Isolation and characterisation of azoxystrobin degrading bacteria from soil

Chemosphere 95 (2014) 370–378

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Isolation and characterisation of azoxystrobin degrading bacteria from soil Christopher C. Howell a,⇑, Kirk T. Semple b, Gary D. Bending a a b

School of Life Sciences, University of Warwick, Gibbet Hill Road, Coventry, Warwickshire CV4 7AL, UK Lancaster Environment Centre, Lancaster University, Lancaster LA1 4YQ, UK

h i g h l i g h t s  We studied the degradation of azoxystrobin by enriched bacterial communities.  Azoxystrobin-degrading bacterial strains CCH1 and CCH2 were isolated.  Strains CCH1 and CCH2 also degraded three other strobilurin compounds.

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Article history: Received 28 May 2013 Received in revised form 22 August 2013 Accepted 10 September 2013 Available online 11 October 2013 Keywords: Azoxystrobin Degradation Cupriavidus sp. Rhodanobacter sp. T-RFLP

a b s t r a c t The first strobilurin fungicides were introduced in 1996, and have since been used in a vast array of disease/plant systems worldwide. The strobilurins now consist of 16 compounds and represent the 2nd most important fungicide group worldwide with 15% of the total fungicide market share. Strobilurins are moderately persistent in soil, and some degradation products (e.g. azoxystrobin acid) have been detected as contaminants of freshwater systems. Little is currently known about the transformation processes involved in the biodegradation of strobilurins or the microbial groups involved. Using sequential soil and liquid culture enrichments, we isolated two bacterial strains which were able to degrade the most widely used strobilurin, azoxystrobin, when supplied as a sole carbon source. 16S rRNA showed that the strains showed homology to Cupriavidus sp. and Rhodanobacter sp. Both isolated strains were also able to degrade the related strobilurin compounds trifloxystrobin, pyraclostrobin, and kresoxim-methyl. An additional nitrogen source was required for degradation to occur, but the addition of a further carbon source reduced compound degradation by approximately 50%. However, 14C radiometric analysis showed that full mineralisation of azosxystrobin to 14CO2 was negligible for both isolates. 16S rRNA T-RFLP analysis using both DNA and RNA extracts showed that degradation of azoxystrobin in soil was associated with shifts in bacterial community structure. However, the phylotypes which proliferated during degradation could not be attributed to the isolated degraders. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction The strobilurin fungicides represent one of the most important groups of pesticides currently in use worldwide for the control of fungal crop pathogens. The first strobilurin compounds, azoxystrobin and kresoxim-methyl, were released in 1996. By 2002, a further six compounds (metominostrobin, trifloxystrobin, picoxystrobin, pyraclostrobin, famoxadone, and fenamidone) had been released (Bartlett et al., 2002), followed by fluoxastrobin and dimoxystrobin in 2003 (Suty-Heinze et al., 2004; Balba, 2007) and orysastrobin in 2006 (van Ravenzwaay et al., 2007). Currently, a total of 16 strobilurin compounds are available worldwide (Stanley Alliance Info-Tech, 2011). In 1999, sales of strobilurins to⇑ Corresponding author. Tel.: +44 2476 575145. E-mail address: [email protected] (C.C. Howell). 0045-6535/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.chemosphere.2013.09.048

talled US$620 million worldwide (Bartlett et al., 2002) and this had increased to US$1.636 billion by 2007 (Stanley Alliance Info-Tech, 2011). The chemical structures of the strobilurin fungicides are based on those of natural products produced by wood-degrading basidiomycete fungi, such as Oudemansiella mucida and Strobilurus tenacellus. The compounds act by disrupting ATP production within the fungal mitochondria and their activities can be either fungicidal or fungistatic (Bartlett et al., 2002). Azoxystrobin is considered relatively immobile in soil, with a Koc of 300–1690 (British Crop Protection Council, 2009), but increased use of the compound has been associated with its detection in surface freshwater. Azoxystrobin was detected in 12 of 18 German streams, with a concentration range of between 0.05 and 29.7 lg L1 (Berenzen et al., 2005), while in the US, azoxystrobin was the most widely detected fungicide in streams, occurring in 46% of 103 samples taken from 29 rivers, with a mean of

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0.16 lg L1 detected (Battaglin et al., 2011). Furthermore, partial degradation of azoxystrobin can produce a metabolite R234886 (also known as azoxystrobin acid) which is much more water soluble and prone to leaching through soil than the parent compound (Ghosh and Singh, 2009), and which has also been identified as a contaminant in surface freshwater (Kern et al., 2009). Jørgensen et al. (2012) observed leaching of metabolite R234886 through loamy, but not sandy, soils at 4 different agricultural sites in Denmark at concentrations up to 2.1 lg L1. Current understanding of the transformation processes involved in strobilurin degradation in soils is limited. Persistence of strobilurins in soil varies widely, with time to 50% degradation (DT50) ranging from days to months (British Crop Protection Council, 2009), and in the case of azoxystrobin, DT50 in soil has been recorded as between 1 to 15 weeks (Joseph, 1999; Bending et al., 2006, 2007; Adetutu et al., 2008; Ghosh and Singh, 2009). An EFSA (2010) report suggested that azoxystrobin DT50 values in aerobic soils ranged from 8 to 40 weeks depending on the soil microbial and chemical properties (EFSA, 2010; Rodrigues et al., 2013). Microbial degradation of most strobilurins, including azoxystrobin, is known to involve the hydrolysis of the carboxyl ester bonds of the parent compound (Katagi, 2006), and a subtilisin- like carboxypeptidase group of enzymes may play an essential role in this process (Clinton et al., 2011). In contrast, pyraclostrobin is degraded by demethoxylation reactions (Katagi, 2006). However, few strobilurin degrading microbes have been described to date. Clinton et al. (2011) isolated a range of Bacillus spp., Arthrobacter and Stentrophomonas strains from soil which used trifloxystrobin as a C source, but isolation of azoxystrobin degrading bacteria was not successful. Additionally a Klebsiella strain isolated from soil was shown to degrade pyraclostrobin in addition to the triazole fungicide epoxiconazole (Lopes et al., 2010). The chemical structures of azoxystrobin, trifloxystrobin, and pyraclostrobin can be seen in Fig. 1. The current study investigated the microbial populations involved in azoxystrobin degradation in soil. Azoxystrobin-degrading bacterial strains were isolated, characterised and their nutritional requirements determined. Further degradation assays were used to determine whether the isolated strains were able to degrade other strobilurin compounds (Fig. 1). Culture independent methods were then used to investigate whether azoxystrobin degradation in soil could be attributed to increased prevalence of the isolated strains.

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2. Materials and methods 2.1. Soil collection and preparation Soil was collected from Hunts Mill Field, School of Life Sciences, Wellesbourne Campus, UK in January 2009. The soil is a sandy loam of the Wick series with a composition of 73% sand, 12% silt, and 14% clay (Bending et al., 2007). The field had been managed as set-aside for over a decade and thus had received no pesticide applications. Soil was collected from the top 20 cm to comply with OECD guidelines for soil sampling in agricultural soils (OECD, 2011). Prior to azoxystrobin application, the soil was re-wetted to a matric potential of 33 kPa (Bending et al., 2006). This equated to a soil moisture content of 13.5%. 2.2. Soil enrichment of azoxystrobin degraders Azoxystrobin (Greyhound Chromatography, Birkenhead, UK) dissolved in acetone at a solvent:soil ratio of 1:20 (Northcott and Jones, 2000) was added to the soil at a concentration of 25 mg kg1 soil using a stainless steel spoon as described by Doick et al. (2003). Control soil received deionised water only. 120 g portions of control and azoxystrobin amended soils were placed into sterile 250 mL glass Duran bottles, wrapped in aluminium foil and stored at 15 °C in the dark. There were 4 replicates for each treatment. Degradation of azoxystrobin was monitored every 2 weeks for 4 months by HPLC. After 4 months, approximately 40% dissipation of azoxystrobin had occurred, and a further 25 mg kg1 dose of azoxystrobin was applied to both the originally amended and a control soil, using the method described above. Samples were stored as previously and degradation monitored every week for 4 weeks by HPLC. 2.3. Azoxystrobin extraction and analysis Azoxystrobin was extracted by adding 10 g of soil to 50 mL centrifuge tubes and mixing it with 20 mL of HPLC-grade acetonitrile (Fisher Scientific, UK). The tubes were shaken for 1 h, left for 30 min to settle, and centrifuged at 4000 rpm for 2 min. 2 mL of the supernatant was decanted into a 2 mL screw-top glass HPLC vial (Chromacol Ltd., UK). Samples were analysed using an Agilent 1100 series unit with a diode array detector (DAD) and LiChrospherÒ 100 RP-18e (5 lm) HPLC column (Column length:

Fig. 1. The chemical structures of (a) Azoxystrobin (b) Kresoxim-Methyl (c) Trifloxystrobin and (d) Pyraclostrobin.

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125 mm, Pore size: 100 Å, Internal diameter: 4.00 mm) (Agilent, UK). A liquid phase of 75% HPLC-grade acetonitrile (Fisher Scientific, UK) and 25% distilled water was used (Flow rate: 1.30 mL min1). 25 lL of each sample was injected and the azoxystrobin concentration determined by monitoring the absorbance at 230 nm. The run time per sample was 4 min with an azoxystrobin retention time of approximately 3 min. The limit of detection (LOD) of the analytical method was 0.02 mg L1. Azoxystrobin recovery from control samples varied from 94.2% to 88.6% of the applied concentration.

2.4. Liquid enrichments of azoxystrobin-degrading cultures Following the cessation of degradation in the second application of azoxystrobin, once and twice amended soils were used to set up liquid enrichment cultures to select azoxystrobin degraders. 15 g Of each soil sample was added to Duran bottles containing 25 mL of sterile minimal basal salts (MBS) medium (Skerman, 1967), which included 0.02 g of MnSO44H2O and 0.005 g of filter sterilised FeSO4 as trace elements (Roberts et al., 1998). Azoxystrobin was added as the sole carbon source to a concentration of 25 mg L1. The Duran bottles were wrapped in aluminium foil and shaken at 150 rpm at a temperature of 15 °C. Degradation of azoxystrobin was monitored by HPLC analysis, as described above, on a weekly basis for 4 weeks. After 4 weeks, 5 mL of each culture was re-enriched with 20 mL of MBS containing 25 mg l1 azoxystrobin and samples monitored as previously. The process was repeated after a further 4 weeks. Control samples containing only MBS and azoxystrobin solution were also prepared to monitor any non-biological degradation of the compound.

2.5. Culture-independent analyses of bacterial community structure changes during enrichment of soil and liquid cultures with azoxystrobin DNA was extracted from soil and liquid cultures 4 weeks postenrichment using the FastDNAÒ Spin Kit (Qbiogene, UK), and amplified using PCR reactions containing MegaMix (Microzone Ltd., UK) following the manufacturer’s guidelines, and the 16S rRNA primer pair 63f-NED/1087r-VIC (5 lM for each primer) (Hauben et al., 1997; Marchesi et al., 1998). PCR products were used for T-RFLP analysis as described by Hunter et al. (2010) using the restriction enzymes HhaI and MspI. All samples were analysed using an Applied Biosystems 3130XL Genetic Analyzer (Applied Biosystems, Warrington, UK). Data was analysed using GeneMarkerÒ software (SoftgeneticsÒ, USA). TRF sizes were determined by reference to LIZ-1200 standards, and the default software settings. Only peaks with an intensity value of 50 or over were selected and used for further analysis (Hackl et al., 2004). A clone library was produced from azoxystrobin-amended soil samples 1-month following the first application as described by Hunter et al. (2010). Clone sequences were determined at the Genome Centre at the University of Warwick, School of Life Sciences, Wellesbourne Campus. Sequences were analysed using the SeqMan programme (DNASTAR Inc., USA). Sequence homologies were identified using the nucleotide BLAST (NCBI, url: http://blast. ncbi.nlm.nih.gov/Blast.cgi?PROGRAM=blastn&BLAST_PROGRAMS=megaBlast&PAGE_TYPE=BlastSearch&SHOW_DEFAULTS= on&LINK_LOC=blasthome) database. A total of 90 16S rRNA sequences were obtained by cloning. The EditSeq programme (DNASTAR Inc.) was used to determine the position of HhaI and MspI restriction sites in each sequence.

2.6. Isolation of azoxystrobin degrading bacteria After the third liquid enrichment, 5 mL of each culture was serially diluted with sterile distilled water (101 to 107), and degradation cultures were set up as described in Section 3.4. Samples were analysed by HPLC every 4 d for 16 d. 50 lL aliquots of the highest dilution cultures exhibiting degradation were sub-cultured onto minimal basal salts agar containing azoxystrobin (MBSA) (Roberts et al., 1998; Rosenzweig et al., 2008) (azoxystrobin concentration: 10 mg L1). Plates were stored at 15 °C in the dark, and monitored for growth on a daily basis. After 10 d, 60 single colonies were selected from across plates for further analysis. Single colony degradation assays were set up in CellstarÒ sterile plastic 6-well TC-plates (Greiner Bio-One, Stonehouse, UK). Each well contained 7 mL of MBS and azoxystrobin solution (25 mg L1) and was inoculated with a loop of cells taken from well separated bacterial colonies growing on MBSA agar. Five un-inoculated controls were also set up. Samples for HPLC analysis were taken every 4 d for 16 d. 1 mL of culture was taken at each time point, and azoxystrobin concentration measured as described in Section 3.4. After 16 d, 50 lL aliquots were taken from the 16 cultures with the lowest percentages of azoxystrobin (40 to 80%) recovery. These were re-inoculated onto MBSA + azoxystrobin plates for 10 d, and used for a second round of degradation assays using the same methods as described previously. The 16 degrader cultures were then spread on MBSA agar plates to check for purity. 2.7. Colony sequencing and identification Two strains were chosen for 16S rRNA gene sequencing and identification. These had degraded 2–3 times more of the azoxystrobin than any other single colony culture. Ten single colonies from each plate were analysed. Single colonies were transferred to sterile Eppendorf tubes containing 100 lL of sterile distilled water. The colonies were re-suspended and 5 lL of the suspension was used in a 50 lL PCR reaction containing 43 lL of MegaMix (Microzone Ltd., UK) and 1 lL of each primer (63f/1087r, 5 lM). The PCR and sequencing reaction conditions used, along with the analytical methods, were the same as described in Section 3.4. The two strains obtained, designated CCH1 and CCH2, were maintained on MBSA for 10 d at 15 °C in the dark as described in Section 3.6, before being used in characterisation assays. 2.8. The effects of carbon (C) and nitrogen (N) availability on azoxystrobin degradation by pure cultures The capacity of CCH1 and CCH2 to degrade azoxystrobin as a sole C and N source was determined. Four variations of the MBS liquid media were set up. These were: (1) azoxystrobin as the sole C and N source (–C–N); (2) azoxystrobin as the sole C source + 6 mM (NH4)2SO4 as a N source (–C + N); (3) azoxystrobin as the sole N source but with 120 mM acetic acid (C2H4O2) as an alternative C source (+C–N); (4) Both the alternative C and N sources present (+C + N). These assays were set up in 6-well TC-plates with 3 replicates per treatment. Each culture was grown in MBS containing 25 mg L1 of azoxystrobin, and cells at an exponential growth stage were removed and inoculated into the 4 media to give a concentration of 106 cells mL1. Un-inoculated control samples were also set up, and azoxystrobin degradation measured every 4 d for 4 weeks, as described in Section 3.4. In order to determine whether there was bacterial proliferation during azoxystrobin degradation by strains CCH1 and CCH2, approximate cell numbers were calculated for each time point based on optical density (OD) measurements taken at a wavelength of 600 nm using a Unicam 5625 UV/VIS spectrometer (Unicam, UK).

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2.9. The degradation of three alternative strobilurin fungicides by azoxystrobin-degrading cultures The capacity of CCH1 and CCH2 to degrade the strobilurin fungicides trifloxystrobin, pyraclostrobin, and kresoxim-methyl was determined (Greyhound Chromatography, UK). Control uninoculated samples were also prepared. Degradation was determined using the procedures outlined in Section 3.4, using MBS containing 25 mg L1 of each strobilurin as the sole C source. The HPLC conditions used were the same as those described in Section 3.3. The retention times were approximately 3 min for azoxystrobin, 4.5 min for pyraclostrobin, 5 min for trifloxystrobin, and 4 min for kresoxim-methyl. Recovery of control samples varied between 100.2% and 87.4% of the applied concentration. 2.10. Mineralisation of azoxystrobin by degrader cultures The method of Reid et al. (2001) was used to determine the extent to which CCH1 and CCH2 mineralised the cyanophenyl ring of azoxystrobin. Three degrader treatment cultures (1. Cupriavidus spp.; 2. Rhodanobacter spp.; 3. Cupriavidus spp. + Rhodanobacter spp.; along with an un-inoculated control) containing [14C]Cyanophenyl-labelled azoxystrobin (>98% purity) were set up using procedures outlined in 3.4. The concentration of evolved 14CO2 was determined using a Canberra Packard Tri-Carb 2250CA liquid scintillation analyser (Canberra Packard, UK). 2.11. Statistical analyses Since azoxystrobin degradation did not always extend beyond 50%, the time taken for 25% of the applied azoxystrobin to degrade (DT25) was calculated (Bending et al., 2006) using the vinterpolate function in the GenStat Version 12 statistics programme (VSN International, UK). Two-way ANOVAs and least significant difference (LSD) analysis was used to determine significant differences between treatments at each time point. NMDS profiles were produced using the Primer 6 programme (Primer-E, UK). ANOSIM analysis was used to determine significant bacterial community structure changes, and SIMPER analysis detailed the individual TRFs responsible for the most community variation between treatments. 3. Results 3.1. The degradation of azoxystrobin in once- and twice-amended soils Azoxystrobin degradation was significantly higher in the twiceamended soils compared to the once-amended treatment (p = 0.019). The once- and twice-amended treatments had degraded 14% and 30% of the applied compound, respectively, after 1 week, and 32 and 43% respectively after 4 weeks (Fig. 2). The DT25 values for each treatment were significantly different (p 6 0.01), at 12 and 5.5 d for the once- and twice-amended soils, respectively. 3.2. Sequential enrichments of azoxystrobin-degrading cultures: HPLC analysis of azoxystrobin degradation There was a significant difference in degradation between the once- and twice amended treatments over the course of the three liquid enrichments (p = 0.028), with more rapid degradation in enrichments derived from the twice amended soil (Fig. 3). Differences were particularly noticeable in enrichment 1, in which DT25 values for the once- and twice-amended treatments were

Fig. 2. Percentage of extractable azoxystrobin recovered using HPLC analysis. j Once-amended h Twice-amended. Each data point represents the mean of 3 experimental replicates.

11 and 5 d, respectively (significantly different, p = 0.039). In all enrichments, degradation initially proceeded rapidly, but had ceased by 3 weeks, when 25–60% azoxystrobin remained. In the control samples, between 87.4% and 90.0% of the initially applied azoxystrobin was recovered after 4 weeks. 3.3. T-RFLP analysis of bacterial community structure changes There was no significant difference (p = 0.407) in the number of TRFs recorded in the once-amended treatments compared with the twice-amended ones through the soil and liquid enrichment process, although the number of TRFs declined from 44 in soil to 15 in enrichment 3. ANOSIM analyses showed significant community structure differences between the once- and twice-amended soil and liquid culture treatments (p 6 0.01), and between each of the soil and liquid enrichments (p 6 0.01). This was supported by NMDS analysis (Fig. 4) which showed a grouping of sample points based both on the treatment (once- or twice-amended) and the enrichment number. SIMPER analysis showed that the presence of the TRFs at 470 bp (HhaI) and 501 bp (MspI) were responsible for a high level of the variation (12% each) between the onceand twice-amended treatments across the soil and liquid enrichments. Analysis of the 16S rRNA sequences obtained by cloning revealed that three different sequences with 100% homology to Rhodococcus spp. (Accession numbers: AY027586.1, AB183437.1, GU815137.1) had predicted TRFs of 470 bp using HhaI. A sequence with 95% homology to Methylibium sp (Accession number: AB1621105.1) produced a TRF of 471 bp with HhaI. Lastly, a sequence with a 100% homology to Haliea sp. (Accession number: JN177653.1) produced a TRF of 501 bp with MspI. 3.4. Isolation of azoxystrobin degrading bacteria Of the 60 colonies tested initially for azoxystrobin degradation, 35 degraded <10% of the compound, 9 degraded from 10% to 25%, whilst 16 degraded between 30% and 65% of the compound (Data not shown). There was no degradation in the uninoculated controls. Of the 16 single cultures selected for further testing, 7 showed degradation of between 30% and 35% of the applied azoxystrobin, and 7 degraded 50–60% (Data not shown). The remaining two single colony cultures (strains CCH1 and CCH2) degraded 85 and 91% of the azoxystrobin, respectively.

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Fig. 3. Percentage of extractable azoxystrobin recovered from the once- and twice-amended samples over a series of 3  4 week liquid culture enrichments X Control j Once-enriched h Twice-enriched. The black arrows denote the points at which the cultures were enriched. Each data point represents the mean of 3 experimental replicates.

3.5. Colony sequencing and identification The colonies of strain CCH1 were relatively large (5 mm), circular, yellow, and had a ridged surface. The colonies from strain CCH2 were smaller, and white/grey with a smooth, raised surface. Neither colony type produced any kind of halo in the agar. BLAST searches of the 16S rRNA gene sequence showed that strain CCH1 had a 98% sequence homology to an uncultured Rhodanobacter sp. (Accession number: JF968486.1), whilst strain CCH2 had a 99% sequence similarity to Cupriavidus sp. B-8(2011) (Accession number: JN128831.1). The sequences were submitted to the EMBL database under the accession numbers HE598561.1 and HE576771.1 for strains CCH1 and CCH2, respectively. Further 16S rRNA gene sequence analysis showed that strain CCH1 would have produced TRFs of 175 and 104 bp using HhaI and MspI, respectively. In contrast, strain CCH2 would have produced TRFs of 154 bp using HhaI and 377 bp using MspI. 3.6. The effects of carbon and nitrogen availability on azoxystrobin degradation There was no difference between CCH1 and CCH2 in the rate or extent of azoxystrobin degradation across the four treatments (–C–

100 90 80 70 60 50 40

LSD = 17.48

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Sampling Time (Days) Applied Azoxystrobin Recovered (% of uninoculated control)

Fig. 4. NMDS plot of the bacterial community structures in the once- and twiceamended treatments during 4 different enrichment series. Filled symbols denote the once-amended treatments and open symbols represent the twice-amended treatments. jh Soil Enrichment; N 4 Liquid Enrichment 1; d s Liquid Enrichment 2; . O Liquid Enrichment 3. Each data point represents an experimental replicate.

Applied Azoxystrobin Recovered (% of uninoculated control)

N, –C + N, +C–N, and +C + N) (p = 0.907). However, carbon and nitrogen availability did affect azoxystrobin degradation by the strains (p 6 0.01) (Fig. 5a and b). There was limited degradation of azoxystrobin when either no additional C and N, or C without additional N was supplied. Azoxystrobin loss was greatest in cultures where it was present as the sole carbon source and an additional nitrogen source was supplied, with 88.5% and 85.5% of the azoxystrobin degraded after 16 d for strains CCH1 and CCH2, respectively. The presence of an alternative carbon source significantly reduced azoxystrobin degradation, with 39.0% and 41.0% of the azoxystrobin degraded for strains CCH1 and CCH2 after 16 d, respectively. Azoxystrobin recovery from the uninoculated

100 90 80 70 60 50 40 LSD = 17.00

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Sampling Time (Days) Fig. 5. The percentage of azoxystrobin recovered from (a) Strain CCH1 and (b) Strain CCH2 liquid cultures under different carbon and nitrogen source treatments. j – C – N h + C – N N – C + N 4 + C + N. Each data point represents the mean of 3 experimental replicates.

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control samples varied from 83.3% to 91.7% of the applied compound after 16 d (data not shown). There was a marked increase in bacterial cell growth for the – C + N and +C + N treatments for both strains over the course of the study, which is indicative of bacterial growth during degradation (Fig. 6a and b). For strain CCH1, the highest growth rate was recorded between days 0 and 4 for the –C + N treatment where cell numbers increased from 1  106 to 5  107 cells mL1. In contrast, the greatest increase in cell number for the +C + N treatment was recorded between days 4 and 8 (from 5  106 to 2  107 cells mL1). Bacterial growth was significantly lower in the +C–N and –C–N treatments. The growth rate of strain CCH2 was highest between days 0 and 4 for both the –C + N and +C + N treatments (1  106 to 8.5  107 cells mL1). However, at the end of the experimental period the cell number was higher in the –C + N treatment (5.5  109 cells mL1) compared to the +C + N treatment (6.0  108 cells mL1). For both strains, lower growth was recorded in the –C–N and +C–N treatments which compared well with the lower levels of degradation in these treatments. 3.7. The degradation of three alternative strobilurin fungicides by azoxystrobin-degrading cultures There was no difference between CCH1 and CCH2 in the rate or extent of degradation of the different strobilurins (p = 0.235) (Fig. 7a and b). However, there were differences in the rate of degradation of the different compounds (p 6 0.01). Trifloxystrobin was degraded more slowly than the other compounds. For strain CCH1, average DT25 for azoxystrobin, pyraclostrobin, trifloxystrobin, and kresoxim-methyl were 2.0, 2.3, 4.2, and 2.6 d respectively. For strain CCH2, average DT25 values were 2.2, 2.8, 4.2, and 2.9 d for

Fig. 7. The percentage of 4 different strobilurin fungicides recovered from (a) strain CCH1 and (b) strain CCH2 liquid cultures. j Azoxystrobin h Pyraclostrobin N Trifloxystrobin 4 Kresoxim-Methyl. Each data point represents the average of 3 experimental replicates.

the azoxystrobin, pyraclostrobin, trifloxystrobin, and kresoximmethyl, respectively. Compound recovery rates from control samples ranged from 87% to 100% of the applied pesticide for azoxystrobin, 90–98% for pyraclostrobin, 87–94% for trifloxystrobin, and 85–99% for kresoxim-methyl after 16 d (data not shown). 3.8. Mineralisation of azoxystrobin by degrader cultures There was a very low level of azoxystrobin mineralisation by CCH1 and CCH2 when grown individually and together (Data not shown). Strain CCH1 mineralised 0.20% of the azoxystrobin after 14 d, compared with 0.83%, and 0.13% for strain CCH1, and the mixed culture, respectively. As the purity of the azoxystrobin used was only >98%, it could not be confirmed that the 14CO2 evolved resulted from the mineralisation of azoxystrobin. 4. Discussion

Fig. 6. Bacterial growth curves for (a) Strain CCH1 and (b) Strain CCH2 liquid cultures under different carbon and nitrogen source treatments. j – C – N h + C – N N – C + N 4 + C + N. Each data point represents the mean of 3 experimental replicates.

The two catabolic bacterial strains isolated from liquid cultures showed 98 and 99% sequence homologies to members of the genera Rhodanobacter and Cupriavidus sp., respectively. While bacterial isolates capable of trifloxystrobin and pyraclostrobin degradation have previously been isolated from soil (Lopes et al., 2010; Clinton et al., 2011), azoxystrobin degrading strains have not been characterised before. Furthermore, unlike the other described strobilurin degraders, those isolated in the current work had wide specificity and were also able to degrade pyraclostrobin, trifloxystrobin, and kresoxim-methyl. The genus Rhodanobacter was first proposed by Nalin et al. (1999) who identified a novel lindane-degrading bacterium,

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Rhodanobacter lindaniclasticus. In addition to R. lindaniclasticus, R. xiangquanii is also capable of degrading xenobiotic compounds such as the herbicide, anilofos (Zhang et al., 2011). Cupriavidus is a genus of b-Proteobacteria and encompasses some of the former members of the genus Wautersia. Members of this genus including C. taiwanensis (Chen et al., 2008), C. pinatubonensis (Mayer et al., 2010), C. pampae (Cuadrado et al., 2010), and C. necator (Streber et al., 1987) have been associated with the degradation of a wide range of xenobiotic compounds. Pérez-Pantoja et al. (2008) described how C. necator JMP134 was capable of using 60 out of 140 aromatic compounds tested (including 2,4-D, halobenzoates, chlorophenols and nitrophenols) as a sole carbon source. The bacterium was capable of performing the majority of the common ring cleavage metabolic pathways for aromatic compounds. The degradation of azoxystrobin by strains CCH1 and CCH2 appears to have been growth linked. This was illustrated by increases in cell number when the compound was supplied as a sole C source. The absence of an additional nitrogen source dramatically reduced the growth of strains CCH1 and CCH2 and the subsequent degradation of azoxystrobin. This suggests that the isolates were able to use the compound as a carbon source only. Similar results have also been observed in a microbial consortium degrading the herbicide alachlor (Chirnside et al., 2007), and also in para-nitrophenol (PNP) degrading cultures (Rehman et al., 2007; Zhang et al., 2009). These studies suggest that, as with azoxystrobin, the microbial communities analysed were better adapted for using xenobiotic amendments as sources of carbon rather than nitrogen. Azoxystrobin degradation was significantly decreased (50%) when an alternative carbon source (acetic acid) was provided. This suggests that strains CCH1 and CCH2 did not preferentially degrade either carbon source. This could be more indicative of azoxystrobin degradation in soil environments, where microbial communities are exposed to a constantly changing pool consisting of a variety of carbon and other nutrient sources (Rüegg et al., 2007). There was no significant mineralisation of azoxystrobin to CO2 by either of the two isolates. This may indicate that the isolates were able to utilise the carbon present in the azoxystrobin side chains, but were unable to break down the ring structure where the 14C label was attached. These observations support previous studies into the mineralisation of the compound in soil (Adetutu et al., 2008) and water (Singh et al., 2010). To date, the only strobilurin compounds for which degraders have been isolated are trifloxystrobin (Clinton et al., 2011) and pyraclostrobin (Lopes et al., 2010). The trifloxystrobin isolates isolated by Clinton et al. (2011) were unable to degrade azoxystrobin, but degradation could be attributed to a non-specific subtilase like protease which showed low rates of hydrolysis indicative of cometabolism. Using strains from culture collections with no history of exposure to strobilurins, Clinton et al. (2011) showed that the mechanisms associated with trifloxystrobin were widespread among Bacillus spp., Arthrobacter and Stentrophomonas strains, indicating a widespread promiscuous catabolic enzyme in these genera of bacteria. The azoxystrobin-degrading isolates obtained in the current study were able to degrade three other strobilurin fungicides, including trifloxystrobin and pyraclostrobin. Furthermore, degradation in the current study was associated with growth-linked catabolism. Clearly specificity of the catabolic enzymes associated with strains isolated in the current study was lower than that of strains isolated in the study by Clinton et al. (2011). Similarly, previous research using herbicides also showed that some bacterial isolates were able to degrade several structurally-related compounds (Cullington and Walker, 1999), whereas others were not (Batisson et al., 2009; Hussain et al., 2009). In this study, 16S rRNA T-RFLP analysis showed that the bacterial community structure was significantly different between the once- and twice-amended treatments in both soil and liquid

enrichments. The TRFs at 470 and 501 bp were responsible for 12% of the community structure variation each and were more prevalent in the twice-amended treatment. Sequence analysis of strains CCH1 and CCH2 showed that they would have produced TRFs of 175 and 154 bp using the restriction enzyme HhaI, and 104 and 377 bp using the enzyme MspI. These TRFs occurred in the 16S rRNA TRFLP profiles but were their abundance was not affected by the sequential enrichment process. Lozada et al. (2004) recorded that c-Proteobacteria were most commonly isolated from activated sludge communities adapted to degrade nonylphenol ethoxylate (NPnEO). In contrast, culture-independent methods showed that the community was in fact dominated by b-Proteobacteria with the c-Proteobacteria only representing a small proportion of the community. Other examples of this have been seen in degradation cultures of TNT (Travis et al., 2008), the nematicide, fenamiphos (Chanika et al., 2011) and isotproturon (Bending et al., 2003; Bending and Rodríguez-Cruz, 2007). Such differences have previously been attributed to the transfer of genes conferring compound degradation from one strain/species to another, during the enrichment process (Newby et al., 2000). Whilst the use of culture-independent methods indicated that other, noncultured organisms may also have been involved in azoxystrobin degradation, application of further techniques such as stable isotope probing (SIP) would be required to link these processes to specific bacteria (Cupples and Sims, 2007). In the soil enrichments, there was an initially high degradation rate within the first month after which degradation slowed before stopping after three months. This indicated that, over time, azoxystrobin becoming less bioavailable. A similar pattern of degradation was observed for the nematicide ethoprophos (Karpouzas and Walker, 2000). It has been suggested that bioavailability and biodegradation are intrinsically linked with abiotic factors such as compound sorption to soils, particularly organic matter (Semple et al., 2004). However, there have been contrasting reports as to the role that sorption per se plays in degradation. Laor et al. (1999) suggested that the chemical characteristics of a compound may be a stronger determining factor than the sorption itself. This theory has been supported by the results of research using both hydrophobic (Ahmad et al., 2004) and hydrophilic (Schnürer et al., 2006) compounds However, in some cases, no correlation has been found between pesticide sorption and biodegradation (Kah et al., 2007). There was a decrease in azoxystrobin degradation rates over time even in the second and third liquid enrichment cultures in which the size of the soil fraction was negligible compared to liquid enrichment 1. This could suggest that azoxystrobin bioavailability was reducing as a result of sorption of the compound onto the bacterial biomass within the culture. Similarly, trifloxystrobin isolates were not able to degrade more than 50% of the compound when applied as a sole C source in liquid culture (Clinton et al., 2011).

5. Conclusions Two azoxystrobin-degrading bacterial strains showing sequence homologies to Cupriavidus sp. and Rhodanobacter sp. were successfully isolated from a soil enrichment series. Both isolates were capable of partially degrading azoxystrobin and other strobilurin fungicides when an additional nitrogen source was present. Fungicide degradation was significantly reduced by the presence of an alternative carbon source. However, culture independent analysis indicated that non-culturable strains may have been more important agents of degradation in situ. While isolated catabolic strains can be valuable for providing information about degradation pathways and as agents

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