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Applied and Environmental Microbiology, February 2003, p. 827-834, Vol. 69, No. 2
0099-2240/03/$08.00+0     DOI: 10.1128/AEM.69.2.827-834.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.

In-Field Spatial Variability in the Degradation of the Phenyl-Urea Herbicide Isoproturon Is the Result of Interactions between Degradative Sphingomonas spp. and Soil pH

Gary D. Bending,1* Suzanne D. Lincoln,1 Sebastian R. Sørensen,2 J. Alun W. Morgan,1 Jens Aamand,2 and Allan Walker1

Horticulture Research International, Wellesbourne, Warwick CV35 9EF, United Kingdom,1 Department of Geochemistry, Geological Survey of Denmark and Greenland, DK-1350 Copenhagen, Denmark2

Received 8 July 2002/ Accepted 25 October 2002


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ABSTRACT
 
Substantial spatial variability in the degradation rate of the phenyl-urea herbicide isoproturon (IPU) [3-(4-isopropylphenyl)-1,1-dimethylurea] has been shown to occur within agricultural fields, with implications for the longevity of the compound in the soil, and its movement to ground- and surface water. The microbial mechanisms underlying such spatial variability in degradation rate were investigated at Deep Slade field in Warwickshire, United Kingdom. Most-probable-number analysis showed that rapid degradation of IPU was associated with proliferation of IPU-degrading organisms. Slow degradation of IPU was linked to either a delay in the proliferation of IPU-degrading organisms or apparent cometabolic degradation. Using enrichment techniques, an IPU-degrading bacterial culture (designated strain F35) was isolated from fast-degrading soil, and partial 16S rRNA sequencing placed it within the Sphingomonas group. Denaturing gradient gel electrophoresis (DGGE) of PCR-amplified bacterial community 16S rRNA revealed two bands that increased in intensity in soil during growth-linked metabolism of IPU, and sequencing of the excised bands showed high sequence homology to the Sphingomonas group. However, while F35 was not closely related to either DGGE band, one of the DGGE bands showed 100% partial 16S rRNA sequence homology to an IPU-degrading Sphingomonas sp. (strain SRS2) isolated from Deep Slade field in an earlier study. Experiments with strains SRS2 and F35 in soil and liquid culture showed that the isolates had a narrow pH optimum (7 to 7.5) for metabolism of IPU. The pH requirements of IPU-degrading strains of Sphingomonas spp. could largely account for the spatial variation of IPU degradation rates across the field.


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INTRODUCTION
 
Concern about the environmental impact of pesticides most frequently arises from their ability to leach from soil and contaminate water resources. The phenyl-urea herbicides are of particular significance in this respect, since several members of the group, including isoproturon (IPU) [3-(4-isopropylphenyl)-1,1-dimethylurea] and diuron [3-(3,4-dichlorophenyl)-1,1-dimethylurea] are degraded slowly in soil and are susceptible to leaching. As a result, IPU and diuron are frequently detected as contaminants of agricultural catchments in Europe (23).

Studies of the fate of IPU in agricultural fields on contrasting soil types have revealed considerable spatial variability in degradation rates across fields (2, 26, 27). At two such sites in the United Kingdom, Deep Slade field in Warwickshire and Brimstone farm in Oxfordshire, IPU half-life in soil was found to vary between 6 and 30 days, with degradation rate linked to soil pH. Further studies by Bending et al. (3) demonstrated that soil pH controlled the ease of induction of growth-linked metabolism, with slow degradation rates at lower soil pH linked to apparent cometabolic degradation of the compound. At the Deep Slade site, various bacteria capable of degrading IPU have been isolated (8, 19, 21). However, the relative importance of each in the degradation of IPU in situ is unclear.

IPU-degrading isolates from Deep Slade field include an Arthrobacter sp., which degrades phenyl-ureas to their aniline derivatives by hydrolysis of the urea side chain at the carbonyl group (8, 24, 25), and a Sphingomonas sp. (strain SRS2), which degrades IPU by sequential N-demethylation of the urea side chain followed by mineralization of the phenyl-urea structure (21). However, Sphingomonas sp. strain SRS2 requires the presence of other bacteria in coculture, or amino acid supplements, in order to extensively degrade IPU in liquid culture and agricultural soil (22).

The IPU-degrading bacteria obtained in these studies were isolated following sequential enrichment procedures, which are known to be susceptible to shifts in the nature of organisms carrying degradative genes (10, 11, 18), and it is unclear which, if any of the IPU-degrading isolates contributes to IPU degradation in situ. Further, it is not known how the spatial variability in IPU degradation at the field scale, which appears to be controlled by pH, relates to the distribution and activities of IPU-degrading organisms.

In the present study we investigated the characteristics and dynamics of IPU-degrading bacteria in soil from Deep Slade field using most-probable-number (MPN) techniques, enrichment-isolation, and denaturing gradient gel electrophoresis (DGGE) of bacterial 16S rRNA genes. Further, we investigated effects of soil pH on the metabolism of IPU by Sphingomonas sp. strain SRS2, for which we show evidence of proliferation in Deep Slade soil during IPU degradation.


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MATERIALS AND METHODS
 
Soil.
Soil was collected from Deep Slade field at HRI-Wellesbourne, Warwickshire, United Kingdom. The soil is a sandy loam of the Wick series, with (wt/vol) 2.5% organic matter, 14% clay, 74% sand, and 12% silt (28). The field had received regular applications of IPU for 20 years prior to the study and annually for the previous 5 years. Soil was collected from two parallel transects, separated by 50 m, across areas of the field known to show contrasting degradation characteristics. Soil was collected from a depth of 0 to 15 cm at 20-m intervals, with five samples collected from each transect (3). Transect 1 was located in an area of the field previously identified as giving rapid biodegradation of IPU, and the pH values of the samples ranged from 7.1 to 7.5. Transect 2 was taken from an area known to give slow degradation of IPU, and the pH of samples was 6.3 to 6.5. Soil samples were sieved (particle size, <3 mm) and allowed to air dry to approximately 8% moisture content.

Soil-IPU incubation.
An aqueous suspension of the commercial formulation of IPU (Aventis Crop Science, Lyon, France) was prepared at a concentration of 0.94 mg IPU ml-1. Each portion of soil (fresh weight [fw], 500 g) was spread over a fresh polyethylene sheet, and 8 ml of the IPU suspension was dropped evenly over the soil to give an IPU concentration of 15 mg kg of soil (fw)-1. This is equivalent to concentrations found in the top 1 cm of soil following application of the recommended dose of 2.5 kg ha-1. Further distilled H2O was added as necessary to amend each soil sample to 10% moisture content, which was equivalent to a matric potential of -33 kPa. Each soil sample was mixed thoroughly by hand using a new pair of latex disposable gloves for each sample. The soil was transferred to screw-top polypropylene bottles. For each soil sample, parallel bottles were set up in which distilled H2O was added to give 10% moisture content but which received no application of IPU. Samples were incubated at 15°C in the dark. The moisture content of each bottle was maintained at 10% through the experiment.

IPU extraction and analysis.
Immediately after application of IPU, and at regular intervals thereafter, samples were removed from each bottle for analysis of residual IPU. Soil samples (15 g [fw]) were extracted by shaking with 90:10 (vol/vol) acetonitrile-H2O (25 ml) for 1 h. Once the soil had settled, 250 µl of the supernatant was diluted with 500 µl of acetonitrile, and the IPU concentration was determined by high-performance liquid chromatography (HPLC). The HPLC system (Konitron series 300) was equipped with a Lichrosorb RP18 column (250 by 4.6 mm; Merck, Poole, United Kingdom). IPU was eluted using a mobile phase of acetonitrile-water-orthophosphoric acid of 75:25:0.25, with a flow rate of 1 ml min-1, detected by UV absorbance at 240 nm, and its concentration was determined by reference to an analytical standard (British Greyhound, Birkenhead, United Kingdom).

Prior to IPU application, at the point of approximately 40 and 90% IPU degradation, and 9 months following IPU addition, samples of soil were taken for MPN analysis of IPU-degrading organisms and characterization of the microbial community using DGGE of 16S rRNA genes.

MPN analysis of IPU-degrading organisms.
Soil (25 g [fw]) was suspended in 100 ml of Ringer's solution (Fisher Scientific, Loughborough, United Kingdom) and shaken for 20 min, following which 11 serial fivefold dilutions were prepared in Ringer's solution. A 100-µl aliquot of each dilution was inoculated into four microtiter plate wells containing 100 µl of double-strength mineral salts liquid medium (MSL) (19) plus IPU (40 mg liter-1). Plates were incubated in the dark at 20°C for 4 weeks. IPU concentrations in the inoculated microtiter plate wells were determined by HPLC using the system described above. IPU degradation was counted as positive if the mean amount of IPU remaining plus 2 standard deviations amounted to less than 100%. The number of positive and negative wells at each dilution was used to give an MPN of degraders using Genstat (version 5.3; Lawes Agricultural Trust, Rothamsted Experimental Station, Harpenden, United Kingdom).

DNA extraction and DGGE analysis.
DNA was extracted from 1-g (fw) portions of soil by bead beating in a 0.12 M NaHPO4-Na EDTA buffer (pH 8.0) as described by Cullen and Hirsch (7). DNA was purified using a Geneclean II kit (Bio 101, Nottingham, United Kingdom) and diluted 10-fold in MilliQ H2O prior to use. Partial eubacterial 16S rRNA gene fragments were amplified using primers described by Muyzer et al. (17), at positions 341f and 534r (Escherichia coli numbering), using a Hybaid (Ashford, United Kingdom) Omnigene thermocycler. The 50-µl PCR mixtures contained 1 U of DNA polymerase (Finnzymes, Espoo, Finland), and a 200 µM concentration (each) of dATP, dCTP, dGTP, and dTTP (Abgene, Surrey, United Kingdom) in a 10 mM Tris-HCl buffer (pH 8.8), with 1.5 mM MgCl2, 50 mM KCl, and 0.1% Triton X-100 (Finnzymes). The reaction conditions were 1 cycle of 95°C for 5 min, 35 cycles of 94°C for 1 min, 55°C for 1 min, and 72°C for 2 min, with a final extension of 72°C for 5 min. DGGE gels were set up according to the method of Muyzer et al. (17) using a DCode Universal Mutation System (Bio-Rad, Hemel Hempstead, United Kingdom) with 8% acrylamide, and a denaturant gradient of 20 to 70% (100% denaturant was equivalent to 7 M urea with 40% [vol/vol] formamide). The gels were run at 60 V and 60°C for 18 h. The gels were stained with ethidium bromide (0.5 mg liter-1) and visualized under UV light on an Imago Imaging system (B and L systems, Maarssen, The Netherlands). Bands of interest were cut from the gel and left overnight in 50 µl of MilliQ H2O at 4°C for 18 h. After centrifuging, DNA in the supernatant was amplified as described above. Reamplified bands were run against the original sample to check motility and purity. A mixture comprising of partial 16S rRNA of Shewanella sp., Pseudomonas sp., Alcaligenes sp., Sphingomonas sp., and Desulfovibrio sp. (4) was used as a standard marker for the DGGE analysis.

Cloning and sequencing.
The PCR products were purified using a QIAquick PCR purification kit (Qiagen Ltd., Dorking, United Kingdom) and then cloned into pGEM-T Easy vector (Promega, Southampton, United Kingdom). The DNA was heated at 65°C for 5 min, dialyzed, and electroporated (field strength, 12.5 kV/cm2) into electrocompetent cells of E. coli strain JM109. Colonies containing inserts were selected according to manufacturer's instructions. Plasmid DNA was extracted and purified from selected colonies using a Qiagen Ultra Plasmid kit (Qiagen). Sequencing reactions were performed according to manufacturer's instructions, on a Hybaid PCR multiblock system, using a PRISM BigDye Terminator Cycle Sequence reaction kit (Applied Biosystems, Warrington, United Kingdom) and the SP6 promoter primer (Promega). Products were analyzed on an Applied Biosystems 377 DNA sequencer.

Isolation of IPU-degrading bacteria.
At the point of 90% IPU degradation, 5 g of soil (fw) from each of the fast-degrading microsites was inoculated into 25 ml of MSL containing IPU at 20 mg liter-1 (MSLIPU). The flasks were incubated at 25°C in an orbital shaker at a speed of 100 rpm. Degradation of IPU was monitored by HPLC. At the point of 50% degradation, a 0.5-ml aliquot of the culture was transferred to a fresh flask containing 25 ml of MSLIPU. After three successive enrichment cycles, 10-fold dilutions of the culture were prepared in Ringer's solution, and 100-µl aliquots were spread onto plates of MSLIPU agar. Forty colonies from the plates were inoculated into 0.5-ml aliquots of MSLIPU, and degradation after 14 days was determined. Cultures were found to stop degrading IPU after two successive enrichments in MSLIPU. Addition of peptone or Casamino Acids to the MSL medium was subsequently found to facilitate continued degradation by the culture. The single IPU-degrading culture obtained via enrichment (designated strain F35) was characterized by partial 16S rRNA sequence analysis, using the primers and conditions described above. General characterization of the strain using traditional tests was also carried out.

Sequence analysis.
The partial 16S rRNA sequences were edited and assembled using the DNAstar II sequence analysis package (Lasergene Inc., Madison, Wis.). Sequences were compared to those on the EMBL DNA database using the program FASTA3. Reference partial 16S rRNA sequences were gathered from the EMBL database, and the partial sequences corresponding to the 173-bp fragment were used for phylogenetic analysis. The DGGE, isolate and reference partial 16S rRNA sequences were analyzed using the PHYLIP (version 3.5c) packages SEQBOOT, DNADIST, and NEIGHBOR. The dendrogram was generated using neighbor-joining analysis and the results were viewed using DRAWTREE.

Effect of pH on IPU degradation by Sphingomonas sp. strain SRS2.
Sphingomonas sp. strain SRS2, obtained from Deep Slade field by Sørensen et al. (21), was used to investigate the effect of pH on degradation of IPU in soil and pure culture. Soil was taken from soil sites with pH of 6.5 and 7.5 which were located within Hunts Mill field, which is next to Deep Slade on the farm at HRI Wellesbourne. Hunts Mill is an organic area, which had received no pesticides for at least 5 years. Soil was autoclave sterilized (121°C for 30 min at 0.1 MPa) twice, with a 24-h interval between treatments. Portions of sterilized or nonsterile soil (200 g [fw], sieved to a particle size of <3 mm) were placed into 500-ml polypropylene jars. For sterilized and nonsterile soil at each pH, three replicate jars were inoculated with strain SRS2 or left uninoculated. Jars were inoculated with a suspension of strain SRS2 in exponential growth phase, to give 6 x 105 cells g of soil (fw)-1. The jars were incubated at 15°C in the dark. IPU remaining was determined weekly over a 60-day period.

For the liquid culture experiment 14C-ring-labeled IPU ([phenyl-U-14C]-IPU; 914 MBq mmol-1; 97% radiochemical purity; Amersham Life Science, Little Chalfont, United Kingdom) and unlabeled IPU in acetone were added to sterile 100-ml flasks to give 1 mg of IPU and 40,000 dpm. The acetone was allowed to evaporate before 49 ml of MS medium supplemented with 0.1 g of Casamino Acids liter-1 (21) and adjusted to a pH of 6.5, 7.0, 7.5, or 8.0 using 0.1 M NaOH was added. A suspension of SRS2 in exponential growth phase was inoculated into the flasks to give a density of 107 cells ml-1. IPU mineralization was determined by trapping 14CO2 in 2 ml of 0.5 M NaOH contained in a 5-ml test tube within the flask. Mineralization was followed over a 10-day period, with fresh NaOH solution added at each sampling. The NaOH was mixed with 10 ml of Wallac (Turku, Finland) OptiPhase HiSafe 3 scintillation liquid, and the mixture was analyzed on a Wallac 1409 liquid scintillation counter. At all the tested pHs exponential mineralization of [14C]IPU to 14CO2 was observed and based on log-transformation and estimation of the slope, specific mineralization rates were calculated individually for each sample with a minimum of four data points.

Nucleotide sequence accession numbers.
Nucleotide sequences for clones DS1 and DS2 and culture F35 have been deposited in the EMBL nucleotide sequence database under accession numbers AJ509085, AJ509086, and AJ509087, respectively.


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RESULTS
 
IPU degradation.
Soils from the two transects showed distinct patterns of IPU degradation (Fig. 1). All soils from transect 1, together with a single soil from transect 2 (site E) degraded IPU rapidly. In these samples approximately 25% of applied IPU was degraded after 5 days, following which there was a period of rapid loss, with complete degradation after 10 days. Similarly, approximately 20% of the IPU had been degraded in soils from transect 2 after 5 days. There was a period of rapid degradation in soil from two further sites along transect 2 (sites D and F), although this was delayed relative to that in soils from transect 1, so that complete degradation did not occur until 23 and 40 days following that in soil from transect 1. In soil from two sites along transect 2 (sites B and C), degradation showed first-order kinetics, with no rapid phase of degradation, and approximately 15% IPU remained after 65 days. In all sites, HPLC analysis revealed the temporary appearance of peaks with identical retention times to 3-(4-isopropyl-phenyl)-1-methylurea (MDIPU) and 3-(4-isopropyl-phenyl)-urea (data not shown), indicating that metabolism of IPU occurred by sequential N-demethylation, as outlined by Mudd et al. (16).



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  		FIG. 1. IPU degradation in soils from transects 1 (open symbols) and 2 (closed symbols). Symbols: circles, site B; inverted triangles, site C; squares, site D; diamonds, site E; triangles, site F.

MPN analysis of IPU-degrading organisms.
IPU-metabolizing communities were detected in all soils before the addition of IPU (Fig. 2), with an average of 5 x 103 and 7 x 102 IPU degraders g-1 soil detected in soils from transect 1 and 2, respectively. In soils from transect 1, application of IPU stimulated proliferation of IPU-degrading organisms, with an average of 2.5 x 106 g-1 soil at the point of 90% IPU degradation. After 9 months, numbers of IPU-degrading organisms had declined to an average of 5 x 102 g-1 soil.



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  		FIG. 2. MPN of IPU-degrading organisms prior to application (white bars), at the points of 40 (diagonally striped bars) and 90% (black bars) degradation, and 9 months (horizontally striped bars) after the start of the experiment. (a) Transect 1; (b) transect 2.

In transect 2 there was considerable variability in the dynamics of IPU-degrading organisms. By the point of 40% degradation, there was evidence for proliferation of degrading organisms in 1 site (site E), in which the dynamics of IPU-degrading organisms, and degradation of IPU were similar to sites in transect 1. By the point of 90% degradation, there was evidence for proliferation of IPU-degraders in 2 further sites (sites D and F) that had shown a phase of rapid IPU degradation (Fig. 1). However, in the sites showing first-order IPU degradation kinetics (sites B and C), there was little or no evidence for proliferation of IPU-degrading organisms.

DGGE analysis.
In sites from transect 1, degradation of IPU was associated with changes to 2 DGGE bands (Fig. 3a) in soil from all sites. At the point of 90% degradation, a single band (DS2) sharply increased its intensity in IPU treated, relative to untreated soil. In four of the five sites, a further band (DS1) which was not present in untreated soil, appeared in IPU-treated soil at the point of 90% degradation. There were no other consistent changes to banding pattern. At the point of 40% degradation, these changes to banding had been evident in only 1 of the 5 IPU treated soils (site D, data not shown). In soil from transect 2, changes to banding pattern occurred only in soil from the single site (site E) in which the MPN analysis had revealed identical proliferation of IPU-degrading organisms to sites from transect 1. In this site, the point of 90% IPU degradation coincided with appearance of a band in the same position as band DS1 recorded in transect 1 (Fig. 3b). Nine months following the start of the experiment, there were no consistent differences in DGGE banding pattern in IPU treated and untreated soil from either transect. By this time band DS1 had disappeared from IPU treated soils, and the intensity of band DS2 had returned to that of the control soil (data not shown).



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  		FIG. 3. Analysis of 16S rRNA PCR products by DGGE in control and IPU treated samples, at the point of 90% IPU degradation. Lanes: M, marker; 1 to 5, control, unamended soil; 6 to 10, IPU-treated soil. Lanes 1 to 5 and lanes 6 to 10 correspond to sites B to F in transects 1 and 2. Positions of cloned bands DS1 and DS2 indicated. (a) Transect 1; (b) transect 2.

Phylogenetic relationships of 16S rRNA sequences from isolates and DGGE bands.
Enrichment culture provided a Gram-negative isolate, strain F35, which was able to degrade IPU with the temporary appearance of the metabolites MDIPU and 3-(4-isopropylphenyl)-urea. The partial 16S rRNA sequence of strain F35 was found to be different to that of DGGE bands DS1 and DS2 (Fig. 4). However, strain F35 and both DGGE bands showed homology to members of the genus Sphingomonas. DGGE band DS2 showed 100% sequence identity with Sphingomonas sp. strain SRS2 (accession number AJ251638), an IPU-degrading culture obtained from Deep Slade field by Sørensen et al. (21), and Sphingomonas trueperi (accession number X97776). Strain SRS2 and Sphingomonas trueperi (X97776) have a 16S rRNA gene sequence similarity of 95.4% (415-bp overlap). DGGE band DS1 was clearly separated from band DS2, showing 98.8% sequence identity to Sphingomonas terrae (accession number D13727). Strain F35 had 100% homology to Porphyrobacter sp. (accession number AB033325), which is classified within the genus Sphingomonas.



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  		FIG. 4. Distance tree constructed with partial (169 bp) 16S rRNA sequences, showing the relationships of the IPU-degrading isolates and DGGE PCR-products to members of the genus Sphingomonas. Scale, 0.1% estimated changes.

Effect of pH on IPU metabolism by Sphingomonas sp. strain SRS2.
SRS2 showed no capacity to degrade IPU when inoculated into sterile soil at pH 6.5, and inoculation of strain SRS2 into nonsterile pH 6.5 soil had no effect on the rate of IPU degradation (Fig. 5a). When strain SRS2 was inoculated into sterile pH 7.5 soil, there was a delay of 20 days before IPU degradation was initiated, with complete degradation occurring within 15 days. When inoculated into nonsterile soil, SRS2 caused a rapid stimulation of IPU degradation, so that after 15 days only 10% of the IPU remained in inoculated soil, compared to 25% in uninoculated nonsterile soil (Fig. 5b).



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  		FIG. 5. Degradation of IPU in sterile and nonsterile soil by Sphingomonas sp. strain SRS2. ({blacktriangleup} sterile, inoculated; • nonsterile, uninoculated; {blacksquare} nonsterile, inoculated). Error bars represent standard errors of the mean. (a) Soil at pH 6.5; (b) soil at pH 7.5.

In liquid culture experiments, strain SRS2 showed a pH optimum of 7.5, with activity dropping sharply above this pH, and below pH 7 (Fig. 6). Isolate F35 was found to have similar pH optima to strain SRS2 (data not shown).



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  		FIG. 6. Effect of pH on mineralization of IPU by Sphingomonas sp. strain SRS2 in liquid culture. Error bars represent standard errors of the mean.


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DISCUSSION
 
Most xenobiotic degrading bacteria have been isolated from the environment following successive enrichment of samples with the xenobiotic of interest. There is evidence that the nature of bacteria contributing to degradation can change as enrichment proceeds, because of mechanisms such as gene transfer between related and unrelated bacteria (10, 11, 18). Although numerous pesticide-degrading bacteria have been isolated from soil using enrichment techniques, demonstration that the isolate actually has a role in degradation in situ has not previously been shown. Our results show that Sphingomonas spp. are involved in IPU degradation in situ in agricultural soil.

Other studies have shown that application of xenobiotics, including pesticides, can change bacterial community structure, although the functional significance of these changes has not been demonstrated. Using 16S rRNA-DGGE, El-Fantroussi et al. (9), showed that repeated application of phenyl-urea herbicides to field plots reduced bacterial diversity, with certain unculturable bacteria disappearing from herbicide treated plots. Similarly, Crecchio et al. (6) found that application of the herbicide prometryne to soil reduced bacterial diversity, although there was evidence for increases in the intensity of some DGGE bands, which was interpreted as evidence for involvement of those organisms in growth-linked metabolism of the compound. MacNaughton et al. (15) showed that members of the {alpha}-proteobacteria and Flexibacter-Cytophaga-Bacteroides groups increased their representation within oil contaminated soil, although the contribution of these organisms to oil degradation was not determined.

Our data, together with that of earlier studies (3, 26) suggests that spatial variability in IPU degradation rates across Deep Slade field is largely the result of direct effects of soil pH on IPU degradation by Sphingomonas spp., with a pH above 7 generally necessary for rapid growth-linked degradation. The results indicate a close similarity between isolate SRS2 and the DGGE band DS2, and therefore a role for strains closely related to SRS2 in IPU degradation in the pH 7.5 soils. Although growth-linked metabolism of IPU can evidently occur at sites with pH of 6.5 or less, it is delayed relative to that of the high pH sites. The components of the Sphingomonas community contributing to degradation at the low pH sites is unclear, because changes to 16S rRNA DGGE band patterns were observed only at a single low pH site, at which only the DS1 Sphingomonas sp. band was shown to increase in intensity.

The role of the DGGE band DS1 and strain F35 components of the Sphingomonas sp. community in IPU degradation remains unclear. The appearance of the DS1 DGGE band during growth-linked IPU degradation in soil from transect 1, and the single soil from transect 2 which exhibited rapid growth-linked metabolism, suggests that it represents a bacterium involved in metabolism of IPU.

Sphingomonas isolate SRS2 is able to metabolize IPU to CO2 and biomass, involving successive N-demethylations followed by cleavage of the urea group and mineralization of the phenyl structure (21). The main metabolite detected in strain SRS2 cultures is MDIPU, and recent results suggest that the metabolites beyond MDIPU occur only intracellularly, without release of intermediates from SRS2 cells (12). This could indicate that either the DS1 and DS2 band Sphingomonas sp. compete for IPU, or that one utilizes cell products and biomass of the other as an energy source. The latter mechanism has recently been suggested for an uncharacterized soil bacterium (designated strain SRS1) which proliferates during growth in coculture with Sphingomonas sp. strain SRS2 with IPU as sole carbon and nitrogen source (22). Another possibility is that DS1 represents an MDIPU-degrading bacterium, given that this metabolite has previously been shown to be readily metabolized by communities of soil bacteria (20).

The role of Sphingomonas sp. strain F35 in IPU degradation in the soil is also uncertain. Although an involvement of F35 in the degradation of IPU in soil cannot be ruled out, it is possible that the serial enrichment steps, prior to isolation, could have resulted in the transfer of IPU-degradative genes to strain F35. Newby et al. (18) showed that when a 2,4-dichlorophenoxyacetic acid (2,4-D) degrading strain of Ralstonia eutropha was inoculated into a soil bioreactor, the degradative genes, which were located on plasmids, could transfer to other Ralstonia strains, as well as strains of Burkholderia, during degradation of applied 2,4-D. Further, the diversity of transconjugants increased with further addition of 2,4 D. It is possible that similar processes have occurred in Deep Slade field during the 20 years over which IPU has been applied to it, resulting in a variety of Sphingomonas sp. contributing to IPU degradation.

Sørensen et al. (22) found that Sphingomonas sp. strain SRS2 requires the presence of other bacteria to facilitate IPU degradation in soil, and that the role of other bacteria could be simulated by the provision of specific amino acids. The degradation of IPU by SRS2 in sterile soil that occurred at pH 7.5 could have been permitted by amino acids released from biomass on autoclaving.

The structure of soil microbial communities is known to vary considerably within ecosystems, reflecting vegetation induced differences in soil properties and C inputs (14). However, little is known of the extent of variability in the spatial distribution of specific functional microbial groups or genes within soil. Further, although soil bacteria with the potential to metabolize diverse xenobiotics have been described (1), the distribution of such organisms within the soil, and factors influencing their activities have rarely been elucidated. Our findings indicate that the sharp pH optimum for degradation by IPU-degrading Sphingomonas spp., which limits degradation in soil with pH less than 7.0, is a key factor controlling field scale patterns of IPU degradation.

In the case of the phenyl-urea herbicide diuron, Cullington and Walker (8) found that degradation at the field scale was highly variable, with a diuron degrading Arthrobacter sp. strain apparently located at only 1 of 10 sites sampled within a field, with distribution associated with neither application history or soil pH (8, 24). However, other pesticide degrading bacteria do have pH requirements similar to those of strain SRS2 and F35, and which could similarly affect patterns of pesticide degradation at the field scale (5, 13).


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ACKNOWLEDGMENTS
 
We thank the Biotechnology and Biological Sciences Research Council and the Department of Environment, Food and Rural Affairs, for financial support; Eve Shaw for technical assistance; John Stephen for providing the DGGE marker; and Leo Calvo-Bades, Margaret Ousley, and Oliver Price for useful advice and discussion.


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FOOTNOTES
 
* Corresponding author. Mailing address: Horticulture Research International, Wellesbourne, Warwick CV35 9EF, United Kingdom. Phone: 44 (0) 1789 470382. Fax: 44 (0) 1789 470382. E-mail: gary.bending{at}hri.ac.uk. Back


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Applied and Environmental Microbiology, February 2003, p. 827-834, Vol. 69, No. 2
0099-2240/03/$08.00+0     DOI: 10.1128/AEM.69.2.827-834.2003
Copyright © 2003, American Society for Microbiology. All Rights Reserved.



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