Chemosphere 66 (2007) 664–671
www.elsevier.com/locate/chemosphere
Microbial aspects of the interaction between soil depth
and biodegradation of the herbicide isoproturon
Gary D. Bending *, M. Sonia Rodriguez-Cruz
1
Warwick HRI, University of Warwick, Wellesbourne, Warwick CV35 9EF, UK
Received 22 June 2006; received in revised form 28 July 2006; accepted 28 July 2006
Available online 22 September 2006
Abstract
Factors controlling change in biodegradation rate of the pesticide isoproturon with soil depth were investigated in a field with sandyloam soil. Soil was sampled at five depths between 0–10 and 70–80 cm. Degradation rate declined progressively down the soil profile,
with degradation slower, and relative differences in degradation rate between soil depths greater, in intact cores relative to sieved soil.
Neither the maximum rate of degradation, or sorption, changed with soil depth, indicating that there was no variation in bioavailability.
Differences in degradation rate between soil depths were not associated with the starting population size of catabolic organisms or the
number of catabolic organisms proliferating following 100% degradation. Decreasing degradation rates with soil depth were associated
with an increase in the length of the lag phase prior to exponential degradation, suggesting the time required for adaptation within communities controlled degradation rates. 16S rRNA PCR denaturing gradient gel electrophoresis showed that degradation in sub-soil
between 40–50 and 70–80 cm depths was associated with proliferation of the same strains of Sphingomonas spp.
2006 Elsevier Ltd. All rights reserved.
Keywords: Pesticide; Leaching; Biodegradation; Bioavailability; Sub-soil; Sphingomonas spp.
1. Introduction
The major environmental concern arising from pesticide
use is the capacity of pesticides to leach from soil and contaminate water resources (Kookana et al., 1998). The
amount of pesticides leaching through soil reflects the interaction of degradation and sorption processes in both topsoil and sub-soil (Fomsgaard, 1995). Many pesticides are
degraded by cometabolism in which degradation follows
first order kinetics, with the organisms responsible apparently showing no capacity to proliferate following degradation of the compound. Other pesticides are degraded by
growth-linked metabolism, in which organisms responsible
for biodegradation have adapted to use the pesticide as an
*
Corresponding author. Tel./fax: +44 24 76575057.
E-mail address: gary.bending@warwick.ac.uk (G.D. Bending).
1
Present address: Institute of Natural Resources and Agrobiology
(CSIC), Department of Environmental Chemistry and Geochemistry,
Salamanca 37008, Spain.
0045-6535/$ - see front matter 2006 Elsevier Ltd. All rights reserved.
doi:10.1016/j.chemosphere.2006.07.099
energy and nutrient source, resulting in cell proliferation
and an increase in degradation rate over time (Aislabie
and Lloyd-Jones, 1995).
Degradation rates of pesticides are usually assumed to
decrease down the soil profile (Fomsgaard, 1995). However,
in some instances degradation rates of pesticides susceptible
to both cometabolic and growth-linked degradation can be
greater in sub-soil than in top-soil. The precise relationship
between top- and sub-soil degradation rate can vary between
different compounds at single sites, and at different sites for
individual compounds (Di et al., 1998; Karpouzas et al.,
2001; Mills et al., 2001). The reasons for contrasting patterns
of degradation rates in sub- and top-soil are unclear.
A number of counteracting biotic and abiotic factors
could be important for determining differences in pesticide
degradation rate through the soil profile. The size of the
microbial community, which decreases with soil depth
(Fomsgaard, 1995) could determine both the availability
of suitable microbial strains and genetic elements for adaptation to enable pesticide degradation, the survival of
G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
adapted strains and the extent to which the activities of
adapted strains are limited by competition. Sorption, which
may reflect bioavailability (Jensen et al., 2004), is closely
linked to organic matter content, and declines with soil
depth. A range of environmental factors such as oxygen
and temperature, which can affect microbial growth rates
and pesticide degradation, also change with soil depth
(Fomsgaard, 1995; Vink and van der Zee, 1997; Williams
et al., 2003).
Microbes contributing to degradation of xenobiotics in
top-soil have traditionally been investigated using enrichment methods (e.g. Sørensen et al., 2001), although the catabolic organisms obtained using these procedures may not
reflect those organisms acting in situ (Newby et al., 2000).
Little is known about the nature of catabolic communities
acting in sub-soil. Furthermore, catabolic strains obtained
from top-soil by isolation, or communities characterised
using in situ molecular profiling techniques, have typically
been investigated in soil samples derived from single environmental locations (e.g. Cullington and Walker, 1999;
Sørensen et al., 2001; Singh et al., 2003). The extent to
which there is spatial variability in the structure of catabolic communities, at both the field and landscape scale,
has received little attention.
Bending et al. (2003) found that catabolism of the herbicide isoproturon was associated with proliferation of
Sphingomonas spp. in a transect over a high pH (7.0–7.5)
area of a field, but within a low pH (pH 6.0–6.5) area located
just 50 m away, it appeared that different strains were
involved in catabolism. Within agricultural fields, vertical
changes in soil properties with soil depth are much greater
than the horizontal spatial variability in soil properties
which can occur within top-soil (Rodriguez-Cruz et al.,
2006). Clearly, such changes in soil physico-chemical and
biological properties with soil depth could result in the selection of different catabolic communities between top- and
sub-soil, although this has not previously been investigated.
The aims of the current study were to investigate the
interactions between catabolic communities, physico-chemical properties and pesticide degradation through the soil
profile. Experiments were performed to address the following questions: 1. Are changes in pesticide bioavailability
with soil depth associated with changes in the rate of biodegradation and its kinetics? 2. Does change in biodegradation rate with soil depth reflect differences in the size of the
initial catabolic community, or the extent to which the catabolic community is able to proliferate? 3. Is there vertical
spatial variability in the nature of communities contributing to pesticide catabolism?
2. Materials and methods
2.1. Pesticides and pesticide treatment history
Studies focussed on the herbicide isoproturon (3-(4-isopropylphenyl)-1,1-dimethylurea). Isoproturon is a member
of the phenylurea group of herbicides, and together with
665
related compounds are frequently detected as contaminants
of groundwater and surface freshwater in Europe (Sørensen et al., 2003). Sampling occurred in Long Close field
on the farm at Warwick HRI, Wellesbourne, Warwickshire, UK. The soil is a sandy loam of the Wick series
(Whitfield, 1974), and the field had a history of isoproturon
use (1999, 2001).
2.2. Soil collection
Soil was collected from five depths at three sampling
locations. Three pits (A–C) separated by 60 m were excavated to 1 m depth using a mechanical digger, in February
2003. One side of each pit was further excavated using a
surface sterilised trowel, so that the face was free of loose
soil. Soil was collected from 0–10, 20–30, 40–50, 60–70
and 70–80 cm depth using two methods. 1. From each
depth approximately 2 kg soil was collected using a trowel
and placed into a polythene bag. The trowel was surface
sterilised with ethanol between the collection of each soil
sample. Soil was spread onto clean polythene bags and left
on the bench overnight to reduce moisture content, before
being sieved (<3 mm) using surface sterilised sieves. 2. In
order to maintain the physical and microbiological integrity of the soil, further samples were taken using intact
10 · 5 cm pre-sterilised stainless steel cores. Two cores were
obtained at each depth from each hole by hammering the
core horizontally into soil. Following removal, the top
and bottom of each core were sealed with parafilm.
2.3. Analysis of soil characteristics
In the pre-sieved soil, total organic matter, microbial
biomass-N, dehydrogenase activity and pH were measured
by procedures described in Bending et al. (2006). Clay,
sand and silt content were determined according to Day
(1965).
2.4. Pesticide application
For pre-sieved soil, commercial isoproturon formulation
(Atlas Crop protection, Doncaster, UK) was dissolved in
distilled H2O and added to single 300 g fw portions of soil
from each location to provide 5 mg pesticide kg 1 soil, and
further H2O was added to bring the water holding capacity
to 40%. Each soil was mixed thoroughly by hand, and then
further mixed by passing through a <3 mm sieve five times.
Each soil was transferred to a sterile polypropylene container which was loosely capped and incubated at 15 C.
Moisture content was maintained by the addition of sterile
distilled water as necessary (usually once each week).
In the case of the two soil cores collected from each sampling location, four 250 ll aliquots of the commercial formulation of isoproturon in water were injected centrally
at 2 cm depths to give a final concentration of 5 mg kg 1
soil. The soil cores were sealed base and top with parafilm
and incubated vertically at 15 C in the dark.
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G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
2.5. Pesticide extraction and analysis
The pre-sieved soils were sampled at regular intervals
over a 2-month period, with extraction and HPLC as
described by Bending et al. (2006). The cores were sampled
after 20 days incubation. Soil was pushed from the cores
and allowed to dry for 2 h, before the soil was mixed by
hand and sieved (<3 mm) five times. Sub-samples (10 g)
of each soil were dried in an oven at 110 C overnight to
determine soil moisture content. Pesticides were extracted
from each soil sample and analysed using the procedures
described above. Sorption of isoproturon was determined
using a batch mixing method, and adsorption distribution
coefficients (Kd) measured as described by Rodriguez-Cruz
et al. (2006).
2.6. Interactions between depth and the isoproturon
degrading community
Prior to isoproturon application, and at the point of
100% degradation, the size of the isoproturon catabolising community was determined in pre-sieved soil by most
probable number analysis according to Bending et al.
(2001). PCR-16S rRNA-denaturing gradient gel electrophoresis (DGGE) was conducted in order to determine
whether organisms proliferating following treatment with
isoproturon could be identified. Portions of pre-sieved soil
were removed immediately following isoproturon addition,
and at the point of 100% pesticide degradation. DNA was
extracted from 1 g fw portions of soil by bead-beating
using a Cambio (Cambridge, UK) Ultraclean soil DNA
extraction kit. Partial eubacterial 16S rRNA gene fragments were amplified using primers described by Muyzer
et al. (1993) at positions 341f and 534r (Escherichia coli
numbering), using a Hybaid Omnigene thermocycler (Ashford, UK). DGGE gels were set up according to Muyzer
et al. (1993) using an Ingeny PhorU System (Amsterdam)
with 8% acrylamide, and a denaturant gradient of 20–
70% (100% denaturant was equivalent to 7 M urea with
40% vol/vol formamide). The gels were run at 70 V and
60 C for 18 h. The gels were stained with ethidium
bromide (0.5 mg l 1) and visualised under UV light on an
Imago Imaging system (B and L systems, the Netherlands).
Bands of interest were cut from the gel and left overnight
in 50 ll MiliQ H2O at 4 C for 18 h. After centrifuging,
DNA in the supernatant was amplified as described above.
Re-amplified bands were run against the original sample to
check motility and purity.
The PCR products were purified using a QIAquick PCR
purification kit (Qiagen Ltd., Dorking, UK), and then
cloned using a TOPO TA Cloning kit (Invitrogen, Paisley,
UK). Sequencing reactions were performed according to
manufacturer’s instructions, on a Hybaid PCR multiblock
system (Hybaid, Middlesex, UK), using a PRISM BigDye
Terminator Cycle Sequence reaction kit (Applied Biosystems, Warrington, UK). Products were analysed on an
Applied Biosystems 377 DNA sequencer.
The partial 16S rRNA-DGGE sequences were edited
and assembled using the DNAstar II sequence analysis
package (Lasergene Inc., Madison, Wisconsin, USA).
Sequences were compared to those on the EMBL database
using the programme FASTA3. Reference partial 16S
rRNA sequences were gathered from the EMBL database,
and from previous studies describing isolated isoproturon
degrading bacteria, and environmental sequences shown
previously to proliferate during degradation of isoproturon
in Deep Slade field, which is adjacent to Long Close (Bending et al., 2003). The partial DGGE and reference sequences
corresponding to the 169 bp fragment were used for phylogenetic analysis using the PHYLIP packages SEQBOOT,
DNADIST and NEIGHBOR. The dendrogram was generated using neighbour-joining analysis. The nucleotide
sequences for LC1 and LC2 have been deposited in the
EMBL nucleotide sequence database under the accession
numbers AM295188 and AM295189 respectively.
2.7. Statistical analysis
Analysis of variance was used to determine the significance of differences in soil parameters and degradation
characteristics between soil depths. The Gompertz model
was found to provide best fit to the degradation kinetics,
and was used to obtain time to 50% degradation (DT50)
values, the length of lag phase prior to exponential degradation and the maximum mineralization rate (i.e. rate of
decline during the exponential degradation phase). All statistical analyses were performed using GenStat (7th edition,
VSN International Ltd.).
3. Results
3.1. Variation in soil biological and physico-chemical
properties with soil depth
There were significant progressive declines in % organic
matter (OM), biomass and dehydrogenase, and a significant
increase in pH, down the soil profile, demonstrating a clear
gradient in soil chemical and biological properties with depth
(Table 1). Percentage of organic matter declined from 2.70 at
0–10 cm to 1.11 at 70–80 cm depth. pH increased from 7.01
at 0–10 cm to 8.17 at 70–80 cm depth. Biomass declined from
68.8 mg C kg 1 soil to 9.5 mg C kg 1 soil at 70–80 cm depth.
Similarly, there was a decrease in dehydrogenase activity
from 43.9 lg triphenyl formazan (TPF) g 1 soil at 0–10 cm
to 1.7 lg TPF g 1 soil at 70–80 cm. There were no significant
changes in clay, silt or sand with depth.
3.2. Variation in isoproturon degradation rates down the soil
profile
In pre-sieved top-soil, isoproturon degradation proceeded rapidly without a lag phase (Fig. 1). In sub-soil, there
was a lag phase followed by a period of rapid degradation.
Degradation rate and DT50 declined progressively down
G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
667
Table 1
Changes in soil chemical and microbial properties with soil depth
Soil depth
OM (%)
0–10 cm
20–30 cm
40–50 cm
60–70 cm
70–80 cm
LSD (P > 0.05)
Significance of effect of depthb
2.70
2.37
2.24
1.54
1.11
0.29
***
Clay (%)
21.4
20.9
23.6
32.2
31.9
10.9
NS
Silt (%)
8.7
8.8
8.0
11.0
11.7
4.5
NS
Sand (%)
pH
69.9
70.3
68.4
56.8
56.4
20.2
NS
Biomass
(mg C kg
7.01
7.08
7.74
8.05
8.17
0.45
**
1
Dehydrogenase
(lg TPFa g 1 dw soil)
soil)
68.8
66.9
45.6
16.3
9.5
16.0
***
43.9
23.0
16.4
5.7
1.7
14.7
***
Data represent average of the three replicate sampling locations (A–C) at each depth.
a
Triphenyl formazan.
b
NS, not significant; ** significant, P < 0.01; *** significant, P < 0.001.
3.3. Interactions between depth and isoproturon degrading
communities
Prior to isoproturon application the number of isoproturon degrading organisms was lower than the detection
100
LSD (P<0.05)
% isoproturon remaining
the soil profile (Fig. 1, Table 2). In top-soil, DT50 was 16.8
and 14.4 days, at 0–10 and 20–30 cm respectively, while in
sub-soil it declined from 20.5 days at 40–50 cm depth to
29.5 days at 70–80 cm. The variation in DT50 was the result
of the length of the lag phase prior to the period of rapid
degradation. In sub-soil samples the length of the lag phase
increased down the soil profile, from 18.3 days at 40–50 cm,
to 26.6 days at 70–80 cm. Soil depth had no significant effect
on the maximum degradation rate of isoproturon, which
ranged between 0.54 and 0.86 lg isoproturon g 1 soil day 1
(Table 2). Additionally, there was no significant change in
Kd with soil depth (Table 2). Isoproturon DT50 was significantly correlated with biomass (r = 0.56, P < 0.05) and
dehydrogenase (r = 0.57, P < 0.05).
In the case of soil cores, degradation rate decreased significantly with soil depth, with 37.4% isoproturon remaining after 20 days at 0–10 cm depth, and 90.9% remaining at
70–80 cm depth (Table 2). Compared with pre-sieved soil,
degradation was slower in intact cores at all depths (Table
2, Fig. 1).
80
60
40
20
0
0
10
20
30
40
50
60
70
Time (days)
Fig. 1. Degradation of isoproturon in top- and sub-soil samples. Data
represent average of three replicate sampling locations at each depth: (d)
0–10 cm; (.) 20–30 cm; (s) 40–50 cm; ($) 60–70 cm; (h) 70–80 cm.
limit of 100 degraders g 1 soil in all samples. At the point
of 100% degradation numbers of isoproturon degrading
organisms had increased in all samples, although there
were no significant differences in the number proliferating
at the different soil depths (Table 2).
Table 2
Changes in isoproturon degradation and sorption characteristics with soil depth
Soil depth
% Isoproturon remaining
in cores after 20 days
DT50 (days)a
Lag phase (days)a
Maximum degradation
rate (lg g 1 fw soil day 1)a
Number of degraders
(log MPN g 1 fw soil)a,b
Kd
0–10 cm
20–30 cm
40–50 cm
60–70 cm
70–80 cm
LSD (P > 0.05)
Significance of
effect of depthc
37.4
42.7
68.7
88.1
90.9
26.0
***
16.8
14.4
20.5
26.2
29.5
2.4
**
0.0
0.0
18.3
23.0
26.6
2.1
***
0.54
0.54
0.84
0.75
0.86
0.46
NS
3.90
3.42
3.43
3.75
3.56
2.20
NS
0.96
0.85
0.61
0.40
0.66
0.40
NS
Data represent average of the three replicate sampling locations (A–C) at each depth.
a
In sieved soil.
b
At the point of 100% isoproturon degradation.
c
NS, not significant; ** significant, P < 0.01; *** significant, P < 0.001.
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G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
Sphingomonas stygialis U20775
Erythrobacter longus M59062
Sphingomonas rosa D13945
Sphingomonas sp. JSS-7 AF131295
Sphingomonas adhaesiva D13722
Sphingomonas sp. JSS-26 AF131296
Sphingomonas terrae D13727
DS1 AJ509085
Sphingomonas yanoikuyae D16145
Porphyrobacter sp. KK348 ab033325a
Sphingomonas sp. Isolate 782
Sphingomonas sp. BN6 X94098
Erythrobacter litoralis AB013354
Zymomonas mobilis AF281031
Sphingomonas paucimobilis U37341
Sphingomonas sp. UN1F1 U37345
Sphingomonas sp. SAFR-028 AY167833
Sphingomonas trueperi X97776
DS2 AJ509086
Sphingomonas sp. Isolate SRS2 AJ251638
LC2
Sphingomonas parapaucimobilis D13724
Sphingomonas paucimobilis U37337
LC1
Sphingomonas sp. SYK6 D16149
Sphingomonas herbicidovorans AB042233
0.005
Fig. 2. Results of 16S rRNA PCR–DGGE profiling. (a) Analysis of DGGE profile in control (C) and isoproturon treated (+IPU) soil at the point of 100%
degradation. Example represents 70–80 cm depth at location A. LC1 and LC2, which appeared in isoproturon treated sub-soil, are indicated. (b)
Neighbour-joining tree showing phylogenetic relationships of partial (169 bp) 16S rRNA sequences of bands LC1 and LC2. Isoproturon catabolising
isolates (F35 and SRS2) and 16S rRNA DGGE PCR products associated with isoproturon degradation in Deep Slade field (DS1 and DS2) are highlighted
in bold for comparison.
Treatment with isoproturon had no effect on DGGE
banding in top-soil samples (data not shown). However,
in all sub-soil samples from 60–70 and 70–80 cm depth,
and two samples from 40–50 cm depth, isoproturon degradation was associated with the appearance of two new
DGGE bands, LC1 and LC2, which were not present in
untreated soil (Fig. 2a). The sequence of the bands differed
in three positions over the 169 bp sequenced. There were no
other consistent changes to banding, which occurred in
more than one sample, following isoproturon addition.
Sequencing of bands LC1 and LC2 demonstrated homology to the Sphingomonas spp. group (Fig. 2b). LC1 and
LC2 clustered with the Sphingomonas paucimobilis group.
This contrasted with the groupings of isoproturon catabolic strains which had been characterised by DGGE and
isolation in earlier studies in the adjacent Deep Slade field
(Bending et al., 2003).
4. Discussion
Most studies have found decreased degradation rates of
pesticides with increasing soil depth (Fomsgaard, 1995).
However, a number of studies with compounds with
similar sorption to isoproturon have found higher rates
of degradation in sub-soil relative to top-soil (Di et al.,
1998; Karpouzas et al., 2001; Mills et al., 2001). Decrease
in sorption, and an increase in bioavailability, associated
with a decrease in organic matter content with soil depth
has been proposed as the mechanism by which degradation
rates could increase with soil depth. In our study, degradation rate determined using two alternative methods showed
clear decline with soil depth.
While sorption may not provide an accurate measure of
bioavailability, differences in sorption between samples for
individual pesticides may be indicative of contrasting
bioavailability (Jensen et al., 2004). Our data demonstrated that sorption did not change with soil depth,
despite a substantial reduction in organic matter. Boivin
et al. (2005) showed that for 13 top-soils with OM ranging
from between 1% and 6%, isoproturon sorption was
strongly correlated with organic matter content. However,
variation in % OM in our study ranged between only 2.7%
and 1.1%, and our study agrees with that of Boivin et al.
(2005), which suggests that differences in isoproturon
sorption do not occur over this range of OM contents.
Furthermore, there was no significant change with soil
depth in the maximum rate of degradation during the
exponential degradation phase, suggesting that soil
depth had no significant effect on pesticide bioavailability.
These findings indicate that bioavailability was not a
G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
factor influencing the adaptation of organisms to degrade
the pesticide or the growth and dynamics of adapted
organisms.
The most probable number of isoproturon degrading
organisms was below detection limits in samples from all
soil depths prior to application. The starting number of
catabolic organisms was clearly very low in all samples,
and cannot therefore account for differences in degradation
rate through the soil profile. Similarly, there was no difference in the number of catabolic organisms which had proliferated following complete degradation of isoproturon, or
in the maximum rate of degradation, suggesting that differences in the rate of proliferation of adapted catabolic
organisms, or the population density achieved, were not
responsible for the decline in degradation rate with soil
depth. The decline in degradation rate with depth clearly
reflected differences in the length of the lag phase prior to
exponential decay. The absence of a lag phase in top-soil
samples indicated that either adapted organisms were present, or that the process of adaptation occurred very rapidly. Since the soil had received previous applications of
the compound, the presence of an adapted population
would be expected (Bending et al., 2003, 2006). The fact
that MPN analysis could not detect isoproturon degraders
even in those samples in which there was no lag phase prior
to exponential degradation indicates that the technique
lacks sensitivity.
The extended lag phase in sub-soil samples could reflect
the time taken for organisms to adapt to degrade the pesticide. Amounts of pesticide leaching to sub-soil are likely
to be small and spatially variable (Walker et al., 2001).
Furthermore, the patterns of distribution of microbial
communities in sub-soil are different to those in top-soil,
and in particular show greater spatial variability (Nunan
et al., 2002). Adaptation to xenobiotic catabolism appears
to involve a variety of processes, including assembly of horizontally transmitted genes, mutations and gene rearrangements (Springael and Top, 2004). Clearly, community
structure, the patterns of contact between microbial communities and the pesticide, spatial patterns in the growth
of catabolic microbes, and the rate of gene exchange
between microbes which could control adaptation, could
be fundamentally different in sub-soil and top-soil, and
contribute to in the increasing length of the lag phase prior
to exponential degradation with soil depth. The close relationship between DT50 and biomass/dehydrogenase suggests that the size of the overall microbial community
contributed to the time taken for organisms to adapt to
degrade the pesticide, even though a specific catabolic
strain appeared to be involved in degradation.
Degradation of isoproturon was slower in the intact
cores relative to sieved soil, and the relative difference in
degradation rate through the soil profile was greater in
the cores relative to the sieved soil. In addition to maintaining microbiological and structural integrity of the soil, the
pesticide application method was by necessity different in
the sieved soil and intact cores, so that the sieved soil
669
received a uniform distribution of pesticide, and in cores
the pesticide was likely to have be localised close to the
points of application. Both factors could potentially have
influenced biodegradation. Clearly differences in soil handling and pesticide application method have a major role
in determining pesticide degradation dynamics, and pesticide fate studies should ideally be made in soil in cores in
preference to the highly artificial sieved condition.
In the current study, no changes occurred to 16S rRNA
DGGE banding following isoproturon metabolism in
top-soil, although in most sub-soil samples, isoproturon
degradation was associated with the appearance of two
new bands which showed homology to Sphingomonas
spp. Furthermore, the same bands appeared at the three
spatially separated sampling locations, and at depths
between 40–50 and 70–80 cm. Similarly, degradation of a
range of pesticides in environmental samples has been
shown to be associated with the appearance of single or
several bands in DGGE profiles, reflecting proliferation
of strains involved in catabolism (Bending et al., 2003;
Singh et al., 2003; Sørensen et al., 2005).
Several Sphingomonas spp. are known to contain multiple heterogeneous 16S rRNA gene copies, and it is therefore unclear whether LC1 and LC2 represent the same or
different strains (Leys et al., 2004). Since DGGE profiles
only dominant members of the microbial community
(van Elsas et al., 1998), in order to change DGGE banding,
changes in population sizes must reach a threshold relative
to the size of background microbial communities. The size
of the isoproturon degrading community that had developed following complete catabolism of the compound
was similar in all soil samples. However, microbial biomass
declined markedly with soil depth. Clearly the proliferation
of IPU degrading organisms relative to the overall size of
the microbial community was higher in sub-soil than topsoil, so that DGGE banding changes occurred.
Previous work in Deep Slade field, which is next to Long
Close, demonstrated that a strain of Sphingomonas
spp. (SRS2) was involved in growth-linked degradation
of isoproturon in top-soil (Bending et al., 2001, 2003).
The degradation pathway for isoproturon, involving
sequential demethylation of the isoproturon side chain is
the same in the two fields. However, the current study suggests that the specific Sphingomonas spp. strains involved in
isoproturon catabolism in Long Close field are different to
those in Deep Slade. However, due to the small size of the
DNA fragment sequenced, in situ RNA profiling and stable
isotope probing in combination with isolation methods
(Mahmood et al., 2005; Sørensen et al., 2005) would be
required to provide unequivocal evidence. The strains
involved in degradation of individual xenobiotics typically
vary between different sites, even if the genera are the same
(Haggblom, 1992), although the catabolic genes involved
can be highly conserved at different geographical locations
(Haggblom, 1992; Ralebitso et al., 2002). Our data suggest
that different strains of Sphingomonas spp. have adapted to
degrade isoproturon in adjacent fields, suggesting that the
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G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
landscape scale over which adaptation occurs within specific bacterial strains is small.
5. Conclusions
Rates of pesticide degradation generally decline with soil
depth, although there are many reports of increasing degradation rates with soil depth. This study determined the
relative importance of bioavailability, the population size
of catabolic organisms and adaptation time to the change
in degradation rate of the pesticide isoproturon with soil
depth. The rate of decline in the biodegradation rate of isoproturon with soil depth was not associated with changes
in bioavailability, as measured by sorption and the exponential degradation rate, or with the number of catabolic
organisms prior to application, or the number proliferating
following complete degradation. The decline in degradation rate could be accounted for solely by the length of
the lag phase prior to exponential degradation of the compound, suggesting that the time taken for microbial communities to adapt to degrade the compound was the
factor controlling the decline in degradation rate with soil
depth. Adaptation to degrade the pesticide was related to
the size of the microbial biomass. 16S rRNA-DGGE analysis suggested that degradation of isoproturon from different sites across the field and sub-soil depths was associated
with appearance of the same Sphingomonas spp., suggesting no vertical or horizontal variation in the composition
of the catabolic community within the field. However these
strains were different to catabolic Sphingomonas spp.
strains isolated from an adjacent field in earlier studies,
indicating that specific communities of catabolic organisms
can vary subtly over small landscape scales.
Acknowledgements
We thank the late Professor Allan Walker for useful discussion, Dr. Julie Jones for statistical advice, Su Lincoln
and Lucille Marot for technical assistance, and the Department for Environment, Food and Rural Affairs for funding.
M.S. Rodriguez-Cruz thanks the Spanish Ministry of Education and Science for a postdoctoral fellowship award.
References
Aislabie, J., Lloyd-Jones, G., 1995. A review of bacterial degradation of
pesticides. Aust. J. Soil Res. 33, 925–942.
Bending, G.D., Shaw, E., Walker, A., 2001. Spatial heterogeneity in the
metabolism and dynamics of isoproturon degrading microbial communities in soil. Biol. Fertil. Soils 33, 484–489.
Bending, G.D., Lincoln, S.D., Sørensen, S.R., Morgan, J.A.W., Aamand,
J., Walker, A., 2003. 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.. Appl. Environ.
Microbiol. 69, 827–834.
Bending, G.D., Lincoln, S.D., Edmondson, R.N., 2006. Spatial variation
in the degradation rate of the pesticides isoproturon, azoxystrobin and
diflufenican in soil and its relationship with chemical and microbial
properties. Environ. Pollut. 139, 279–287.
Boivin, A., Cherrier, R., Schiavon, M., 2005. A comparison of five
pesticides adsorption and desorption processes in thirteen contrasting
field soils. Chemosphere 61, 668–676.
Cullington, J.E., Walker, A., 1999. Rapid biodegradation of diuron and
other phenylureas by a soil bacterium. Soil Biol. Biochem. 31, 677–686.
Day, P.R., 1965. Particle fractionation and particle-size analysis. In:
Black, C.A., Evans, D.D., White, J.L., Ensminger, L.E., Clark, F.E.
(Eds.), Methods of Soil Analysis, Part 1, Agron. Monogr., vol. 9. ASA,
Madison, WI, USA, pp. 545–566.
Di, H.J., Aylmore, L.A.G., Kookana, R.S., 1998. Degradation rates of
eight pesticides in surface and subsurface soils under laboratory and
field conditions. Soil Sci. 163, 404–411.
Fomsgaard, I.S., 1995. Degradation of pesticides in subsurface soils,
unsaturated zone – a review of methods and results. Int. J. Environ.
Anal. Chem. 58, 231–245.
Haggblom, M.M., 1992. Microbial breakdown of halogenated aromatic
pesticides and related-compounds. FEMS Microbiol. Rev. 103, 29–72.
Jensen, P.H., Hansen, H.C.B., Rasmussen, J., Jacobsen, O.S., 2004.
Sorption-controlled degradation kinetics of MCPA in soil. Environ.
Sci. Technol. 38, 6662–6668.
Karpouzas, D.G., Walker, A., Drennan, D.S.H., Froud-Williams, R.J.,
2001. The effect of initial concentration of carbofuran on the
development and stability of its enhanced biodegradation in top-soil
and sub-soil. Pest Manage. Sci. 57, 72–81.
Kookana, R.S., Baskaran, S., Naidu, R., 1998. Pesticide fate and
behaviour in Australian soils in relation to contamination and
management of soil and water: a review. Aust. J. Soil Res. 36, 715–764.
Leys, N.M.E.J., Ryngaert, A., Bastiaens, L., Verstraete, W., Top, E.M.,
Springael, D., 2004. Occurrence and phylogenetic diversity of Sphingomonas strains in soil contaminated with polycyclic aromatic hydrocarbons. Appl. Environ. Microbiol. 70, 1944–1955.
Mahmood, S., Paton, G.I., Prosser, J.I., 2005. Cultivation-independent
in situ molecular analysis of bacteria involved in degradation of
pentachlorophenol in soil. Environ. Microbiol. 7, 1349–1360.
Mills, M.S., Hill, I.R., Newcombe, A.C., Simmons, N.D., Vaughan, P.C.,
Verity, A.A., 2001. Quantification of acetochlor degradation in the
unsaturated zone using two novel in situ field techniques: comparisons
with laboratory-generated data and implications for groundwater risk
assessments. Pest Manage. Sci. 57, 351–359.
Muyzer, G., Dewaal, E.C., Uitterlinden, A.G., 1993. Profiling of complex
microbial-populations by denaturing gradient gel-electrophoresis analysis of polymerase chain reaction-amplified genes-coding for 16S
ribosomal-RNA. Appl. Environ. Microbiol. 59, 695–700.
Newby, D.T., Gentry, T.J., Pepper, I.L., 2000. Comparison of 2,4dichlorophenoxyacetic acid degradation and plasmid transfer in soil
resulting from bioaugmentation with two different pJP4 donors. Appl.
Environ. Microbiol. 66, 3399–3407.
Nunan, N., Wu, K., Young, I.M., Crawford, J.W., Ritz, K., 2002. . In situ
spatial patterns of soil bacterial populations, mapped at multiple
scales, in an arable soil. Microbial Ecol. 44, 296–305.
Ralebitso, T.K., Senior, E., van Verseveld, H.W., 2002. Microbial aspects
of atrazine degradation in natural environments. Biodegradation 13,
11–19.
Rodriguez-Cruz, M.S., Jones, J.E., Bending, G.D., 2006. Field-scale study
of the variability in pesticide biodegradation with soil depth and its
relationship with soil characteristics. Soil Biol. Biochem. 38, 2910–2918.
Singh, B.K., Walker, A., Morgan, J.A.W., Wright, D.J., 2003. Effects of
soil pH on the biodegradation of chlorpyrifos and isolation of a
chlorpyrifos-degrading bacterium. Appl. Environ. Microbiol. 69,
5198–5206.
Sørensen, S.R., Ronen, Z., Aamand, J., 2001. Isolation from agricultural
soil and characterization of a Sphingomonas sp able to mineralize the
phenylurea herbicide isoproturon. Appl. Environ. Microbiol. 67,
5403–5409.
Sørensen, S.R., Bending, G.D., Jacobsen, C.S., Walker, A., Aamand, J.,
2003. Microbial degradation of isoproturon and related phenylurea
herbicides in and below agricultural fields. FEMS Microbiol. Ecol. 45,
1–11.
G.D. Bending, M.S. Rodriguez-Cruz / Chemosphere 66 (2007) 664–671
Sørensen, S.R., Rasmussen, J., Jacobsen, C.S., Jacobsen, O.S., Juhler,
R.K., Aamand, J., 2005. Elucidating the key member of a linuronmineralizing bacterial community by PCR and reverse transcription-PCR denaturing gradient gel electrophoresis 16S rRNA gene
fingerprinting and cultivation. Appl. Environ. Microbiol. 71, 4144–
4148.
Springael, D., Top, E.M., 2004. Horizontal gene transfer and microbial
adaptation to xenobiotics: new types of mobile genetic elements and
lessons from ecological studies. Trends Microbiol. 12, 53–58.
van Elsas, J.D., Duarte, G.F., Rosado, A.S., Smalla, K., 1998. Microbiological and molecular biological methods for monitoring microbial
inoculants and their effects in the soil environment. J. Microbiol.
Methods 32, 133–154.
671
Vink, J.P.M., van der Zee, S.E.A.T.M., 1997. Effect of oxygen status on
pesticide transformation and sorption in undisturbed soil and lake
sediment. Environ. Toxicol. Chem. 16, 608–616.
Walker, A., Jurado-Exposito, M., Bending, G.D., Smith, V.J.R., 2001.
Spatial variability in the degradation rate of isoproturon in soil.
Environ. Pollut. 111, 407–415.
Whitfield, W.A.D., 1974. The soils of the national vegetable research
station, Wellesbourne. In: Report of the national vegetable research
station for 1973, pp. 21–30.
Williams, G.M., Harrison, I., Carlick, C.A., Crowley, O., 2003. Changes
in enantiomeric fraction as evidence of natural attenuation of
mecoprop in a limestone aquifer. J. Contam. Hydrol. 64, 253–
267.