 C ring labeled 3,5-dibromo-4octanoylbenzonitrile ([14 C]bromoxyniloctanoate; Fig. 1; radiochemical purity >98%, specific radioactivity 604 MBq mmol1 ) was purchased from the Institute of Isotopes (Budapest, Hungary). Uniformly 14 C ring labeled 3,5-dibromo-4hydroxybenzonitrile ([14 C]bromoxynil, Fig. 1) was prepared by cleaving the octanoyl group of the [14 C]bromoxyniloctanoate by an esterase reaction (esterase from rabbit liver in 3.6 M (NH4 )2 SO4 , 0.1 M Tris, pH 8.5, Sigma–Aldrich, Steinheim,
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
Fig. 1. Chemical structure of bromoxynil (I) and bromoxyniloctanoate (II).
Germany). Ten units esterase were dissolved in 1000 ll bidistilled water and incubated with 145 ll of a solution of [14 C]bromoxyniloctanoate in methanol (0.7912 lg ll1 , 71,000 dpm ll1 ) overnight at 20 °C (pers. commun., O. Fr€ oscheis, GSFInstitute for Biochemical Plant Pathology). [14 C]Bromoxynil was extracted from the aqueous solution, acidified with 85% H3 PO4 to a pH < 2, by solid phase extraction (Bond Elut-ENV, Varian, Darmstadt, Germany). The quantitative transformation of [14 C]bromoxyniloctanoate to [14 C]bromoxynil was assayed with HPLC and was found to be >98%. The unlabeled reference substances bromoxynil and 3,5-dibromo-4-hydroxybenzamide were purchased from Sigma–Aldrich (Steinheim, Germany), with a purity of >98 and 99%, respectively. Bromoxyniloctanoate was purchased from Dr. Ehrenstorfer GmbH (Augsburg, Germany) with a purity of 98.7%. For determination of radioactivity in liquid samples, aliquots of the sample were mixed with a scintillation cocktail (Ultima Gold XR, Packard, Dreieich, Germany) and measured in a liquid scintillation counter (Tri-Carb 1900TR, Packard, Dreieich, Germany). 2.3. Preparation of cell wall bound residues of [14 C]bromoxynil in maize plants
51
heim, Germany) for 25 days (14 h light and 25 °C, 10 h darkness and 20 °C; 70% rel. air humidity) until they reached the six-leaf stage. Before [14 C]bromoxynil was applied onto the leaves (third to sixth eldest leaves), the leaves were moistened with a Tween 80 solution (1.3 g L1 ) with a fine brush to improve the wettability of the plant surfaces. Then, the leaves of each plant were brushed with 150 ll [14 C]bromoxynil water–methanol solution (14.5% methanol, 18.2 kBq). After a preincubation of 20 h in the dark to protect the applied substance against photodegradation, the plants were kept under light for 14 h per day. The plants were harvested after 6 days of incubation. Shoots and leaves were cut into 2 cm pieces, freeze dried, and pulverized in a mill (Retsch, Haan, Germany). The radioactivity in the maize meal was determined by combustion (Oxidizer 306, Packard, Dreieich, Germany), trapping the 14 CO2 in Carbo-Sorb E (Packard, Dreieich, Germany) and mixing it with Permafluor E (Packard, Dreieich, Germany) prior to scintillation counting. It amounted to 12.2 Bq mg1 . The labeled maize meal was extracted twice with Bligh–Dyer solution (CH3 OH:CH2 Cl2 :H2 0, 2:1:0.8) followed by an extraction with water, 80% methanol, and 1% sodium dodecylsulfonate [16]. The remaining plant material was extracted twice with distilled water. After each extraction step, plant material was separated from the solvent by centrifugation (15 min, 7240g). The radioactivity of the extracted maize meal was determined by combustion as described above and amounted to 3.6 Bq mg1 , representing nonextractable residues (NER) of the applied herbicide. To characterize the [14 C]bromoxynil-binding cell wall fractions, aliquots of the extracted maize meal (500 mg, n ¼ 3) were fractionated into their cell wall constituents starch, pectin, protein, hemicellulose, cellulose, and lignin by enzymatic processes [17]. 2.4. Spiking of the soil
Maize plants were grown on quartz sand (0.1– 0.5 mm, dried at 105 °C, Georg Schuchmacher GmbH, Lauingen, Germany) with 25% w/w Hoagland solution (preparation according to the manufacture instructions, Sigma–Aldrich, Stein-
[14 C]Bromoxynil and [14 C]bromoxyniloctanoate, mixed with unlabeled bromoxynil and bromoxyniloctanoate, respectively, were dissolved in 250 ll methanol and applied to a 1000 mg dried,
52
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
homogenized soil sample in a 10 ml glass vial. After evaporation of methanol the soil aliquot was carefully stirred with a spatula and then transferred to a 200 ml beaker glass. Here it was mixed with a total amount of 49 g fresh soil (dry weight equivalent, 20% of WHC) in three subsequent mixing steps. The spiked soil sample was then transferred to the incubation flask, adjusted to 40% WHC and connected to the biodegradation system. Spiking the soil in this way ensured a homogeneous distribution of the xenobiotic in the soil. The initial radioactivity added to each soil sample was 133 kBq, corresponding to a concentration of 1.62 lg g1 [14 C]bromoxynil and 2.16 lg g1 [14 C]bromoxyniloctanoate, respectively. For the degradation experiments with maizebound residues, 500 mg freeze dried pulverized [14 C]NER meal (1.8 kBq) was mixed with 50 g (dry weight equivalent) fresh soil, in the same way as described above. 2.5. Design of biodegradation experiments The biomineralization experiments were carried out in 500 ml flasks with three replicates per treatment. The spiked soil samples (50 g dry weight) were incubated for 60 days ([14 C]bromoxynil, [14 C]bromoxyniloctanoate) or 70 days ([14 C]NER) at 22 °C in the dark. The flasks were discontinuously aerated for one hour per day with moistened air (15 ml min1 ). After passage through the flasks, the air was trapped in a system of absorption tubes. The first tube was filled with 10 ml ethylene glycol monomethylether (Merck, Darmstadt, Germany) to absorb volatilized substances. The two following tubes were filled with 10 ml of a mixture of ethanolamine and diethylene glycol monobutylether (1:1, Merck, Darmstadt, Germany) to absorb 14 CO2 . With this sequence of absorption tubes a strict separation of volatile 14 Ccompounds and mineralized 14 CO2 is achieved and a correct measurement of the mineralization is ensured. For 14 C counting the absorption liquids were mixed with 10 ml Permablend I scintillation cocktail (22 g Permablend I, Packard, Dreieich, Germany, dissolved in 1 L toluene). Cumulative 14 CO2 in percentage of 14 C applied was plotted as a function of time. Analysis of variance was used to
analyze the different biodegradability of the tested substances. Means were compared by Students– Newman–Keuls test (n ¼ 3, p ¼ 0:05). 2.6. Quantification of nonextractable residues and detection of metabolites in soil The mass balance of radioactive carbon was determined by addition of radioactivity trapped as 14 CO2 , radioactivity volatilized during the experiment and residual radioactivity (nonextractable and extractable) in the soil after the experiments. The soil samples were soxhlet extracted with methanol for 24 h. After extraction aliquots of the soil samples were combusted as described above to determine the amount of nonextractable residues. Radioactivity in the extract was determined by liquid scintillation counting. The extracts were evaporated in a rotary evaporator to dryness and dissolved in a mixture of ethyl acetate and cyclohexane (1:1, chromatography grade, Merck, Darmstadt, Germany). The clean up of soil extracts was performed with an automated gel permeation chromatography system (GPC, Abimed, Langenfeld, Germany). Mobile phase: ethyl acetate:cyclohexane 1:1, flow: 5 ml min1 , column: biobeads S-X3 (Bio-Rad Laboratories GmbH, Munich, Germany). The concentrated GPC fractions were analyzed by HPLC (Merck, Darmstadt, Germany), equipped with a UV/VIS detector and a radioactivity detector LB 507 B (Berthold, Wildbad, Germany). Column: 250 4 mm LiChrospher 100 RP-18, 5 lm (Merck, Darmstadt, Germany). Mobile phase (HPLC grade): A ¼ bidistilled water, B ¼ methanol, C ¼ 0.5% acetic acid. Gradient: T0 : A ¼ 60%, B ¼ 39%, C ¼ 1%; T20 : A ¼ 10%, B ¼ 89%, C ¼ 1%; T25 : A ¼ 10%, B ¼ 89%, C ¼ 1%; T30 : A ¼ 60%, B ¼ 39%, C ¼ 1%; T35 : A ¼ 30%, B ¼ 69%, C ¼ 1%. Flow: 1 ml min1 ; column temperature: 20 °C. 2.7. Degradation of bromoxynil in bacterial enrichment cultures The soil was mixed with a bromoxynil solution (water, 10% methanol), resulting in a water content of 40% WHC and a bromoxynil concentration of 2.2 lg g1 soil dry weight. Then, the soil was
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
incubated for 10 days at room temperature in a water saturated atmosphere. After this adaptation phase, 2 g soil (fresh weight) was transferred into 20 ml of a mineral salt medium containing bromoxynil and a nitrogen source (MS-BO-Nþ ) (NaNO3 0.5 g L1 , K2 HPO4 0.65 g L1 , KH2 PO4 0.17 g L1 , MgSO 4 205 mg L1 , bromoxynil 40 mg L1 ) and incubated at 28 °C. In this enrichment culture, the soil inoculum served as carbon source. After enrichment, 1 ml of this culture was transferred, first, into fresh MS-BO-Nþ , second, into mineral salt medium with an additional carbon source MS-BO-Nþ -Cþ (glucose 500 mg L1 ), and third, into mineral salt medium with an additional carbon source but without nitrogen MSBO-Cþ -N (glucose 500 mg L1 ). The bromoxynil biodegradation in these cultures was recorded by detecting the metabolites in the HPLC-UV/VIS system described above. For this purpose 50 ll of culture medium was injected directly into the HPLC after removing particles by centrifugation (10 min, 7240g) or the culture medium was extracted by solid phase extraction (SPE) described above. Further investigations of formed metabolites were done by analyzing SPE extracts of culture medium in a LC/MS system (ThermoQuest, Egelsbach, Germany, negative ionisation).
3. Results 3.1. Characterization of maize cell wall bound residues About 76% of the radioactivity applied onto the maize plants could be recovered by combustion after 6 days of incubation. As much as 34% of the radioactivity remaining adsorbed or absorbed by the plant was due to nonextractable residues whereas about 66% were extractable with water and organic solvents. The formed nonextractable residues were distributed between the specific cell wall fractions as shown in Fig. 2. The lignin fraction is the main and probably most resistant cell wall fraction, which is involved in the bound residue formation. About 55% of the total formed bound residues were detected in the lignin fraction. The nonextractable
53
Fig. 2. Distribution of bound [14 C]bromoxynil residues in different cell wall fractions of maize plants.
residues in the easily degradable starch, protein and pectin cell wall fractions (8, 24, and 12%, respectively), will not play an important role concerning the persistence of bromoxynil residues in soils. 3.2. Fate of bromoxynil, bromoxyniloctanoate, and maize cell wall bound residues in soil As shown in Fig. 3, mineralization of free applied [14 C]bromoxynil and [14 C]bromoxyniloctanoate started without a lag phase. Within the first 5 days 14 CO2 -production from [14 C]bromoxynil and [14 C]bromoxyniloctanoate increased up to 25 and 30%, respectively. Then mineralization
Fig. 3. Cumulative release of 14 CO2 from [14 C]bromoxynil (14C-BO), [14 C]bromoxyniloctanoate (14C-BOO), and nonextractable [14 C]bromoxynil residues of maize plants (14C-NER) in an agricultural soil.
54
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
rates decreased and within 60 days up to 42% of [14 C]bromoxynil and 49% of [14 C]bromoxyniloctanoate were mineralized (Table 1). The amounts of 14 CO2 released from [14 C]bromoxynil and [14 C]bromoxyniloctanoate after an incubation period of 60 days did not differ significantly. Biomineralization of [14 C]NER occurred with nearly constant mineralization rates (Fig. 3) and was significantly lower than mineralization of the free applied herbicide. After 70 days 21% of [14 C]NER was mineralized, indicating that the soil microorganisms were able to attack the cell wall bound bromoxynil residues and to degrade them to 14 CO2 (Table 1). The extraction of soil material from the experiments with free applied [14 C]bromoxynil and [14 C]bromoxyniloctanoate showed that only a small part of the applied 14 C-substances were extractable (Table 1). After an incubation time of 60 days only 2.9 and 4.4% of the initally applied radioactivity could be extracted in the [14 C]bromoxynil and [14 C]bromoxyniloctanoate degradation experiments, respectively. Consequently, 52.3 and 44.3% of the applied radioactivity could be recovered by combustion as nonextractable residues from [14 C]bromoxynil and [14 C]bromoxyniloctanoate, respectively. Thus, the decreasing mineralization rates could be explained by the decreasing bioavailability in the course of the experiments. The soil residues of the degradation experiments with the plant cell wall bound [14 C] bromoxynil ([14 C]NER) showed a negligible extractability of 1.4% of the applied radioactivity, the main portion (76%) persisted as nonextractable residues in the soil (Table 1). The analysis of the [14 C]bromoxynil metabolites, formed in soil during the incubation time, were
complicated by the low extractability of the remaining residues, due to the strong tendency to form nonextractable residues (Table 1). The extract of the soil from the [14 C]bromoxynil biodegradation experiment was subjected to a GPC cleanup and the main portion of the radioactivity was found in the fractions collected between the retention volumes 150 and 195 ml. This fraction contained 58% of the radioactivity sampled in GPC fractions. In the other fractions the radioactivity was below the detection limit. HPLC analyses of this fraction showed beside the [14 C]bromoxynil itself, only one polar metabolite. To overcome the difficulties of the high formation rate of nonextractable residues in the degradation experiments with soil, the formation of metabolites was investigated in microbial enrichment cultures. In the enrichment cultures with an additional carbon source and bromoxynil as sole source of nitrogen, the formation of two polar metabolites could be observed. HPLC analyses revealed that metabolite 1 had the same retention time as the metabolite that was observed in the soil extract. Further analysis of this metabolite by LC/MS showed a molecular ion of m=z 292/294/296 and the typical isotope distribution pattern of a dibrominated molecule (Fig. 4). Thus, metabolite 1 could be tentatively described as 3,5-dibromo-4-hydroxybenzamide. Metabolite 2 could be identified as 3,5-dibromo-4-hydroxybenzoic acid by comparing the retention time with an authentic reference compound.
4. Discussion From the total bromoxynil residues in plants, 34% nonextractable residues were formed within 6
Table 1 Distribution and mass balance of the radioactivity applied to soil in the biodegradation experiments (in % of Experiment
14
[14 C]Bromoxynila [14 C]Bromoxyniloctanoatea [14 C]NERb
42.4 1.3 49.3 9.4 21.6 8.5
a
60 days incubation. 70 days incubation. c n.d., not detectable. b
CO2
14
C volatile
0.03 0.01 0.06 0.03 n.d.c
14
C nonextractable
52.3 2.3 44.3 3.5 76.5 2.6
14
C extractable
2.9 0.3 4.4 0.9 1.4 0.8
14
C applied) 14
C recovery
97.6 3.4 98.0 13.4 99.5 6.7
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
Fig. 4. Mass spectrum of metabolite 1 in a negative ionization mode.
days of plant growth. The main portion of the nonextractable residues were bound to the lignin fraction, whereas to the more easily degradable cell wall fractions like starch, protein or pectin only minor amounts were bound. The potential adverse long term effects of plant incorporated bromoxynil residues will therefore depend on the degradability of lignin in soil. White rot fungus is known to posses lignolytic enzymes which enable this fungus to degrade lignin in wood. For different soil fungi species like Fusarium [18,19], Trichoderma [20], and Aspergillus [21] the degration of lignin has also been reported. The potentiality for the release of bromoxynil and/or its metabolites from lignin bound residues is therefore present in soil. In the biodegradation experiments, the [14 C] bromoxynil plant cell wall bound residues ([14 C]NER) showed significantly lower mineralization rates than the free applied herbicides bromoxynil and bromoxyniloctanoate. In other
55
experiments with [14 C]isoproturon and plant-incorporated as well as cell wall bound residues of [14 C]isoproturon, the incorporated and bound residues showed also lower mineralization rates than the directly applied herbicide [22,23]. In our study, after an incubation time of 70 days, 21% of the [14 C]NER were completely degraded to 14 CO2 , indicating that the indigenous microbial community was able to release the bound bromoxynil or its metabolites and to mineralize them. Whether the released 14 C-residues were originally bound to the lignin fraction or to the more easily degradable starch, protein or pectin fractions cannot be decided. Since the main portion of the bound 14 Cresidues (55%) was bound to the lignin fraction it can be assumed that the mineralization of the minor 21% portion is probably due to the release of the 14 C-residues bound to the easily degradable cell wall fractions. But for a definite answer to this question further investigations with lignin residues are necessary. The degradation of the free applied herbicides bromoxynil and bromoxyniloctanoate occurred without a lag phase and with high initial mineralization rates, reflecting a good degradability of this herbicide in soils which has been already reported [1,2]. Therefore it can be assumed, that the enzymes which are involved in the mineralization of bromoxynil were already produced by the microorganisms in the soil. Since there was no significant difference in the mineralization of bromoxynil and bromoxyniloctanoate, the cleavage of the ester-binding of the octanoate seems to happen very quickly, as has already been reported in the literature [2], and the following mineralization is referred to the resulting bromoxynil. The decrease of the mineralization rates with time are probably due to a decreasing bioavailability because of formation of nonextractable residues. After 60 days of incubation about 50% of the applied radioactivity was not extractable. The strong tendency for the formation of nonextractable residues can be explained by the reaction of reactive groups within the bromoxynil molecule (hydroxyl group) or its metabolites (amide or carboxyl group) with natural soil components, i.e., humic acids. The low extractability of the 14 Cresidues (2–4% of applied radioactivity) also
56
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
reflects the high transformation rate—mainly mineralization and formation of nonextractable residues—of bromoxynil in soils. Two polar metabolites were observed as degradation products of bromoxynil. One metabolite could be identified as 3,5-dibromo-4-hydroxybenzoic acid and the other was characterized as 3,5-dibromo-4-hydroxybenzamide. As generally known, hydrolysis of the nitrile group is the basic reaction in the transformation of bromoxynil. Principally, the hydrolysis can occur in a one step or a two step process (Fig. 5). Klebsiella pneumonia subsp. ozaenae converts bromoxynil by the enzyme nitrilase directly to the carboxylic acid derivative in the absence of any detectable amide intermediate [7]. Other bromoxynil decomposers like Pseudomonias putida 13XF [4–6] and Flexibacterium BR4 [8] convert the nitrile group in a first step to the amide group by the enzyme nitrile hydratase and in a second step the resulting amide is subsequently hydrolysed to the corresponding carboxylic acid by amidase. In A. radiobacter 8/4, however, no amidase activity was found, since this strain transforms
bromoxynil only to the corresponding amide without further degradation. Bromoxynil degradation not only occurs via hydrolysis of the nitrile group, Topp et al. [9] found a Flavobacterium species to be able to replace the nitrile group with a hydroxyl group, yielding the corresponding chinone by the action of pentachlorophenol hydroxylase (Fig. 5). The results of the present study indicate that in the investigated soil a microbial community was present which was able to mineralize bromoxynil. In the whole process probably various microorganisms were involved, initiating the bromoxynil degradation via hydrolysis of the nitrile group to the corresponding amide and carboxylic acid. The acid was quickly decarboxylized and after ring cleavage the 14 C-ring atoms were completely metabolised to 14 CO2 . No accumulation of the intermediate products could be observed, they were only detected in small amounts. The results of the current study show, that the main degradation and transformation pathways of directly applied bromoxynil in soil were mineralization and formation of bound residues. Plant cell
Fig. 5. Initial degradation reactions of bromoxynil and some enzymes and microorganisms involved, according to the literature [5–9].
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
wall bound residues of bromoxynil were also mineralized but in a lower extent than the free compound. A part of the plant cell wall bound bromoxynil residues could be released in the soil and subjected to complete degradation whereas the main portion persisted as nonextractable residues.
References [1] A.E. Smith, Degradation of bromoxynil in regina heavy clay, Weed Res. 11 (1971) 276–282. [2] R.F. Collins, Perfusion studies with bromoxynil octanoate in soil, Pestic. Sci. 4 (1973) 181–192. [3] D. M€ uller, J. Gabriel, Bacterial degradation of the herbicide bromoxynil by Agrobacterium radiobacter in biofilm, Folia Microbiol. 44 (1999) 377–379. [4] L.A. Golovleva, R.N. Pertsova, F. Kunc, M. Vokounova, Decomposition of the herbicide bromoxynil in soil and in bacterial cultures, Folia Microbiol. 33 (1988) 491–499. [5] M. Vokounov a, O. Vacek, F. Kunc, Effect of glucose and ribose on microbial degradation of the herbicide bromoxynil continuously added to soil, Folia Microbiol. 37 (1992) 128–132. [6] J. Gabriel, J. Vekova, J. Vosahlo, High-performance liquid chromatographic study of the aromatic nitrile metabolism in soil bacteria, J. Chromatogr. B 681 (1996) 191–195. [7] K.E. McBride, J.W. Kenny, D.M. Stalker, Metabolism of the herbicide bromoxynil by Klebsiella pneumoniae subsp. ozaenae, Appl. Environ. Microbiol. 52 (1986) 325–330. [8] A.E. Smith, D.R. Cullimore, The in vitro degradation of the herbicide bromoxynil, Can. J. Microbiol. 20 (1974) 773–776. [9] E. Topp, L. Xun, C.S. Orser, Biodegradation of the herbicide bromoxynil (3,5-dibromo-4-hydroxybenzonitrile) by purified pentachlorophenol hydroxylase and whole cells of Flavobacterium sp. strain ATCC 39723 is accompanied by cyanogenesis, Appl. Environ. Microbiol. 58 (1992) 502–506. [10] B. Graß, H. Mayer, J. Nolte, G. Preuß, N. Zullei-Seibert, Studies on the metabolism of hydroxybenzonitrile herbicides: I. mass spectrometric identification, Pest. Manag. Sci. 56 (2000) 49–59.
57
[11] M. Millet, W.-U. Palm, C. Zetzsch, Abiotic degradation of halobenzonitriles: investigation of the photolysis in solution, Ecotoxicol. Environ. Saf. 41 (1998) 44–50. [12] I. Texier, C. Giannotti, S. Malato, C. Richter, J. Delaire, Solar photodegradation of pesticides in water by sodium decatungstate, Catal. Today 54 (1999) 297–307. [13] B. Schaller, B. Schneider, H.R. Sch€ utte, Metabolism of the herbicide bromoxynil in Hordeum vulgare and Stellaria media, Z. Naturforsch. C 47 (1992) 126–131. [14] D.M. Stalker, K.E. McBride, L.D. Malyi, Herbicide resistance in transgenic plants expressing a bacterial detoxification gene, Science 242 (1988) 419–423. [15] G. Freyssinet, B. Pelissier, M. Freyssinet, R. Delon, Crops resistant to oxynils: from the laboratory to the market, Field Crops Res. 45 (1996) 125–133. [16] W. Ebing, A. Haque, I. Schuphan, Ecochemical assessment of environmental chemicals: draft guideline of the test procedure to evaluate metabolism and degradation of chemicals by plant cell culture, Chemosphere 13 (1984) 947–957. [17] D. Komoßa, I. Gennity, H. Sandermann Jr., Plant metabolism with C–P bonds: glyphosate, Pestic. Biochem. Physiol. 43 (1992) 85–94. [18] D.M. Norris, Degradation of 14 C-labeled lignins and 14 Clabeled aromatic acids by Fusarium solani, Appl. Environ. Microbiol. 40 (1980) 376–380. [19] J.B. Sutherland, A.L. Pometto, D.L. Crawford, Lignocellulose degradation by Fusarium species, Can. J. Bot. 61 (1983) 1194–1198. [20] T.W. Flegel, V. Meevootisom, S. Kiatpapan, Indications of lignolysis by Trichoderma strains isolated from soil during simultaneous screening for fungi with cellulase and laccase activity, J. Ferment. Technol. 60 (1982) 473–475. [21] K.L. Kadam, S.W. Drew, Study of lignin biotransformation by Aspergillus fumigatus and white-rot fungi using 14 Clabeled and unlabeled kraft lignins, Biotech. Bioeng. 28 (1986) 394–404. [22] S. Lehr, I. Scheunert, F. Beese, Mineralization of free and cell-wall-bound isoproturon in soils in relation to soil microbial parameters, Soil Biol. Biochem. 28 (1996) 1–8. [23] S. von Wiren-Lehr, I. Scheunert, U. D€ orfler, Mineralization of plant-incorporated residues of 14 C-isoproturon in arable soils originating from different farming systems, Geoderma 105 (2002) 351–366.
Pesticide Biochemistry and Physiology 78 (2004) 49–57 www.elsevier.com/locate/ypest
Biodegradation of the herbicide bromoxynil and its plant cell wall bound residues in an agricultural soil P. Rosenbrock, J.C. Munch, I. Scheunert, and U. D€ orfler* GSF-National Research Center for Environment and Health, Institute of Soil Ecology, DE-85764 Neuherberg, Germany Received 2 May 2003; accepted 9 September 2003
Abstract The mineralization and formation of metabolites and nonextractable residues of the herbicide [14 C]bromoxyniloctanoate ([14 C]3,5-dibromo-4-octanoylbenzonitrile) and the corresponding agent substance [14 C]bromoxynil ([14 C]3,5dibromo-4-hydroxybenzonitrile) was investigated in a soil from an agricultural site in a model experiment. The mineralization of maize cell wall bound bromoxynil residues was also investigated in the agricultural soil material. The mineralization of [14 C]bromoxynil and [14 C]bromoxyniloctanoate in soil within 60 days amounted up to 42 and 49%, respectively. After the experiments, 52% of the originally applied [14 C]bromoxynil and 44% of the [14 C]bromoxyniloctanoate formed nonextractable residues in soil. Plant cell wall bound [14 C]bromoxynil residues were also mineralized to an extent of about 21% within 70 days; the main portion of 76% persisted as nonextractable residues in the soil. In bacterial enrichment cultures and in soil two polar metabolites were observed; one of it could be identified as 3,5-dibromo-4-hydroxybenzoate and the other could be described tentatively as 3,5-dibromo-4-hydroxybenzamide. Ó 2003 Elsevier Inc. All rights reserved. Keywords: Bromoxynil; Bromoxyniloctanoate; Cell wall bound residues; Mineralization; Soil; Microbial enrichment cultures
1. Introduction Bromoxynil (3,5-dibromo-4-hydroxybenzonitrile) is a widely used postemergence herbicide for the control of broad leaved weeds in cereal crops. Since atrazine was banned in Germany, bromoxynil is an important substitute for this herbicide. Biodegradation experiments showed a rapid disappearance of bromoxynil [1] and a high mineralization of bromoxyniloctanoate [2] in soil. In a * Corresponding author. Fax: +49-89-3187-3376. E-mail address: doerfl[email protected] (U. D€ orfler).
biofilm reactor Agrobacterium radiobacter was able to degrade bromoxynil within a short time [3]. In soil and in cultures of Pseudomonas putida 13XF, various metabolites resulted from stepwise hydrolysis of the cyano group (formation of the corresponding benzamide and benzoic acid), from partial debromination, and from hydroxyl methylation [4]. Vokounova et al. [5] and Gabriel et al. [6] also found Pseudomonas putida 13XF to be able to convert bromoxynil to 3,5-dibromo-4-hydroxybenzamide and 3,5-dibromo-4-hydroxybenzoic acid in the presence of a carbon and energy source, indicating that this strain possesses nitrile
0048-3575/$ - see front matter Ó 2003 Elsevier Inc. All rights reserved. doi:10.1016/j.pestbp.2003.09.002
50
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
hydratase activity (formation of the benzamide) as well as amidase activity (formation of the benzoic acid). A. radiobacter 8/4 however lacks of the amidase activity, since 3,5-dibromo-4-hydroxybenzamide was found as the only metabolite from bromoxynil degradation [6]. Klebsiella pneumoniae subsp. ozaenae uses bromoxynil as nitrogen source by direct formation of 3,5-dibromo-4-hydroxybenzoic acid and NHþ 4 with the enzyme nitrilase [7]. Smith and Cullimore [8] observed a bromoxynil degradation to 3,5-dibromo-4-hydroxybenzamide and 3,5-dibromo-4-hydroxybenzoic acid and a further nonidentified metabolite by a strain of Flexibacterium. In contrast, a Flavobacterium strain degrades bromoxynil by cleavage of the nitrile group and formation of cyanide and 2,6dibromohydrochinone [9]. The responsible enzyme is a hydroxylase capable to transform pentachlorophenol. In model aquifers two main pathways of bromoxynil metabolism were found: hydrolysis of the nitrile group and replacement of bromine by chlorine when chloride was present in the matrix [10]. Abiotic degradation of bromoxynil by photolysis is also reported [11,12]. The postemergence application of the herbicide results in the uptake into the cultivated plants. The susceptibility of a plant species decreases with increasing rates of bromoxynil metabolism [13]. In the last years, gene engineered plants were constructed, which are resistant against oxynil herbicides, by insertion of the nitrilase gene into various dicotyledonous species, such as tobacco, tomato, cotton, oilseed rape, carrot, potato, and eggplant [14,15]. In metabolism studies of bromoxynil in plants, the metabolites bromohydrochinone and 2,6dibromohydrochinone and the formation of glucosidic conjugates and nonextractable residues (NER) could be observed in Hordeum vulgare and Stellaria media [13]. After harvest, the herbicide residues which are adsorbed or absorbed by the plant are introduced, together with the plant residues, into the soil. Regarding this process, there are two possible ways on which a xenobiotic can enter soil—as the free substance and as a plant adsorbed or incorporated residue. The plant incorporated residues
can be present as extractable or nonextractable residues. Information on the fate of nonextractable ‘‘ plant bound’’ residues introduced into soil is very limited. In the present work, the mineralization of plant cell wall bound residues of bromoxynil in an agricultural soil was studied and compared with the mineralization of free applied bromoxynil and bromoxyniloctanoate. The cell wall bound residues were produced by applying the postemergence herbicide bromoxynil directly to the leaves of maize plants and by isolating the fraction of nonextractable residues after 6 days of plant growth.
2. Materials and methods 2.1. Investigated soil An agricultural soil (Dystric Eutrudept, 0– 30 cm) from the FAM Research Station Scheyern was used for biodegradation experiments. The Research Station is situated in the tertiary hills, 40 km north of Munich, at an altitude of 445– 498 m above sea level at the western end of the hops-growing ‘‘ Hallertau region.’’ After sampling, the soil material was sieved (<2 mm) and stored at 4 °C. Prior to the start of the experiments, the soil samples were equilibrated for 14 days at room temperature (20 °C) at a water content of about 25% of maximum water holding capacity (WHC). The soil characteristics determined, were as follows: 43% sand, 43% silt, 14% clay, 0.16% total nitrogen, 2.78% organic carbon, pH (CaCl2 ) 6.6. 2.2. Materials Uniformly 14 C ring labeled 3,5-dibromo-4octanoylbenzonitrile ([14 C]bromoxyniloctanoate; Fig. 1; radiochemical purity >98%, specific radioactivity 604 MBq mmol1 ) was purchased from the Institute of Isotopes (Budapest, Hungary). Uniformly 14 C ring labeled 3,5-dibromo-4hydroxybenzonitrile ([14 C]bromoxynil, Fig. 1) was prepared by cleaving the octanoyl group of the [14 C]bromoxyniloctanoate by an esterase reaction (esterase from rabbit liver in 3.6 M (NH4 )2 SO4 , 0.1 M Tris, pH 8.5, Sigma–Aldrich, Steinheim,
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
Fig. 1. Chemical structure of bromoxynil (I) and bromoxyniloctanoate (II).
Germany). Ten units esterase were dissolved in 1000 ll bidistilled water and incubated with 145 ll of a solution of [14 C]bromoxyniloctanoate in methanol (0.7912 lg ll1 , 71,000 dpm ll1 ) overnight at 20 °C (pers. commun., O. Fr€ oscheis, GSFInstitute for Biochemical Plant Pathology). [14 C]Bromoxynil was extracted from the aqueous solution, acidified with 85% H3 PO4 to a pH < 2, by solid phase extraction (Bond Elut-ENV, Varian, Darmstadt, Germany). The quantitative transformation of [14 C]bromoxyniloctanoate to [14 C]bromoxynil was assayed with HPLC and was found to be >98%. The unlabeled reference substances bromoxynil and 3,5-dibromo-4-hydroxybenzamide were purchased from Sigma–Aldrich (Steinheim, Germany), with a purity of >98 and 99%, respectively. Bromoxyniloctanoate was purchased from Dr. Ehrenstorfer GmbH (Augsburg, Germany) with a purity of 98.7%. For determination of radioactivity in liquid samples, aliquots of the sample were mixed with a scintillation cocktail (Ultima Gold XR, Packard, Dreieich, Germany) and measured in a liquid scintillation counter (Tri-Carb 1900TR, Packard, Dreieich, Germany). 2.3. Preparation of cell wall bound residues of [14 C]bromoxynil in maize plants
51
heim, Germany) for 25 days (14 h light and 25 °C, 10 h darkness and 20 °C; 70% rel. air humidity) until they reached the six-leaf stage. Before [14 C]bromoxynil was applied onto the leaves (third to sixth eldest leaves), the leaves were moistened with a Tween 80 solution (1.3 g L1 ) with a fine brush to improve the wettability of the plant surfaces. Then, the leaves of each plant were brushed with 150 ll [14 C]bromoxynil water–methanol solution (14.5% methanol, 18.2 kBq). After a preincubation of 20 h in the dark to protect the applied substance against photodegradation, the plants were kept under light for 14 h per day. The plants were harvested after 6 days of incubation. Shoots and leaves were cut into 2 cm pieces, freeze dried, and pulverized in a mill (Retsch, Haan, Germany). The radioactivity in the maize meal was determined by combustion (Oxidizer 306, Packard, Dreieich, Germany), trapping the 14 CO2 in Carbo-Sorb E (Packard, Dreieich, Germany) and mixing it with Permafluor E (Packard, Dreieich, Germany) prior to scintillation counting. It amounted to 12.2 Bq mg1 . The labeled maize meal was extracted twice with Bligh–Dyer solution (CH3 OH:CH2 Cl2 :H2 0, 2:1:0.8) followed by an extraction with water, 80% methanol, and 1% sodium dodecylsulfonate [16]. The remaining plant material was extracted twice with distilled water. After each extraction step, plant material was separated from the solvent by centrifugation (15 min, 7240g). The radioactivity of the extracted maize meal was determined by combustion as described above and amounted to 3.6 Bq mg1 , representing nonextractable residues (NER) of the applied herbicide. To characterize the [14 C]bromoxynil-binding cell wall fractions, aliquots of the extracted maize meal (500 mg, n ¼ 3) were fractionated into their cell wall constituents starch, pectin, protein, hemicellulose, cellulose, and lignin by enzymatic processes [17]. 2.4. Spiking of the soil
Maize plants were grown on quartz sand (0.1– 0.5 mm, dried at 105 °C, Georg Schuchmacher GmbH, Lauingen, Germany) with 25% w/w Hoagland solution (preparation according to the manufacture instructions, Sigma–Aldrich, Stein-
[14 C]Bromoxynil and [14 C]bromoxyniloctanoate, mixed with unlabeled bromoxynil and bromoxyniloctanoate, respectively, were dissolved in 250 ll methanol and applied to a 1000 mg dried,
52
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
homogenized soil sample in a 10 ml glass vial. After evaporation of methanol the soil aliquot was carefully stirred with a spatula and then transferred to a 200 ml beaker glass. Here it was mixed with a total amount of 49 g fresh soil (dry weight equivalent, 20% of WHC) in three subsequent mixing steps. The spiked soil sample was then transferred to the incubation flask, adjusted to 40% WHC and connected to the biodegradation system. Spiking the soil in this way ensured a homogeneous distribution of the xenobiotic in the soil. The initial radioactivity added to each soil sample was 133 kBq, corresponding to a concentration of 1.62 lg g1 [14 C]bromoxynil and 2.16 lg g1 [14 C]bromoxyniloctanoate, respectively. For the degradation experiments with maizebound residues, 500 mg freeze dried pulverized [14 C]NER meal (1.8 kBq) was mixed with 50 g (dry weight equivalent) fresh soil, in the same way as described above. 2.5. Design of biodegradation experiments The biomineralization experiments were carried out in 500 ml flasks with three replicates per treatment. The spiked soil samples (50 g dry weight) were incubated for 60 days ([14 C]bromoxynil, [14 C]bromoxyniloctanoate) or 70 days ([14 C]NER) at 22 °C in the dark. The flasks were discontinuously aerated for one hour per day with moistened air (15 ml min1 ). After passage through the flasks, the air was trapped in a system of absorption tubes. The first tube was filled with 10 ml ethylene glycol monomethylether (Merck, Darmstadt, Germany) to absorb volatilized substances. The two following tubes were filled with 10 ml of a mixture of ethanolamine and diethylene glycol monobutylether (1:1, Merck, Darmstadt, Germany) to absorb 14 CO2 . With this sequence of absorption tubes a strict separation of volatile 14 Ccompounds and mineralized 14 CO2 is achieved and a correct measurement of the mineralization is ensured. For 14 C counting the absorption liquids were mixed with 10 ml Permablend I scintillation cocktail (22 g Permablend I, Packard, Dreieich, Germany, dissolved in 1 L toluene). Cumulative 14 CO2 in percentage of 14 C applied was plotted as a function of time. Analysis of variance was used to
analyze the different biodegradability of the tested substances. Means were compared by Students– Newman–Keuls test (n ¼ 3, p ¼ 0:05). 2.6. Quantification of nonextractable residues and detection of metabolites in soil The mass balance of radioactive carbon was determined by addition of radioactivity trapped as 14 CO2 , radioactivity volatilized during the experiment and residual radioactivity (nonextractable and extractable) in the soil after the experiments. The soil samples were soxhlet extracted with methanol for 24 h. After extraction aliquots of the soil samples were combusted as described above to determine the amount of nonextractable residues. Radioactivity in the extract was determined by liquid scintillation counting. The extracts were evaporated in a rotary evaporator to dryness and dissolved in a mixture of ethyl acetate and cyclohexane (1:1, chromatography grade, Merck, Darmstadt, Germany). The clean up of soil extracts was performed with an automated gel permeation chromatography system (GPC, Abimed, Langenfeld, Germany). Mobile phase: ethyl acetate:cyclohexane 1:1, flow: 5 ml min1 , column: biobeads S-X3 (Bio-Rad Laboratories GmbH, Munich, Germany). The concentrated GPC fractions were analyzed by HPLC (Merck, Darmstadt, Germany), equipped with a UV/VIS detector and a radioactivity detector LB 507 B (Berthold, Wildbad, Germany). Column: 250 4 mm LiChrospher 100 RP-18, 5 lm (Merck, Darmstadt, Germany). Mobile phase (HPLC grade): A ¼ bidistilled water, B ¼ methanol, C ¼ 0.5% acetic acid. Gradient: T0 : A ¼ 60%, B ¼ 39%, C ¼ 1%; T20 : A ¼ 10%, B ¼ 89%, C ¼ 1%; T25 : A ¼ 10%, B ¼ 89%, C ¼ 1%; T30 : A ¼ 60%, B ¼ 39%, C ¼ 1%; T35 : A ¼ 30%, B ¼ 69%, C ¼ 1%. Flow: 1 ml min1 ; column temperature: 20 °C. 2.7. Degradation of bromoxynil in bacterial enrichment cultures The soil was mixed with a bromoxynil solution (water, 10% methanol), resulting in a water content of 40% WHC and a bromoxynil concentration of 2.2 lg g1 soil dry weight. Then, the soil was
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
incubated for 10 days at room temperature in a water saturated atmosphere. After this adaptation phase, 2 g soil (fresh weight) was transferred into 20 ml of a mineral salt medium containing bromoxynil and a nitrogen source (MS-BO-Nþ ) (NaNO3 0.5 g L1 , K2 HPO4 0.65 g L1 , KH2 PO4 0.17 g L1 , MgSO 4 205 mg L1 , bromoxynil 40 mg L1 ) and incubated at 28 °C. In this enrichment culture, the soil inoculum served as carbon source. After enrichment, 1 ml of this culture was transferred, first, into fresh MS-BO-Nþ , second, into mineral salt medium with an additional carbon source MS-BO-Nþ -Cþ (glucose 500 mg L1 ), and third, into mineral salt medium with an additional carbon source but without nitrogen MSBO-Cþ -N (glucose 500 mg L1 ). The bromoxynil biodegradation in these cultures was recorded by detecting the metabolites in the HPLC-UV/VIS system described above. For this purpose 50 ll of culture medium was injected directly into the HPLC after removing particles by centrifugation (10 min, 7240g) or the culture medium was extracted by solid phase extraction (SPE) described above. Further investigations of formed metabolites were done by analyzing SPE extracts of culture medium in a LC/MS system (ThermoQuest, Egelsbach, Germany, negative ionisation).
3. Results 3.1. Characterization of maize cell wall bound residues About 76% of the radioactivity applied onto the maize plants could be recovered by combustion after 6 days of incubation. As much as 34% of the radioactivity remaining adsorbed or absorbed by the plant was due to nonextractable residues whereas about 66% were extractable with water and organic solvents. The formed nonextractable residues were distributed between the specific cell wall fractions as shown in Fig. 2. The lignin fraction is the main and probably most resistant cell wall fraction, which is involved in the bound residue formation. About 55% of the total formed bound residues were detected in the lignin fraction. The nonextractable
53
Fig. 2. Distribution of bound [14 C]bromoxynil residues in different cell wall fractions of maize plants.
residues in the easily degradable starch, protein and pectin cell wall fractions (8, 24, and 12%, respectively), will not play an important role concerning the persistence of bromoxynil residues in soils. 3.2. Fate of bromoxynil, bromoxyniloctanoate, and maize cell wall bound residues in soil As shown in Fig. 3, mineralization of free applied [14 C]bromoxynil and [14 C]bromoxyniloctanoate started without a lag phase. Within the first 5 days 14 CO2 -production from [14 C]bromoxynil and [14 C]bromoxyniloctanoate increased up to 25 and 30%, respectively. Then mineralization
Fig. 3. Cumulative release of 14 CO2 from [14 C]bromoxynil (14C-BO), [14 C]bromoxyniloctanoate (14C-BOO), and nonextractable [14 C]bromoxynil residues of maize plants (14C-NER) in an agricultural soil.
54
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
rates decreased and within 60 days up to 42% of [14 C]bromoxynil and 49% of [14 C]bromoxyniloctanoate were mineralized (Table 1). The amounts of 14 CO2 released from [14 C]bromoxynil and [14 C]bromoxyniloctanoate after an incubation period of 60 days did not differ significantly. Biomineralization of [14 C]NER occurred with nearly constant mineralization rates (Fig. 3) and was significantly lower than mineralization of the free applied herbicide. After 70 days 21% of [14 C]NER was mineralized, indicating that the soil microorganisms were able to attack the cell wall bound bromoxynil residues and to degrade them to 14 CO2 (Table 1). The extraction of soil material from the experiments with free applied [14 C]bromoxynil and [14 C]bromoxyniloctanoate showed that only a small part of the applied 14 C-substances were extractable (Table 1). After an incubation time of 60 days only 2.9 and 4.4% of the initally applied radioactivity could be extracted in the [14 C]bromoxynil and [14 C]bromoxyniloctanoate degradation experiments, respectively. Consequently, 52.3 and 44.3% of the applied radioactivity could be recovered by combustion as nonextractable residues from [14 C]bromoxynil and [14 C]bromoxyniloctanoate, respectively. Thus, the decreasing mineralization rates could be explained by the decreasing bioavailability in the course of the experiments. The soil residues of the degradation experiments with the plant cell wall bound [14 C] bromoxynil ([14 C]NER) showed a negligible extractability of 1.4% of the applied radioactivity, the main portion (76%) persisted as nonextractable residues in the soil (Table 1). The analysis of the [14 C]bromoxynil metabolites, formed in soil during the incubation time, were
complicated by the low extractability of the remaining residues, due to the strong tendency to form nonextractable residues (Table 1). The extract of the soil from the [14 C]bromoxynil biodegradation experiment was subjected to a GPC cleanup and the main portion of the radioactivity was found in the fractions collected between the retention volumes 150 and 195 ml. This fraction contained 58% of the radioactivity sampled in GPC fractions. In the other fractions the radioactivity was below the detection limit. HPLC analyses of this fraction showed beside the [14 C]bromoxynil itself, only one polar metabolite. To overcome the difficulties of the high formation rate of nonextractable residues in the degradation experiments with soil, the formation of metabolites was investigated in microbial enrichment cultures. In the enrichment cultures with an additional carbon source and bromoxynil as sole source of nitrogen, the formation of two polar metabolites could be observed. HPLC analyses revealed that metabolite 1 had the same retention time as the metabolite that was observed in the soil extract. Further analysis of this metabolite by LC/MS showed a molecular ion of m=z 292/294/296 and the typical isotope distribution pattern of a dibrominated molecule (Fig. 4). Thus, metabolite 1 could be tentatively described as 3,5-dibromo-4-hydroxybenzamide. Metabolite 2 could be identified as 3,5-dibromo-4-hydroxybenzoic acid by comparing the retention time with an authentic reference compound.
4. Discussion From the total bromoxynil residues in plants, 34% nonextractable residues were formed within 6
Table 1 Distribution and mass balance of the radioactivity applied to soil in the biodegradation experiments (in % of Experiment
14
[14 C]Bromoxynila [14 C]Bromoxyniloctanoatea [14 C]NERb
42.4 1.3 49.3 9.4 21.6 8.5
a
60 days incubation. 70 days incubation. c n.d., not detectable. b
CO2
14
C volatile
0.03 0.01 0.06 0.03 n.d.c
14
C nonextractable
52.3 2.3 44.3 3.5 76.5 2.6
14
C extractable
2.9 0.3 4.4 0.9 1.4 0.8
14
C applied) 14
C recovery
97.6 3.4 98.0 13.4 99.5 6.7
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
Fig. 4. Mass spectrum of metabolite 1 in a negative ionization mode.
days of plant growth. The main portion of the nonextractable residues were bound to the lignin fraction, whereas to the more easily degradable cell wall fractions like starch, protein or pectin only minor amounts were bound. The potential adverse long term effects of plant incorporated bromoxynil residues will therefore depend on the degradability of lignin in soil. White rot fungus is known to posses lignolytic enzymes which enable this fungus to degrade lignin in wood. For different soil fungi species like Fusarium [18,19], Trichoderma [20], and Aspergillus [21] the degration of lignin has also been reported. The potentiality for the release of bromoxynil and/or its metabolites from lignin bound residues is therefore present in soil. In the biodegradation experiments, the [14 C] bromoxynil plant cell wall bound residues ([14 C]NER) showed significantly lower mineralization rates than the free applied herbicides bromoxynil and bromoxyniloctanoate. In other
55
experiments with [14 C]isoproturon and plant-incorporated as well as cell wall bound residues of [14 C]isoproturon, the incorporated and bound residues showed also lower mineralization rates than the directly applied herbicide [22,23]. In our study, after an incubation time of 70 days, 21% of the [14 C]NER were completely degraded to 14 CO2 , indicating that the indigenous microbial community was able to release the bound bromoxynil or its metabolites and to mineralize them. Whether the released 14 C-residues were originally bound to the lignin fraction or to the more easily degradable starch, protein or pectin fractions cannot be decided. Since the main portion of the bound 14 Cresidues (55%) was bound to the lignin fraction it can be assumed that the mineralization of the minor 21% portion is probably due to the release of the 14 C-residues bound to the easily degradable cell wall fractions. But for a definite answer to this question further investigations with lignin residues are necessary. The degradation of the free applied herbicides bromoxynil and bromoxyniloctanoate occurred without a lag phase and with high initial mineralization rates, reflecting a good degradability of this herbicide in soils which has been already reported [1,2]. Therefore it can be assumed, that the enzymes which are involved in the mineralization of bromoxynil were already produced by the microorganisms in the soil. Since there was no significant difference in the mineralization of bromoxynil and bromoxyniloctanoate, the cleavage of the ester-binding of the octanoate seems to happen very quickly, as has already been reported in the literature [2], and the following mineralization is referred to the resulting bromoxynil. The decrease of the mineralization rates with time are probably due to a decreasing bioavailability because of formation of nonextractable residues. After 60 days of incubation about 50% of the applied radioactivity was not extractable. The strong tendency for the formation of nonextractable residues can be explained by the reaction of reactive groups within the bromoxynil molecule (hydroxyl group) or its metabolites (amide or carboxyl group) with natural soil components, i.e., humic acids. The low extractability of the 14 Cresidues (2–4% of applied radioactivity) also
56
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
reflects the high transformation rate—mainly mineralization and formation of nonextractable residues—of bromoxynil in soils. Two polar metabolites were observed as degradation products of bromoxynil. One metabolite could be identified as 3,5-dibromo-4-hydroxybenzoic acid and the other was characterized as 3,5-dibromo-4-hydroxybenzamide. As generally known, hydrolysis of the nitrile group is the basic reaction in the transformation of bromoxynil. Principally, the hydrolysis can occur in a one step or a two step process (Fig. 5). Klebsiella pneumonia subsp. ozaenae converts bromoxynil by the enzyme nitrilase directly to the carboxylic acid derivative in the absence of any detectable amide intermediate [7]. Other bromoxynil decomposers like Pseudomonias putida 13XF [4–6] and Flexibacterium BR4 [8] convert the nitrile group in a first step to the amide group by the enzyme nitrile hydratase and in a second step the resulting amide is subsequently hydrolysed to the corresponding carboxylic acid by amidase. In A. radiobacter 8/4, however, no amidase activity was found, since this strain transforms
bromoxynil only to the corresponding amide without further degradation. Bromoxynil degradation not only occurs via hydrolysis of the nitrile group, Topp et al. [9] found a Flavobacterium species to be able to replace the nitrile group with a hydroxyl group, yielding the corresponding chinone by the action of pentachlorophenol hydroxylase (Fig. 5). The results of the present study indicate that in the investigated soil a microbial community was present which was able to mineralize bromoxynil. In the whole process probably various microorganisms were involved, initiating the bromoxynil degradation via hydrolysis of the nitrile group to the corresponding amide and carboxylic acid. The acid was quickly decarboxylized and after ring cleavage the 14 C-ring atoms were completely metabolised to 14 CO2 . No accumulation of the intermediate products could be observed, they were only detected in small amounts. The results of the current study show, that the main degradation and transformation pathways of directly applied bromoxynil in soil were mineralization and formation of bound residues. Plant cell
Fig. 5. Initial degradation reactions of bromoxynil and some enzymes and microorganisms involved, according to the literature [5–9].
P. Rosenbrock et al. / Pesticide Biochemistry and Physiology 78 (2004) 49–57
wall bound residues of bromoxynil were also mineralized but in a lower extent than the free compound. A part of the plant cell wall bound bromoxynil residues could be released in the soil and subjected to complete degradation whereas the main portion persisted as nonextractable residues.
References [1] A.E. Smith, Degradation of bromoxynil in regina heavy clay, Weed Res. 11 (1971) 276–282. [2] R.F. Collins, Perfusion studies with bromoxynil octanoate in soil, Pestic. Sci. 4 (1973) 181–192. [3] D. M€ uller, J. Gabriel, Bacterial degradation of the herbicide bromoxynil by Agrobacterium radiobacter in biofilm, Folia Microbiol. 44 (1999) 377–379. [4] L.A. Golovleva, R.N. Pertsova, F. Kunc, M. Vokounova, Decomposition of the herbicide bromoxynil in soil and in bacterial cultures, Folia Microbiol. 33 (1988) 491–499. [5] M. Vokounov a, O. Vacek, F. Kunc, Effect of glucose and ribose on microbial degradation of the herbicide bromoxynil continuously added to soil, Folia Microbiol. 37 (1992) 128–132. [6] J. Gabriel, J. Vekova, J. Vosahlo, High-performance liquid chromatographic study of the aromatic nitrile metabolism in soil bacteria, J. Chromatogr. B 681 (1996) 191–195. [7] K.E. McBride, J.W. Kenny, D.M. Stalker, Metabolism of the herbicide bromoxynil by Klebsiella pneumoniae subsp. ozaenae, Appl. Environ. Microbiol. 52 (1986) 325–330. [8] A.E. Smith, D.R. Cullimore, The in vitro degradation of the herbicide bromoxynil, Can. J. Microbiol. 20 (1974) 773–776. [9] E. Topp, L. Xun, C.S. Orser, Biodegradation of the herbicide bromoxynil (3,5-dibromo-4-hydroxybenzonitrile) by purified pentachlorophenol hydroxylase and whole cells of Flavobacterium sp. strain ATCC 39723 is accompanied by cyanogenesis, Appl. Environ. Microbiol. 58 (1992) 502–506. [10] B. Graß, H. Mayer, J. Nolte, G. Preuß, N. Zullei-Seibert, Studies on the metabolism of hydroxybenzonitrile herbicides: I. mass spectrometric identification, Pest. Manag. Sci. 56 (2000) 49–59.
57
[11] M. Millet, W.-U. Palm, C. Zetzsch, Abiotic degradation of halobenzonitriles: investigation of the photolysis in solution, Ecotoxicol. Environ. Saf. 41 (1998) 44–50. [12] I. Texier, C. Giannotti, S. Malato, C. Richter, J. Delaire, Solar photodegradation of pesticides in water by sodium decatungstate, Catal. Today 54 (1999) 297–307. [13] B. Schaller, B. Schneider, H.R. Sch€ utte, Metabolism of the herbicide bromoxynil in Hordeum vulgare and Stellaria media, Z. Naturforsch. C 47 (1992) 126–131. [14] D.M. Stalker, K.E. McBride, L.D. Malyi, Herbicide resistance in transgenic plants expressing a bacterial detoxification gene, Science 242 (1988) 419–423. [15] G. Freyssinet, B. Pelissier, M. Freyssinet, R. Delon, Crops resistant to oxynils: from the laboratory to the market, Field Crops Res. 45 (1996) 125–133. [16] W. Ebing, A. Haque, I. Schuphan, Ecochemical assessment of environmental chemicals: draft guideline of the test procedure to evaluate metabolism and degradation of chemicals by plant cell culture, Chemosphere 13 (1984) 947–957. [17] D. Komoßa, I. Gennity, H. Sandermann Jr., Plant metabolism with C–P bonds: glyphosate, Pestic. Biochem. Physiol. 43 (1992) 85–94. [18] D.M. Norris, Degradation of 14 C-labeled lignins and 14 Clabeled aromatic acids by Fusarium solani, Appl. Environ. Microbiol. 40 (1980) 376–380. [19] J.B. Sutherland, A.L. Pometto, D.L. Crawford, Lignocellulose degradation by Fusarium species, Can. J. Bot. 61 (1983) 1194–1198. [20] T.W. Flegel, V. Meevootisom, S. Kiatpapan, Indications of lignolysis by Trichoderma strains isolated from soil during simultaneous screening for fungi with cellulase and laccase activity, J. Ferment. Technol. 60 (1982) 473–475. [21] K.L. Kadam, S.W. Drew, Study of lignin biotransformation by Aspergillus fumigatus and white-rot fungi using 14 Clabeled and unlabeled kraft lignins, Biotech. Bioeng. 28 (1986) 394–404. [22] S. Lehr, I. Scheunert, F. Beese, Mineralization of free and cell-wall-bound isoproturon in soils in relation to soil microbial parameters, Soil Biol. Biochem. 28 (1996) 1–8. [23] S. von Wiren-Lehr, I. Scheunert, U. D€ orfler, Mineralization of plant-incorporated residues of 14 C-isoproturon in arable soils originating from different farming systems, Geoderma 105 (2002) 351–366.