Journal of Environmental Science and Health, Part B ...

This article was downloaded by: [RWTH Aachen University] On: 06 February 2014, At: 01:27 Publisher: Taylor & Francis Informa Ltd Registered in England and Wales Registered Number: 1072954 Registered office: Mortimer House, 37-41 Mortimer Street, London W1T 3JH, UK

Journal of Environmental Science and Health, Part B: Pesticides, Food Contaminants, and Agricultural Wastes Publication details, including instructions for authors and subscription information: http://www.tandfonline.com/loi/lesb20

A correlation between the fate and non-extractable residue formation of activities in soil a

14

C-metalaxyl and enzymatic

a

a

a

Jens Botterweck , Daniela Claßen , Thordis Zegarski , Christian Gottfroh , Roshni b

a

b

Kalathoor , Andreas Schäffer , Jan Schwarzbauer & Burkhard Schmidt a

a

Institute for Environmental Research , RWTH Aachen University , Aachen , Germany

b

EMR RWTH Aachen University, Institute of Petroleum & Coal , Structural Geology , Aachen , Germany Published online: 11 Dec 2013.

To cite this article: Jens Botterweck , Daniela Claßen , Thordis Zegarski , Christian Gottfroh , Roshni Kalathoor , Andreas Schäffer , Jan Schwarzbauer & Burkhard Schmidt (2014) A correlation between the fate and non-extractable residue 14

formation of C-metalaxyl and enzymatic activities in soil, Journal of Environmental Science and Health, Part B: Pesticides, Food Contaminants, and Agricultural Wastes, 49:2, 69-78, DOI: 10.1080/03601234.2014.844600 To link to this article: http://dx.doi.org/10.1080/03601234.2014.844600

PLEASE SCROLL DOWN FOR ARTICLE Taylor & Francis makes every effort to ensure the accuracy of all the information (the “Content”) contained in the publications on our platform. However, Taylor & Francis, our agents, and our licensors make no representations or warranties whatsoever as to the accuracy, completeness, or suitability for any purpose of the Content. Any opinions and views expressed in this publication are the opinions and views of the authors, and are not the views of or endorsed by Taylor & Francis. The accuracy of the Content should not be relied upon and should be independently verified with primary sources of information. Taylor and Francis shall not be liable for any losses, actions, claims, proceedings, demands, costs, expenses, damages, and other liabilities whatsoever or howsoever caused arising directly or indirectly in connection with, in relation to or arising out of the use of the Content. This article may be used for research, teaching, and private study purposes. Any substantial or systematic reproduction, redistribution, reselling, loan, sub-licensing, systematic supply, or distribution in any form to anyone is expressly forbidden. Terms & Conditions of access and use can be found at http:// www.tandfonline.com/page/terms-and-conditions

Journal of Environmental Science and Health, Part B (2014) 49, 69–78 C Taylor & Francis Group, LLC Copyright ISSN: 0360-1234 (Print); 1532-4109 (Online) DOI: 10.1080/03601234.2014.844600

A correlation between the fate and non-extractable residue formation of 14C-metalaxyl and enzymatic activities in soil JENS BOTTERWECK1, DANIELA CLAßEN1, THORDIS ZEGARSKI1, CHRISTIAN GOTTFROH1, 1 ¨ ROSHNI KALATHOOR2, ANDREAS SCHAFFER , JAN SCHWARZBAUER2 and BURKHARD SCHMIDT1 1

Institute for Environmental Research, RWTH Aachen University, Aachen, Germany EMR RWTH Aachen University, Institute of Petroleum & Coal, Structural Geology, Aachen, Germany

Downloaded by [RWTH Aachen University] at 01:27 06 February 2014

2

Extracellular, oxidative soil enzymes like monophenol oxidases and peroxidases play an important role in transformation of xenobiotics and the formation of organic matter in soil. Additionally, these enzymes may be involved in the formation of non-extractable residues (NERs) of xenobiotics during humification processes. To examine this correlation, the fate of the fungicide 14C metalaxyl in soil samples from Ultuna (Sweden) was studied. Using different soil sterilization techniques, it was possible to differentiate between free, immobilized, and abiotic (“pseudoenzyme”-like) oxidative activities. A correlation between the formation of metalaxyl NER and soil organic matter content, biotic activities, as well as extracellular phenoloxidase and peroxidase activities in the bulk soil and its particle size fractions was determined. Extracellular soil-bound enzymes were involved in NER formation (up to 8% of applied radioactivity after 92 days) of the fungicide independently from the presence of living microbes and different distributions of the NER in the soil humic subfractions. Keywords: Metalaxyl, soil, extracellular soil enzymes, non-extractable residues, metabolism.

Introduction Anthropogenic chemicals, e.g., pesticides, are released in significant amounts to the lithosphere, hydrosphere, and atmosphere due to industrial and agricultural practice. Soil as a complex medium with a large number of interaction sites is a major sink for these xenobiotic substances.[1] In the soil, they may be volatilized or leached to the groundwater, taken up by living organisms, and degraded by microorganisms and abiotic processes.[2] Microbial degradation of organic contaminants in soil results in the formation of metabolites, microbial biomass, mineralization products (CO2 and H2 O), as well as bound or non-extractable residues (NERs).[3,4] Microbial activity and organic matter content play a major role in the formation of NER in soil.[5–7] It is suggested that extracellular oxidoreductases like monophenol oxidases and peroxidases may lead to formation of NER of organic contaminants in soil during humification.[7,8] These enzymes are known to oxidize phenolics to aryloxy radicals, which then polymerize to form insoluble humic acid-like complexes during natural humification processes[9–13] and may therefore Address correspondence to Jens Botterweck, Institute for Environmental Research, RWTH Aachen University, Worringerweg 1, 52074, Aachen, Germany; E-mail: [email protected] Received June 5, 2013.

also act as microbial-derived catalysts creating ether and carbon–carbon linkages between xenobiotics and the soil matrix.[14] Like all extracellular enzymes in soil, they can be associated with active cells (animal, plant, and microorganisms), dead cells, cell debris, as well as complexed with clay minerals and humic colloids.[15] These immobilized or soil-bound enzymes may become readily inactivated due to the adverse soil conditions, e.g., protease activities, whereas the enzymes bound to soil organic matter (SOM) and clay minerals and tannins persist in soil with remaining catalytic activities depending on the specific enzyme.[16–19] Mineral components in soil support the adsorption and binding of organic chemicals (enzymes and substrates) and act as a reactive surface for the oxidation of organic compounds, catalyzed by, e.g., oxides/hydroxides of Fe3+, Mn3+, or Mn4+.[20,21] These “pseudoenzyme” activities may also lead to NER formation in the soil ecosystem.[22] Until now, the contribution and localization of extracellular enzymes in soil involved in the formation of nonextractable xenobiotics residues are hardly understood. The use of soil sterilization techniques is one possibility to distinguish between different localizations and sources of oxidative activities in soil. The efficiency and impact of different soil sterilization methods on the biological, chemical, and physical properties of soils have previously been studied by a number of researchers and have been reviewed.[23] It was concluded that every sterilization technique may alter the chemical and physical properties of soil, and the best


70 method to be applied will depend on the research being intended.[23] Metalaxyl [N-(2,6-dimethylphenyl)-N-(2-methoxyacetyl)-alanine methyl ester] belongs to the group of systemic acylanilide fungicides used against downy mildews, late blight, and damping off of root stem and fruit roots in many agricultural crops.[24] Correlations between the fate of metalaxyl in soil and its microbial activity, clay and organic matter content, as well its abiotic catalysts have been investigated since years.[25–30] The aim of the present study was to establish modified soil sterilization techniques to measure and distinguish between different sources of extracellular phenoloxidase and peroxidase activities in a Northern-European soil from Ultuna (Sweden) and its “pseudoenzyme” activities. Specifically, we investigated the influence of these properties on the fate and persistence of 14C-metalaxyl in soil in a time course study for up to 92 days under three conditions: (i) soil samples with no microbial and extracellular enzymatic acitivities; (ii) soil samples without microbial activities but remaining extracellular enzyme activities; and (iii) native bulk soil.

Materials and methods Chemicals Fungicide metalaxyl (N-(methoxyacetyl)-N-(2,6-xylyl)DL-alaninate) was used as (ring-U-14C) labeled radioactive compound (1.13368 MBq/mmol; radioactive purity > 99.9%). The substance was synthesized and provided by the Isotopes Co. (Izotop, Budapest, Hungary); non-labeled metalaxyl used as unlabeled reference compound (99.9% purity) was provided by Dr. Ehrenstorfer (Augsburg, Germany). Metalaxyl acid was synthesized from non-labeled metalaxyl as described.[31]

Botterweck et al. Soil sterilization In order to obtain soil samples without any enzymatic activity, 50 g of the air-dried soil was treated with γ -irradiation (25 kGy for 60 min; beta gamma service BGS; Wiehl, Germany) and subsequently autoclaved two times for 60 min at 121◦ C. The resulting soil samples were kept aseptically and stored in a refrigerator for 2–3 days at 4◦ C, prior to incubation. Each sterilized sample was mixed with 1.08 mL phenylhydrazine solution (68 mM), as inhibitor of oxidative enzymes.[36] Then, the samples were adjusted to 60% of their maximum water holding capacity, using ultrapure water (Millipore equipment, Merck Chemicals GmbH, Schwalbach, Germany) sterilized by autoclaving 5% (v/w) sodium azide.[23] For soil samples without active microorganisms but remaining extracellular enzyme activities, 50 g of soil was fumigated with chloroform in a desiccator as described.[37] The samples were aseptically closed and stored in a refrigerator for one day at 4◦ C, prior to incubation. Then, the samples were mixed with 50 mg of solid mercury chloride according to a published procedure.[38] Another set of samples was only treated with γ irradiation without further addition of inhibitors or sterilants. The sterilization treatments and the corresponding designations of the samples as well as the resulting impacts subsequently with 14 C-metalaxyl incubation are summarized in Table 1. The efficiency of each sterilization procedure was checked prior to incubation experiments by use of growth media plates (standard nutrient medium 1, Carl Roth GmbH, Karlsruhe, Germany; Czapek-Dox medium, DIFCO/BD, Heidelberg, Germany), which were inoculated with 1,000 µL of a soil extract made by using 50 mL phosphate buffer per 5 g of soil. Additionally, the efficiency of sterilization was tested by the use of the DMSO reduction assay.[39]

Enzymatic assays Soil The soil used was derived from an experimental field in Ultuna, near Uppsala, Sweden. The field has been part of a long-term experiment, initiated in 1956 and is planted annually with different kinds of crops.[32,33] The samples used in the present study were taken from the Ap horizon, 0–17 cm. Prior to use, the soil was air-dried, sieved through a 2 mm mesh, and was kept at 4◦ C in darkness for 2 months. The soil characteristics were determined recently.[34,35] Crucial parameters are: C:N ratio: 10; maximum water holding capacity (WHCmax ): 17.5%; cation exchange capacity (CEC): 180 cmol/kg; sand: 23.8%; silt: 52.6%; clay: 23.6%; organic carbon of the bulk soil: 25.9 g/kg; organic carbon of the sand fraction: 3.0 g/kg; organic carbon of the silt fraction: 17.3 g/kg; organic carbon of the clay fraction: 4.6 g/kg.

Extracellular p-diphenoloxygen reductase (EC 1.10.3.2) activities, in the following designated as phenoloxidase activities (PO), were measured colorimetrically using 2,2 azino-bis(3-ethylbenzothiazoline-6-sulphonic acid (ABTS) as substrate as described.[40] The oxidation rate, i.e., the transformation of ABTS to radical cation ABTS+., was measured at 420 nm. Blank samples and substrate calibration were done as described.[40] Phenoloxidase activities were expressed in units defined as µmoles of ABTS+. formed from ABTS per minute (U) and g of dry matter (U/g DM). Extracellular peroxidase activities (PERO) (EC 1.11.1) were measured according to a published procedure with modifications described next.[41] First, 4 g fresh weight of each of the soil samples was mixed with 196 mL of cold acetate buffer (5◦ C, 50 mM, pH 5) in glass beakers for 1 h on a horizontal shaker. After shaking, 250 µL of the mixture

71

Correlation between the fate and non-extractable residue formation

Table 1. Treatment of the soil samples prior to incubation with 14C-metalaxyl, abbreviations of the respective assays, and impact for incubation. Soil treatment γ -Irradiation + autoclaving + phenylhydrazine + sodium azide treatment γ -Irradiation


Chloroform fumigation + mercury chloride treatment Ultuna soil without treatment Annealed seasand without treatment

Abbreviation Soila-γ -N3/phenhyd Soilγ -only SoilHgCl2/CHCl3 Soilnative Sandcontrol

was transferred into 2 mL Eppendorf tubes and mixed with 0.5 mL of preheated (25◦ C) tetramethylbenzidine (TMB) reagent (SIGMA, Aldrich, Steinheim, Germany). The peroxidase reaction was terminated by adding 1.2 mL of 0.2 M sulphuric acid after 5 min. After centrifugation, the absorbance of the supernatants was measured at 450 nm as described.[41] Calibration of the TMB reagent was done accordingly.[41] Peroxidase activities were expressed in units defined as µmoles per minute (U) and g of dry matter (U/g DM). Dimethylsulfoxide reduction assay Microbial activity of the soil samples was determined by means of the dimethylsulfoxide (DMSO) reduction assay as published previously.[39] Four aliquots (1 g fresh weight) of the soil sample were separately transferred into brown glasses (25 mL) and 200 µL of a 5% aqueous DMSO solution was added. The vessels were closed tightly and incubated at 27◦ C in the dark for 24 h. For measurement, 100 µL of the gas phase was removed with a gas-tight syringe and injected into a GC-MS system for analysis, which was calibrated with gaseous dimethylsulfide (DMS) standard dilutions containing 16, 33, and 66 µmol of DMS.

Impact Reduced oxidative activities of intra- and of extracellular enzymes Reduced oxidative activities fof intra- but not of extracellular enzymes Reduced oxidative activities fof intra- but not of extracellular enzymes No reduction of oxidative enzymes No reduction of oxidative enzymes

control samples were wrapped in aluminium foil and incubated at 15◦ C in the dark. Each incubation experiment was done in triplicate. The samples were analyzed 10 and 92 days after application. An aliquot of the soil was removed to determine by combustion analysis the total amount of 14C metalaxyl remaining in the assays. Subsequently, the samples were fractionated into the particle size fractions of sand (63 µm up to 2 mm), silt (2 µm up to 63 µm), and clay (below 2 µm) supported by 110 J/g soil of ultrasonication energy input according to a published method.[42] The obtained fractions were exhaustively extracted under Soxhlet conditions (methanol as solvent, 6 h). The total water used for particle size fractionation was acidified to pH 2 (HCl, 37%) and extracted using ethyl acetate (three times, 100 mL each). Aliquots of the extracted fractions were combusted to determine the amount of non-extractable radioactivity. The extracted silt and clay fractions were further fractionated into non-humics, fulvic acid, humic acid, and humine fractions according to the established alkaline separation method.[43] The amounts of radioactivity in extracts and separated fractions were measured by liquid scintillation counting.

Analytical methods Incubation experiments Prior to application, 50 g of soil contained in one assay was mixed with 50 g of pure annealed sea sand for a better handling of the soil samples. The same sand was also used for the control samples (sandcontrol ). Subsequent metalaxyl application was based on the maximum single application rate (600 g per ha). Concerning soil depth (5 cm) and soil density (1.5 g/cm3), this corresponded to 40 µg of metalaxyl per 50 g of soil. Aliquots of 41.17 µg (0.15 MBq) of metalaxyl in 11.8 mL of deionized water were thus applied to the soil samples in individual 250 mL glass beakers to adjust the final moisture content to 60% of WCHmax. The water containing 14C-metalaxyl was mixed with the soil by intensively stirring for 10 min using a glass rod. Each incubation flask was then equipped with an adsorption device (100 mm × 10 mm) filled with 15 g of soda lime in order to capture evolving 14CO2 from the samples. Samples and

Liquid samples were examined by liquid scintillation counting (LSC) using a Canberra Packard 2250 Tricarb counter (Dreieich, Germany). Solid samples were combusted in Combusto Cones Flexible 1000 (CanberraPackard, Rodgau, Germany) by means of a Biological Oxidizer OX500 (Harvey Instruments/Zinsser, Frankfurt, Germany); the resulting samples were examined by LSC. Extracts were analyzed by radio-thin-layer chromatography (RTLC) on silica gel plates (SIL G-25 UV254 , thick¨ ness: 0.25 mm; Macherey-Nagel, Duren, Germany) using ethyl acetate containing 0.1% formic acid (v/v) as solvent system. The distribution of 14C on the developed RTLC plates was determined using a Fujifilm BAS-1000 Bioimager including Fuji Imaging Plates Type BAS-MS (508) 20 × 40 cm (Raytest, Straubenhardt, Germany). For conventional high-performance liquid chromatography (HPLC), a Beckman System Gold Personal


72 chromatograph with a Programmable Solvent Module 126 ¨ and a Diode Array Detector Module 168 (Munchen, Germany) was used. For radio-HPLC (RHPLC), a Ramona 5 radiodetector equipped with a 1,655 quartz cell (glass, 32–45 µm; internal diameter: 4.0 mm; volume: 0.4 mL; Raytest) was used. Analyses were performed at 25◦ C on a Nucleodur 100- 5 C18 column (5 µm × 4 mm × 250 mm; Macherey-Nagel) with 1 mL min−1 flow rate and 100 µL of injection volume. Non-labeled references metalaxyl and metalaxyl acid were detected at 209 nm. Elution was done with solvents A, bidistilled water with 0.1% formic acid (v/v) and B, acetonitril with 0,1% formic acid (v/v) as follows: A/B 90:10 (v/v) for 10 min, linear 15 min gradient to 90% B, isocratic B (90%) for 10 min, and return to initial conditions within 10 min. Radio-HPLC data calculation was performed by GinaStar software, version 4.08. DMSO samples were measured by gas chromatography mass spectrometry (GC-MS) by an Agilent 6890 system (Agilent, Waldbronn, Germany) with an FS-Supreme 5, 5% phenylpolysilylphenyloxan capillary column under isothermic conditions (50◦ C). The samples were detected by an Agilent MSD detector. Crucial parameters were: flow rate: 1 mL/min; split: 16. 4:1; carrier gas: helium; electron multiplier voltage: 1294.1 V; mass range: 40.0–100.0 m/z). Data calculation was performed by ChemStation software (Ag¨ ilent Technologies GmbH, Boblingen, Germany, version D.01.02.16).

Statistical analysis Each experiment presented in the present study was done in triplicate, and all data are given in mean values and corresponding standard deviations.

Results and discussion Microbial and enzymatic activities in soil The effectiveness of the different sterilization treatments utilized was examined by plating aliquots of the sterilized samples on growth media. However, counting of colonies on plates only targets culturable microorganisms, which represent only a certain amount of the microorganisms present in soil. Thus, the overall microbial activity was additionally checked by measurement of DMSO reduction rates during the incubation experiments with 14C metalaxyl. The corresponding data are shown together with the extracellular phenoloxidase and peroxidase activities in Table 2. Highest DMSO reduction rates (331.9 ng DMS per hour and g dry weight [DW]) and extracellular phenoloxidase (PO) (13.2 mU per g DW) and peroxidase (PERO) activities (50.0 mU per g DW) were determined in native bulk soil 92 days after application of 14C-metalaxyl (Table 2). Soil

Botterweck et al. samples sterilized by γ -irradiation or mercury chloride application and additional CHCl3 fumigation showed lower rates of DMSO reduction after 92 days of incubation (43.2 and 49.0 ng DMS per hour and g DW). As compared to native bulk soil, the extracellular oxidative enzyme activities in these samples were about half of those of native bulk soil enzymatic activities (PO 7.3 and 5.0 mU per g DW; PERO 17.2 and 19.8 mU per g dry weight, respectively) (Table 2). The DMSO reduction rates in soil samples sterilized by autoclaving and γ -irradiation with additional application of sodium azide and phenylhydrazine were the lowest as compared to all the other treatments. DMSO reduction rate was reduced to 16.0 ng DMS per g DW and hour in these samples (92 days after application). The enzymatic activities in these sterilized soil samples were also lowest amounting to PO: 1.1 mU per g DW and PERO: 4.5 mU per g DW (Table 2). As compared to native bulk soil, all three differently sterilized samples, thus, showed very low activities in DMSO reduction, similar to the control sample containing annealed sea sand (Table 2). These results demonstrated that the respective microbial and enzymatic conditions after sterilization were reduced throughout the entire time course study of up to 92, however to different extents. The remaining low amounts of DMS produced in almost all sterile soil samples may be explained by photoreactions, as suggested recently, although care was taken to prevent exposure of the samples to light in the present study.[39] This assumption is corroborated by the finding that the sand control samples also showed similar low DMSO reduction rates. Another explanation is that a small part of the microbial community survived sterilization treatment because DMSO reduction rates increased slightly up to 92 days of incubation in all sterile samples. However, remaining DMSO reduction rates were negligible in absolute terms as compared to those resulting from native bulk soil [331.9 ng]. In contrast, about half of the extracellular oxidative enzyme activities (PO, PERO) of bulk soil remained after the respective sterilization procedure in samples soily-only and soilHgCl2/CHCl3 whereas these were almost completely eliminated in samples treated with autoclaving and γ -irradiation and subsequent application of known inhibitors of respiration and oxidative reactions (soila-y-N3/phenyd ). However, γ -irradiation did not completely delete extracellular phenoloxidase [5 mU] and peroxidase [19.8 mU] activities. It was demonstrated previously that soil is sterilized under such conditions (γ -irradiation) but enzyme activities remained probably due to soil-bound enzymes or endogenous oxidative enzymes released from lysed cells during irradiation.[44,45] Such processes were also observed in the case of phosphatase activity.[46] The amount of enzyme activity remaining after treatment in the samples soilHgCl2/CHCl3 and soilγ -only was about half of that in native bulk soil. This indicates that both treatments (HgCl2 /CHCl3 or γ -irradiation) resulted in quite similar amounts of remaining extracellular oxidative enzyme. The reproducible and stable assays in

73

Correlation between the fate and non-extractable residue formation

Table 2. Dimethylsulfoxide (DMSO) reduction, extracellular phenoloxidase (PO), and extracellular peroxidase (PERO) activities in soil after 10 and 92 days of incubation. Designation (incubation period, days)


Soilnative (10) Soilnative (92) SoilHgCl2/CHCl3 (10) SoilHgCl2/CHCl3 (92) Soily-only (10) Soily-only (92) Soila-y-N3/phenhyd (10) Soila-y-N3/phenhyd (92) Sandcontrol (10) Sandcontrol (92)

DMSO reduction (ng DMS/h−1 g−1 DW)

PO activity (mU g−1 DW)

PERO activity (mU g−1 DW)

232.6 ± 95.4 331.9 ± 19.2 22.6 ± 4.8 43.2 ± 16.7 35.4 ± 19.6 49.0 ± 23.6 13.0 ± 3.1 16.0 ± 3.1 19.6 ± 4.5 16.2 ± 6.9

13.2 ± 1.3 13.3 ± 0.6 5.1 ± 0.3 7.3 ± 0.9 2.2 ± 0.1 5.0 ± 0.2 0.7 ± 0.2 1.1 ± 0.1 1.0 ± 0.2 1.0 ± 0.2

36.1 ± 3.8 42.0 ± 8.1 24.9 ± 4.0 17.2 ± 5.3 23.4 ± 6.8 19.8 ± 0.9 3.4 ± 0.3 4.5 ± 0.3 2.9 ± 1.7 3.1 ± 1.2

Data shown are mean values from three triplicates with corresponding standard deviations DW = dry weight; other abbreviations according to Table 1.

the present study were recently shown to target exclusively extracellular oxidative soil enzymes.[40,41] It is remarkable that these activities did not change much during the entire incubation period and we believe that they were based on soil-bound enzymes, protected and stabilized by immobilization on soil colloid surfaces against photodegradation or denaturation; such processes were recently described.[47] Most likely, these activities are not related to residual enzymes in cell wall fragments or enzymes released from lysed cells because they would be rapidly degraded, since they are not protected and sorbed to the soil.[15] However, slight increases in the PO and PERO activities in the treatments (HgCl2 /CHCl3 or γ -irradiation) may have been due to surviving microbes which produced new extracellular oxidative enzymes. The low oxidative enzyme activities, as measured by the substrates ABTS and TMB in the soila-y-N3/phenhyd samples, did not further decrease during the incubation period. The remaining activities may consequently be traced back to abiotic factors, e.g., the catalysis by ferric oxides and hydroxides in the soil.[22] These activities have recently been described as “pseudoenzyme activities” or “enzyme-like” and are independent from the presence of microflora or microfauna in the soil.[17] The high oxidation activities in native bulk soil comprise “pseudoenzyme” abiotic activities, activities of enzymes immobilized on soil particles, and enzymes associated to and released from active microbial cells. In summary, the experiments could distinguish between different origins of oxidative enzymatic activities in soil using different sterilization treatments as displayed in Figure 1. However, the methods used certainly have noticeable limitations. Firstly, the different sterilization procedures may alter soil parameters, which directly interact with examined enzyme activities; these are, for instance, changes in soil colloid surfaces and soil solution composition and ionic strength.[23] Secondly, the sterilization treatment (irradiation, mercuric chloride, and chloroform fumigation)

may also deactivate portions of the pool of extracellular oxidative enzymes despite stabilization by sorption to soil. Mercuric chloride and chloroform fumigation did not deactivate the oxidative enzymes as shown by an additional experiment: MUB buffer solution (10 mL) containing laccase (1 U) as representative soil phenoloxidase and MUB buffer solution (10 mL) containing lignin peroxidase (1 U) as representative soil peroxidase were treated with mercuric chloride at a concentration similar to that used in soil incubation experiments and fumigated with chloroform. After the treatments, ABTS (0.1 g) was added to the laccase and TMB (1.0 mL) was added to the peroxidase solution. An immediate substrate oxidation was observed in both cases indicating that the oxidative enzyme activities were not reduced by the sterilization techniques.

Fig. 1. Amount and sources of oxidative activities and their amounts in native bulk soil (soilnative ), soil sterilized by γ irradiation (soilγ -only), and soil sterilized by autoclaving + γ irradiation + treatment by sodium azide and phenylhydrazine (soilγ -Na3-Phenyd ).

74

Botterweck et al.

The distributions of extractable, non-extractable, and mineralized radioactivity derived from 14C-metalaxyl are shown in Figure 2. In case of all samples, highest amounts of radioactivity with both incubation periods were found in the water used for fractionation of the soil into the particle size fractions clay, silt, and sand; this portion amounted to more than 65% of applied radioactivity in case of native bulk soil and to about 80% in case of the sterilized samples (Fig. 2). Amounts of radioactivity extracted from the particle size fractions (Soxhlet) were similar for all treatments (ca. 10% of applied radioactivity) (Fig. 2). Slightly lower amounts were extracted from the samples incubated for 92 days compared to the 10-day incubations. The total of radioactivity in both, the fractionation water and the Soxhlet extracts, was defined as extractable radioactivity of the respective sample. The amount of NER contained in the 92-day assays was highest in native bulk soil (about 25% of applied radioactivity); the same holds for mineralized amounts of applied 14C-metalaxyl (about 5% of applied radioactivity) (Fig. 2). The second highest amount of NER was detected in the 92-days assays sterilized by γ -irradiation and in that sterilized using mercury chloride and CHCl3 fumigation: both fractions amounted to about 7% of applied radioactivity (Fig. 2). The lowest amount of non-extractable radioactivity was measured in the soils treated with autoclaving, γ -irradiation, sodium azide, and phenylhydrazine (less than 4% of applied radioactivity after 92 days of incubation; Fig. 2). The portions of NER increased with time in all samples except in the annealed sea sand samples (sandcontrol ) which served as controls: the amount of NER was below 2% of the applied amount after 10 days and did not further increase. Mineralization of metalaxyl in all sterilized soil samples was negligible (below 1% of applied radioactivity) as compared to native bulk soil. 110 100 % of applied radioacvity


Distribution of radioactivity

90 80 70 60 50 40 30 20 10 0

10 92 soilnative NER

10 92 soilHgCl2/CHCl3

ER Fraconaon water

10 92 soil -only

10 92 10 92 soilya- -N3-phenyd sandcontrol

sum of ER sand/silt/clay

Mineralisaon

Fig. 2. Distribution of extractable, mineralized, and nonextractable radioactivity in soil 10 days (left bars) and 92 days after application (right bars). Data are mean values from three triplicates. Abbreviations are according to Table 1.

The total recovery of radioactivity in all samples ranged between 95 and 105% of the applied amount. Recently performed and published studies pointed to a correlation between microbial activity and formation of non-extractable residues of metalaxyl in soil.[25,26] The present study clearly underlines these findings for both, the bulk soil and its particle size fractions. The present microbial activities and extracellular oxidative enzyme activities were closely correlated with the amounts of non-extractable residues formed increasing in the order soila-y-N3/phenhyd < soilHgCl2/CHCl3 < soily-only < soilnative . The assays soilHgCl2/CHCl3 and soily-only with inactivated microbes but remaining enzyme activities formed higher amounts of than the assay in which both activities were almost completely inhibited (soilauto-y-N3/phenhyd ). Obviously, both oxidative enzymes, phenoloxidases and peroxidases, support the degradation and binding of metalaxyl in soil. The residues formed remained in the soil after subsequent, exhaustive extractions by water and organic solvents even after an ultrasonication suggesting strong linkages of metalaxyl residues to the soil. According to published pesticide studies,[48] we, thus, assume that in the present experiments, metalaxyl is covalently bound to humic matter by these oxidative enzymes via generation of radicals as they occur during humification processes. Thus, soil-bound oxidative enzymes seem to support NER formation to a remarkable degree. This correlation observed is in agreement with previously published studies showing that most of all potential extracellular soil enzymatic activities originated from adsorbed enzymes.[49] In Ultuna soil, enzyme activities and metalaxyl-derived NER were most abundant in the silt fraction. This was due to the high organic matter content of this fraction and its stabilization by adsorption on mineral surfaces as described previously.[33] The NER were not only correlated with living microbes and soil-bound extracellular enzyme activities, because they were also formed in the absence of active microbes and active extracellular phenoloxidases and peroxidases. Abiotic influences leading to NER, e.g., by metal oxides and hydroxides, were studied to some extent.[50,51] Correspondingly, we assume that portions of NER of metalaxyl in the samples with completely denatured enzymes and nearly without microbial activity were based on such abiotic reactions at clay surfaces known to catalyze a number of transformation reactions including hydrolysis, reduction, oxidation, oligo- and polymerisation, and rearrangement.[50,51] The present study revealed low mineralization rates of metalaxyl (