Functional Redundancy of Linuron Degradation in MicrobialCommunities in Agricultural Soil and Biopurification Systems

Benjamin Horemans,a Karolien Bers,a Erick Ruiz Romero,b Eva Pose Juan,c Vincent Dunon,a René De Mot,d Dirk Springaela

Division of Soil and Water Management, KU Leuven, Heverlee, Belgiuma; Laboratory of Soil Ecology, Cinvestav, Mexico City, Mexicob; Instituto de Recursos Naturales yAgrobiología de Salamanca, Salamanca, Spainc; Centre of Microbial and Plant Genetics, KU Leuven, Heverlee, Belgiumd

ABSTRACT

The abundance of libA, encoding a hydrolase that initiates linuron degradation in the linuron-metabolizing Variovorax sp.strain SRS16, was previously found to correlate well with linuron mineralization, but not in all tested environments. Recently, analternative linuron hydrolase, HylA, was identified in Variovorax sp. strain WDL1, a strain that initiates linuron degradation in alinuron-mineralizing commensal bacterial consortium. The discovery of alternative linuron hydrolases poses questions aboutthe respective contribution and competitive character of hylA- and libA-carrying bacteria as well as the role of linuron-mineral-izing consortia versus single strains in linuron-exposed settings. Therefore, dynamics of hylA as well as dcaQ as a marker fordownstream catabolic functions involved in linuron mineralization, in response to linuron treatment in agricultural soil andon-farm biopurification systems (BPS), were compared with previously reported libA dynamics. The results suggest that (i) or-ganisms containing either libA or hylA contribute simultaneously to linuron biodegradation in the same environment, albeit tovarious extents, (ii) environmental linuron mineralization depends on multispecies bacterial food webs, and (iii) initiation oflinuron mineralization can be governed by currently unidentified enzymes.

IMPORTANCE

A limited set of different isofunctional catabolic gene functions is known for the bacterial degradation of the phenylurea herbi-cide linuron, but the role of this redundancy in linuron degradation in environmental settings is not known. In this study, thesimultaneous involvement of bacteria carrying one of two isofunctional linuron hydrolysis genes in the degradation of linuronwas shown in agricultural soil and on-farm biopurification systems, as was the involvement of other bacterial populations thatmineralize the downstream metabolites of linuron hydrolysis. This study illustrates the importance of the synergistic metabo-lism of pesticides in environmental settings.

Linuron [3-(3,4-dichlorophenyl)-1-methoxy-1-methyl urea] isa phenylurea herbicide that is widely used in various agricul-

ture crops and in orchards (1) but also forms a contaminant insoil, groundwater, and surface water (1, 2). Biodegradation is animportant mechanism for removal of linuron in the environment.In agricultural soils that were treated with the compound on along-term basis, microbial communities respond to linuron ap-plication and specific bacterial populations that use linuron as asole source of carbon, nitrogen, and energy proliferate (2).

The major bacterial pathway for mineralization of linuron in soilis initiated with the hydrolysis of linuron into 3,4-dicholoroaniline(3,4-DCA) and N,O-dimethylhydroxylamine (N,O-DMHA), whichare subsequently mineralized (2). Various bacterial isolates andconsortia that mineralize linuron were reported. In both cases,members of the genus Variovorax are essential. Most linuron-mineralizing single strains are Variovorax species, and in consor-tia, the organism performs at least the initial hydrolysis step (3–6).Recently, gene functions linked with linuron mineralization inVariovorax sp. were identified. The libA gene was identified inVariovorax sp. strain SRS16 and encodes the linuron amidaseLibA, which hydrolyzes linuron into 3,4-DCA and N,O-DMHA(7). Until recently, libA was the only linuron hydrolysis genelinked with linuron mineralization. In several environments suchas agricultural soil and on-farm biopurification systems (BPS)used for the treatment of agricultural pesticide-contaminatedwastewater, the abundance of libA increased in parallel with in-creasing linuron mineralization capacity as a response to linuron

application, indicating a prominent role for libA in linuron min-eralization in environmental settings (8, 9). However, in additionto LibA, other linuron hydrolases were proposed to contribute tolinuron mineralization in Variovorax and in the environment, assome linuron-degrading Variovorax isolates lack a libA homo-logue and libA did not proliferate in all environments that devel-oped linuron mineralization activity upon exposure to linuron(7). Recently, we identified HylA as a second type of linuron hy-drolase in Variovorax sp. strain WDL1 (10). Interestingly, HylA isevolutionarily unrelated to LibA and shows different enzymatickinetic properties (10). The linuron hydrolysis genes in bothWDL1 and SRS16 are combined with highly similar catabolic genemodules encoding the downstream pathway for 3,4-DCA degra-dation. Apparently, the expansion of a 3,4-DCA catabolic pathway

Received 18 December 2015 Accepted 28 February 2016

Accepted manuscript posted online 4 March 2016

Citation Horemans B, Bers K, Ruiz Romero E, Pose Juan E, Dunon V, De Mot R,Springael D. 2016. Functional redundancy of linuron degradation in microbialcommunities in agricultural soil and biopurification systems. Appl EnvironMicrobiol 82:2843–2853. doi:10.1128/AEM.04018-15.

Editor: R. M. Kelly, North Carolina State University

Address correspondence to Dirk Springael, dirk.springael@ees.kuleuven.be.

Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.04018-15.

Copyright © 2016, American Society for Microbiology. All Rights Reserved.

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toward linuron degradation in the two strains involved divergentevolution and the independent acquisition of nonrelated isofunc-tional linuron hydrolytic gene functions by horizontal gene trans-fer (10). Moreover, in contrast to SRS16, WDL1 is a member of acommensal bacterial consortium in which most of the 3,4-DCAproduced from HylA-dependent linuron hydrolysis is channeledto 3,4-DCA catabolic strains (4).

The identification of HylA as an alternative linuron hydrolasein linuron-degrading Variovorax strains poses questions about itscontribution to environmental linuron degradation in agricul-tural environments in addition to that of LibA, i.e., about thefunctional redundancy of environmental linuron biodegradation.Catabolic enzymes for pesticide degradation are considered to bequite unique, as only a few enzyme types that perform the corre-sponding activity exist, making their genes excellent molecularmarkers for assessing the corresponding biodegradation capacityand activity. Several authors (8, 11, 12) reported on the use ofcatabolic molecular markers to assess herbicide biodegradation insoil. However, isofunctional diversity of enzymes in pesticide bio-degradation and the role of their functional redundancy remainlargely unexplored and are to date reported only for biodegrada-tion of phenoxy acid herbicides (11, 13–15).

In this study, we examined whether hylA-hosting organismscan successfully compete with libA-hosting organisms in linuron-exposed environments and whether hylA gene copy numbers ex-plain increases in linuron mineralization capacities in linuron-exposed environments where libA failed to do so. Moreover, weexamined whether commensalism in linuron metabolism as ob-served for hylA-carrying WDL1 and hence multispecies food websare involved in linuron biodegradation in the environment. Forthose purposes, gene copy numbers of hylA as well as dcaQ weredetermined in available DNA extracts from two different ecosys-tems for which the responses of the resident Variovorax commu-nity, of the libA gene copy number, and of the intrinsic capacity tomineralize linuron to the application of linuron were studiedpreviously (9, 16, 17). The gene dcaQ encodes the glutamine-ami-notransferase-like component of the multicomponent enzyme3,4-dioxygenase converting 3,4-DCA into chlorocatechol andfunctions as a marker for linuron biodegradation beyond 3,4-DCA (18, 19). The first ecosystem is an agricultural soil with ahistory of linuron treatment and was studied as a field experiment

followed by a lab-scale soil microcosm (SM) experiment that wasinitiated to verify results of the field experiment in a controlledway (9). The second ecosystem mimicked the environment of anon-farm BPS. Microcosms (BMs) containing BPS material weretreated with linuron (7, 16), and different stress events were im-posed to study their effects on linuron mineralization (20, 21).

MATERIALS AND METHODSqPCR. Primer3web and Primer-BLAST were used to design primers tar-geting hylA, encoding the linuron hydrolase HylA, and dcaQ, encodingthe glutamine-aminotransferase-like component of the multicomponentenzyme 3,4-dioxygenase. Primers for hylA were designed based on thesequences of the corresponding gene in the genomes of the linuron-de-grading Variovorax sp. strains WDL1 and PBS-H4. Primers for dcaQ weredesigned based on the corresponding gene sequences identified in Vario-vorax strains SRS16, WDL1, and PBS-H4 and in a range of chloroaniline-degrading bacteria, including the 3,4-DCA-degrading Comamonas testos-teroni strain WDL7, which is part of the linuron-mineralizing commensalbacterial consortium that includes WDL1 (Table 1). Phylogenetic analysisof dcaQ genes in chloroaniline-degrading bacteria showed the existence oftwo main groups that show around 80% identity at the nucleotide leveland 79% at the amino acid level. The groups were designated dcaQI anddcaQII, and primer sets that allowed discrimination between the twogroups were designed. Specificities of the primer sets were assessed in silicowith Primer-BLAST and a BLAST search and performance of PCR andquantitative real-time PCR (qPCR) on genomic DNA from several linu-ron-/3,4-DCA-degrading and non-linuron-/3,4-DCA-degrading Vario-vorax strains and bacteria of related and nonrelated genera. qPCR wasperformed in a Rotor Gene real-time centrifugal DNA amplification ap-paratus (Corbett Research, Australia). The real-time PCR mixtures con-tained 7.5 �l of Absolute QPCR SYBR Green mix (Thermo Fisher Scien-tific, United Kingdom), 0.30 �l of forward primer (200 nM), 0.30 �l ofreverse primer (200 nM), 3.90 �l of nuclease-free water, and 3 �l of10-fold-diluted template DNA. The exception was the hylA PCR mixture,which used a 100 nM solution of reverse primer HylA-RT-F instead of a200 nM solution. Reaction conditions were 15 min at 95°C, followed by 40cycles of 15 s at 94°C, 15 s at 60°C, and 15 s at 72°C. Standard curves forqPCR were compiled using 10-fold serial dilutions of amplicons (rangingfrom 1 copy/�l up to 108 copies/�l) of appropriate gene fragments gen-erated by conventional PCR from genomic DNA of strains SRS16 (dcaQI)and WDL1 (for hylA and dcaQII) as reported below. The fragments werepurified from agarose gels using the QIAquick gel extraction kit (Qiagen).DNA concentrations of the purified DNA fragments were determinedwith the NanoDrop 1000 spectrophotometer (Thermo Scientific). The

TABLE 1 Primer pairs used for either regular PCR or real-time PCR targeting linuron-specific catabolic genes

Target Primera Primer sequence Amplicon size (bp)GenBank sequenceaccession no.

hylA HylA-F AGGTCATGTCCACTCGCGTCT 1,905 KC146403; KC146406HylA-R GCCGATGCATAGGGCCATATTTGCTHylA-RT-F GCATGGGTCTGTTGCTGATAC 90HylA-RT-R CTGCGTGGAACTTCACTGTTAG

dcaQI dca I-F CTCTCATGGCCGGATCAATA 272 JN104632.1dca I-R TACAGATCGGCCAGCATCCAdca I-RT F AAGGGATTGAACACGAAGGC 137dca I-RT R TGGCCGGATCAATATGGTCTG

dcaQII dca II-F CGCCCACTGGTCATGTAAAG 377 KC146405.1dca II-R GAAAAGCACGGCATCTGGTCdca II-RT F GCCAAGACAACCGAACCATC 80dca II-RT R GGATACCCAGAAAGCCGCA

a Primers including RT in their designation were used for qPCR. The others were used for conventional PCR. F, forward; R, reverse.

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limit of detection for all genes by qPCR was 1.2 � 103 copies g (dry weight)of soil/BM material�1. For each DNA extract, qPCRs were performed induplicate. Gene abundances are expressed either as the gene copy numberof hylA/dcaQI/dcaQII per copy of bacterial 16S rRNA gene or as the per-centage of the bacterial 16S rRNA gene copy number. The values of bac-terial 16S rRNA gene copy numbers used originated from the work of Berset al. (9, 16, 17). hylA, dcaQI, and dcaQII gene copy numbers were deter-mined on the same DNA extracts that were previously used to determinelibA and 16S rRNA gene copy numbers (9, 16, 17).

Conventional PCR. Conventional PCR targeting hylA, dcaQI, anddcaQII was performed using the primer sets reported in Table 1. PCRmixtures contained 5 �l DreamTaq Green buffer (10 �) (Life Technolo-gies), 5 �l 1% bovine serum albumin (BSA), 4 �l 2.5 mM deoxynucleosidetriphosphates (dNTPs), 0.25 �l 0.1 mM forward and reverse primers, and0.25 �l DreamTaq polymerase (5 units/�l) (Life Technologies) adjustedto a total volume of 50 �l with nuclease-free water. PCRs were performedin a Biometra Thermocycler (AnalytikJena), and reaction conditions were15 min at 95°C, followed by 30 cycles of 1 min at 94°C, 1 min at 60°C, and1 min at 72°C and a final elongation step at 72°C for 10 min. Ampliconswere visualized by agarose gel electrophoresis (1% agarose, 75 min, 90 V)using GelRed (Biotium) as a nucleic acid stain.

Agricultural soil DNA extracts. The agricultural soil DNA extractsused originated from a field experiment and a concomitant SM experi-ment which assessed the responses of the Variovorax community, libAgene copy number, and linuron mineralization potential to linuron ap-plication. A detailed description of those experiments is found in the workof Bers et al. (9). Briefly, two adjacent potato field plots either were nottreated with any herbicide (plot P0) or, at day 0, were treated with 450 glinuron ha�1 (plot PF). Topsoil samples were taken from each plot atthree different positions (marked as A, B, and C in plot PF and D, E, andF in plot P0) at day 0, before the pesticide application, and at days 20 and34 after the treatment. At each position, three soil samples were taken in a

radius of 0.2 m. Soil samples were homogenized and used for DNA ex-traction for molecular analysis, dry weight measurement, and [14C]linu-ron mineralization assays in triplicate (9). After 176 days of pesticidetreatment, soil samples, taken from all six positions in plots P0 and PF,were mixed and six SMs consisting of glass columns (height, 10 cm; di-ameter, 4 cm) filled with the soil mixture were set up. All SMs were incu-bated at 25°C. Three SMs were irrigated with tap water with linuron, andthree were irrigated with linuron-free tap water (9) according to thescheme shown in Fig. 1 (9). For some SM replicates that originally re-ceived linuron, linuron application was intermittently stopped as picturedin Fig. 1. At selected time points, soil samples were taken for DNA extrac-tion as reported previously (9). Other DNA extracts used were those re-covered previously from linuron-mineralizing liquid enrichment culturesthat were started from samples of linuron-fed soil microcosms (L SM A, LSM B, and L SM C) as described previously (9).

DNAs from biopurification systems. The BPS DNA extracts usedoriginated from a BM experiment previously described by Sniegowski etal. (16, 20). A detailed description of the experiment is found in the workof Sniegowski et al. (16, 20). Briefly, the experiment made use of BMs setup in glass columns (height, 10 cm; diameter, 4 cm) filled with a mixtureof 25% (vol/vol) cut straw, 25% (vol/vol) peat, and 50% (vol/vol) soil. Thesoil was either a non-linuron-primed soil, C (BM type C), or a linuron-primed soil, L (BM type L). Soil L originated from the agricultural fieldstudied in the field experiment described above but was taken at anotherlocation within the field and 2 years before the field experiment. Soil C wasa subsurface soil obtained from a construction site. All setups were per-formed in triplicate. The setups were subjected to different treatments andperiods of stress (drought and freezing) as shown in Fig. 1 (16, 20). Sam-ples were taken as outlined in Fig. 1.

Data analysis. libA gene copy numbers, 16S rRNA gene copy num-bers, and linuron mineralization capacity data used in this study weretaken from previous reports (9, 16, 20, 21). The lag times determined in

FIG 1 Treatment schemes used in the microcosm experiments. In the agricultural soil microcosm (SM) experiment (top), as described by Bers et al. (9), SMs A,B, and C were treated with linuron with an intermittent period of no linuron application for SM A and SM B. SMs D, E, and F were never treated with linuron.A drought period without linuron application was imposed as indicated. In the BPS microcosm (BM) experiment (bottom) as described by Bers et al. (21) andSniegowski et al. (20), the BPS matrix contained either linuron-primed soil L or nonprimed soil C. The BMs were treated either with linuron (L�/C�) or withwater (L�/C�). Linuron treatment was stopped between week 12 and week 22 for BMs L� and C�, and an intermittent drought and cold period withoutlinuron application was imposed for all BMs.

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the linuron mineralization kinetics recorded in linuron mineralizationassays using samples taken from the field or SM and BM experiments asinoculum were used as a measure for the linuron mineralization capacity,i.e., the shorter the lag phase, the higher the mineralization capacity (9,16). The results were subjected to Student’s t test on a significance level of0.05 to assess differences between gene copy numbers and to assess effectsof linuron application and perturbations.

RESULTShylA and dcaQ dynamics in agricultural soil. The dynamics ofhylA gene abundances were similar to those previously reportedfor libA, but throughout the experiment, hylA gene copy numberswere at least 2 log10 units higher than those of libA in both thelinuron-treated and nontreated plots (Fig. 2). At day 20, as was thecase for libA, hylA abundances had significantly increased (40- to80-fold) at two of the three sampling positions in the linuron-treated plot PF compared to day 0 and were significantly higherthan those in the nontreated plot P0 (Fig. 2). Similar to libA genecopy numbers at day 34, hylA gene copy numbers had decreasedagain to an abundance similar to this at day 0. Overall, gene copynumbers of dcaQI and dcaQII followed the same dynamics as libAand hylA, with the highest number at day 20 in the linuron-treatedplots. However, dcaQI and dcaQII abundances were never in thesame order as those of libA and hylA. In particular, dcaQII showedhigh copy numbers that were 10 times higher than those of hylAand up to 1,000 times higher than those of libA at day 20 in thelinuron-treated plots. We conclude that, as for libA and hylA,dcaQI and dcaQII abundances were highest when the mineraliza-tion capacity in the linuron-treated plots was highest compared tothe nontreated plots, i.e., at day 20. At that time point, the lagphase of linuron mineralization as a measure for the mineraliza-tion capacity was reduced to approximately 5 days in the treatedplot compared to 8 days in the nontreated plot. Apparently, toobtain this reduction, the gene copy numbers of libA, hylA, dcaQI,and dcaQII have to reach (1.7 � 0.4) � 10�4%, (8.3 � 31) �10�2%, (1.8 � 40) � 10�3%, and (4.7 � 17.3) � 10�1% of thetotal number of 16S rRNA gene copies, respectively.

In the controlled SM experiment, hylA gene copy numberswere initially (before feeding with linuron) relatively high and, asin the field, exceeded libA gene copy numbers by a factor of 100 to1,000 despite the low linuron mineralization capacity (Table 2).These initial high hylA abundances were maintained only in mi-crocosms treated with linuron (Table 2), implying that the main-tenance of the hylA-containing population(s) depended on linu-ron application. At day 149, hylA gene copy numbers, however,also increased in the control microcosms, which can be attributedto the drought period between day 78 and day 110. A similarobservation was previously done for libA (Table 2) (9). In the caseof linuron application, hylA abundance increased further to2.82% � 0.72% of the bacterial 16S rRNA gene copies at day 491.The dependency of hylA gene copy numbers on linuron was fur-ther apparent from the drops in hylA abundance in replicate L SMC and replicate L SM B after stopping linuron application fromday 159 until day 285 and from day 285 until day 491, respectively(Table 2). Compared to hylA, both dcaQI and dcaQII abundanceswere a factor of 1,000 and 10 less at the start of the SM experiment,respectively (Fig. 3 and Table 2). dcaQ gene copy numbers andespecially those of dcaQII responded positively on linuron appli-cation but never coincided with hylA or libA abundances. As withlibA and hylA, both dcaQI and dcaQII increased in abundance in-

dependently of linuron application after the drought period be-tween days 78 and 110. Stopping linuron application in micro-cosm L SM B affected dcaQII abundance (Fig. 3 and Table 2).Overall, it can be concluded that gene copy numbers of most cat-abolic markers (except dcaQI) were highest when the mineraliza-tion capacity was highest. Lag times as low as 0.7 days were reachedin linuron-fed SMs which corresponded with libA, hylA, dcaQI,and dcaQII abundances of, respectively, (5.4 � 1.4) � 10�1%,(1.5 � 0.6) � 10�1%, (8.2 � 3.4) � 10�3%, and (5.1 � 2.2) �100% of the total number of 16S rRNA gene copies. In addition, aspreviously found for libA, a correlation was found between dcaQII

gene copy numbers and lag time (representing the mineralizationcapacity) (Fig. 4). This correlation was less obvious for hylA, par-ticularly due to the gene copy numbers recorded at day 0 and formicrocosm L SM B at day 491, where hylA gene copy numberswere fairly high but mineralization capacity was relatively low(Fig. 4).

In the work of Bers et al. (9), liquid enrichment cultures usinglinuron as the sole source of carbon and energy were obtainedfrom soil samples taken from plot PF 20 days after application oflinuron and from soil samples taken from the linuron-fed SMs onday 134 of incubation. In these enrichment cultures, libA abun-dance fell under the detection limit after a few transfers, althoughVariovorax was still present (9). Conventional PCR targeting hylAshowed that hylA instead of libA became the dominant linuronhydrolysis gene in those cultures upon prolonged enrichment (seeFig. S1 in the supplemental material).

hylA and dcaQ dynamics in biopurification systems. In BMscontaining primed soil L, overall hylA behaved similarly to libA(Fig. 5, top), although hylA abundances were generally higherthan those of libA (2 to 10 times higher). Similarly to libA, hylAclearly responded to the linuron feed with increasing gene copynumbers in the treated microcosms and a decrease of gene copynumbers when linuron feeding stopped. A cold period withoutlinuron application did not affect hylA gene copy numbers. Whilethe drought period without linuron application affected the abun-dance of neither libA nor hylA, consequent rewetting and resum-ing the feed with linuron after the drought period resulted in adramatic increase of hylA gene copy numbers to (2.3 � 0.3) �10�4% of the 16S rRNA gene copy number. In the non-linuron-treated BMs containing primed soil, drought-wetting events af-fected both libA and hylA gene copy numbers positively. Theabundance of dcaQI followed an increasing trend similar to that oflibA in the linuron-treated BMs, although the initial increase be-tween week 0 and week 2 was 10-fold higher than for libA. dcaQII

gene copy numbers remained similar to those of hylA and fol-lowed a similar trend but diverged toward the end to a 10-folddifference (dcaQII � hylA) where dcaQII became the dominantdcaQ variant. In the BMs containing primed soil, the highest linu-ron mineralization capacity (lowest lag phase, 0.8 � 0.1 days) wasreached in the linuron-treated BMs at week 55, corresponding tothe highest percentage of dcaQII ([2.9 � 0.3] � 10�2%), dcaQI

([1.4 � 0.3] � 10�3%), hylA ([2.3 � 0.2] � 10�3%), and libA([5.0 � 2.7] � 10�5%) of the 16S rRNA gene copy number.

In BMs containing nonprimed soil C, libA was previously re-corded in gene copy numbers above the detection limit in only onereplicate BM, i.e., BM3 of setup C�, although similar high linuronmineralization capacities developed in the other two replicates(9). In contrast to libA, hylA was detected at most time pointsexcept when the mineralization capacity was very low (for in-

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stance, week 17). However, even when the mineralization capacitywas relatively high (short lag time), for instance, at week 55, hylAabundances were extremely low (down to [1.4 � 1.6] � 10�7%),even in replicates where libA gene copy numbers were below the

detection limit (Fig. 5, bottom). Despite the low abundances oflibA or hylA gene copies and the low linuron mineralization ca-pacity, both dcaQ genes increased in the first 17 weeks of linurontreatment to reach and remain at a high abundance until the end

FIG 2 Abundances of libA, hylA, dcaQI, and dcaQII in samples taken at days 0, 20, and 34 from positions A, B, and C in the linuron-treated field plot PF andpositions D, E, and F in the nontreated field plot P0. Reported values are the log10 values of the average hylA gene copy numbers expressed as a percentage of thebacterial 16S rRNA gene copy number with standard deviation (n � 6; 3 soil samples per position, 2 qPCRs per sample) (approximately 109 bacterial 16S rRNAgene copies/g soil). Student’s t test was used to determine significant differences in treatment and time points (P 0.05). An asterisk above a bar marks asignificant difference in gene abundance at a specific position (n � 6) in a linuron-treated plot compared to the average abundance of that gene determined atpositions D, E, and F of the nontreated field plot P0 at the same time point. To indicate whether gene abundance is significantly different between different timepoints at a specific position, “a,” “b,” and “c” are used as markers above the bars of each position. “a” marks a significant increase between day 0 and day 20, “b”marks a significant decrease between day 20 and day 34, and “c” marks a significant increase between day 0 and day 34. Lag time (bottom) as a measure for thelinuron mineralization capacity is shown for each position of plots PF and P0 on days 0, 20, and 34. Values are averages (n � 3, 3 soil samples per position) withstandard deviations shown as error bars. Values for libA abundances and mineralization capacity (lag time) were taken from the work of Bers et al. (9).

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of the linuron treatment. Nevertheless, the linuron mineralizationlag time was at a minimum when the copy numbers of dcaQ geneswere at a maximum, i.e., at week 55. No linuron mineralizationoccurred in the samples taken from water-treated BMs withnonprimed soil. libA was not detected in any of these samples,while hylA and both dcaQ genes were detected at low copy num-bers (approximately 10�7% of the 16S rRNA gene copy number).We conclude that in the BMs containing nonprimed soil, thehighest linuron mineralization capacity (lowest lag time, 1.3 � 0.3days) was reached in the linuron-treated BMs at day 55 corre-sponding to the highest percentage of dcaQII ([1.06 � 1.06] �10�4%), dcaQI ([5.6 � 1.2] � 10�4%) of the 16S rRNA gene copynumber but not of hylA ([1.4 � 1.6] � 10�7%), and libA (3.3 �10�5% in only one replicate, BM3).

DISCUSSIONlibA, hylA, and dcaQ dynamics in agricultural soil. Our resultsshow that hylA gene copy numbers, like libA gene copy numbers,clearly depended on and responded to the application of linuronin agricultural soil with a history of linuron application. BothhylA- and libA-containing bacteria were previously isolated fromthat soil and, hence, known to be endogenous to the studied field(22). We conclude that as a response to linuron application, bothhylA- and libA-carrying bacteria grow in the soil, indicating thatbacteria carrying hylA compete successfully with those carryinglibA in the same soil. Neither of the two is really outcompeted,indicating that the two benefit simultaneously from the appliedlinuron to grow and/or maintain their population size. It alsoindicates that hylA-carrying hosts, in addition to libA-carryinghosts, contribute to linuron degradation in the soil. This is incontrast to studies that follow the dynamics of alternative tfdAgenes that encode the enzyme that initiates the metabolism ofphenoxy acid herbicides in soils as a response to treatment withdifferent types of phenoxy acid herbicides. It was found that de-pending on the phenoxy acid substrate, specific tfdA gene groups

became dominant, despite the initial presence of multiple groups(11, 13, 23), implying that in the case of phenoxy acid herbicidesthe initial assessment of functional genes in soils does not neces-sarily reflect the organisms or genes that proliferate and performthe degradation of the compounds in question, which is the case inour study.

The high initial abundance of hylA in the agricultural soil at thestart of the field experiment might be a result of growth of hylA-containing bacteria on residual linuron from previous applica-tions in the field and a higher persistence of those populationsthan of the libA-carrying populations under field conditions. Sur-prisingly, in contrast to libA, relatively high gene copy numbers ofhylA in the agricultural soil did not always correspond to a highlinuron mineralization capacity (9) (Fig. 4 and Table 2, day 0 andday 491 for microcosm L SM B after linuron application wasstopped), indicating that hylA gene copy numbers do not alwayscontribute to the measured mineralization capacity. This is ex-plained by the abundances of dcaQ involved in the downstreammetabolism of linuron and hence actual 14C-labeled CO2 produc-tion. In contrast to other time points, for which a high mineral-ization capacity was recorded, dcaQ, and more precisely dcaQII,showed relatively low gene copy numbers at day 0 and, for micro-cosm L SM B, at day 491. The high gene copy numbers of hylA-containing bacteria and low gene copy numbers of dcaQ-contain-ing bacteria indicate that at those time points, linuron wasconverted to 3,4-DCA but only slowly mineralized beyond 3,4-DCA and, as such, explain the apparent incongruence betweenhigh hylA abundances and low linuron mineralization capacity.These data have other important implications since they show thathylA-containing bacteria do not always contain the downstreampathway and as such must compose a part of consortia, including3,4-DCA-mineralizing organisms that do not convert linuroninto 3,4-DCA. At other time points, though, dcaQ gene copy num-bers often exceeded hylA gene copy numbers, which implies that

FIG 3 Abundances of catabolic genes encoding linuron mineralization in the water-treated (left) and linuron-treated (right) SM setups of the agricultural soilexperiment. Linuron hydrolase genes libA (�) and hylA (Œ) and genes dcaQI (Œ) and dcaQII (o) are expressed as percentages of the bacterial 16S rRNA gene copynumber (approximately 109 bacterial 16S rRNA gene copies/g soil). Reported values are average values with the standard deviation (lag time, n � 3 [3 replicates];qPCR, n � 6 [3 replicate SMs, 2 qPCRs]) indicated by the error bars. Values for libA abundances were taken from the work of Bers et al. (9).

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(i) each dcaQ-containing cell might contain an hylA copy but alsothat (ii) populations that contained only the 3,4-DCA catabolicpathway profited from organisms that perform HylA/LibA activ-ity. The complex dynamics of the examined catabolic genes sug-gest a high plasticity of the different catabolic gene modules in-volved in linuron degradation in the agricultural soil as a responseto linuron application potentially involving horizontal gene trans-fer, as previously suggested by Dunon et al. (24). Interestingly,hylA was identified in Variovorax strains that are part of consortiain which efficient linuron conversion to 3,4-DCA and the furthermineralization of 3,4-DCA depend on synergistic metabolic inter-actions between Variovorax and other bacteria. In the consortiumreported by Dejonghe et al. (4), Variovorax sp. strain WDL1,which carries hylA, converts linuron to 3,4-DCA. Strain WDL1can grow on 3,4-DCA, but the conversion of 3,4-DCA is not thatefficient, leading to a release of 3,4-DCA that is used for growth bya second consortium member, Comamonas testosteroni WDL7.Concomitant removal of 3,4-DCA in the culture in turn results inan improved rate of conversion of linuron by strain WDL1. Inanother similarly composed consortium that depends on hylA forinitial linuron hydrolysis, a similar cooperation exists (3). IncP-1plasmids are part of these consortia and, for instance, carry thegenes for 3,4-DCA biodegradation in strain WDL7 (19). In con-

trast to hylA, libA was primarily found in Variovorax strains, suchas strain SRS16 (6), which show efficient conversion of 3,4-DCAand hence growth on linuron as single strains. libA gene copynumbers were always below dcaQ gene copy numbers, and hence,each libA-containing cell might contain a dcaQ gene copy. Inter-estingly, in liquid cultures that were enriched for organisms thatuse linuron as sole carbon source and were initiated from thestudied agricultural soil, the coexistence of libA- and hylA-con-taining bacteria is lost and hylA-containing consortia start domi-nating. Apparently, libA-containing bacteria are less competitivethan hylA-containing strains under those culture conditions.

libA, hylA, and dcaQ dynamics in BPS. The results show that,as in the agricultural soil, in the BPS matrix containing primedsoil, hylA-containing bacteria compete successfully with libA-containing bacteria for linuron as a carbon source and growalongside. hylA gene copy numbers did not respond on linuronapplication in BMs containing nonprimed soil, indicating thathylA-carrying microorganisms likely originate from soil L, whichwas included in the BM matrix, indicating a successful invasion ofhylA linuron-degrading bacteria from that primed soil in the over-all system. The same was concluded for libA (21). As with libA,hylA abundance appears to correlate with the observed dynamicsof the linuron mineralization capacity in the system containing

FIG 4 Correlation between lag time of linuron mineralization and abundance of the linuron hydrolase (left) (libA and hylA) and dcaQ (right) (dcaQI and dcaQII)genes as log10 value of the percentage of the bacterial 16S rRNA gene copy number for the water-fed (open symbols) and linuron-fed (solid symbols) SMscontaining agricultural soil. Reported values are average values with the standard deviation (lag time, n � 3 [3 replicates]; qPCR, n � 6 [3 replicate SMs, 2qPCRs]) shown by the error bars. Linear regressions are shown as dashed lines. R2 values are shown. Linuron mineralization lag times and libA gene copy numberswere taken from the work of Bers et al. (9).

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primed soil, i.e., hylA gene copy numbers were relatively high andlow when the linuron mineralization capacity was high and low,respectively (Fig. 6). Interestingly, between week 51 and week 55,libA gene copy numbers remained the same or even tended todecrease in linuron-amended BMs while the linuron mineraliza-tion capacity significantly increased. In contrast, hylA gene copynumbers were extremely high and had increased 8-fold from week51 onward. This occurred after the drought-rewetting period im-plemented between weeks 42 and 51. A similar observation wasdone for hylA in the nontreated BMs containing primed soil, in-dicating that this stress situation benefitted hylA-carrying popu-lations in BPS. In contrast, libA-carrying bacteria seem to benefitfirst from the drought stress in the non-linuron-treated micro-cosms. In contrast to the agricultural soil, dcaQ gene copy num-bers always exceeded hylA/libA gene copy numbers, implying thateach hylA/libA-containing cell might also contain a dcaQ genecopy. However, populations that contained only dcaQ clearly ben-efited from linuron application, suggesting that as in the agricul-tural soil, 3,4-DCA-degrading organisms that contain only thepathway beyond 3,4-DCA profited from LibA/HylA activity per-formed by other organisms.

In the linuron-fed BM setup that contained non-linuron-primed soil, some response of hylA gene copy numbers was re-corded upon addition of linuron, but this population seems todeteriorate over time despite the maintenance of a high linuronmineralization capacity. Previously, libA was recorded to respondto linuron application in only one of the replicates of that SMsetup, and these data also did not always explain the observed highmineralization capacity. As such, we conclude that still other linu-ron hydrolases or biochemical systems to initiate linuron miner-alization exist in that environment, likely originating from thenonprimed soil C. PuhA (25) and PuhB (26) are other enzymesisofunctional to LibA and HylA, involved in linuron hydrolysis byGram-positive bacteria, but they have never been linked with min-eralization of linuron. They might form alternatives for initiat-ing linuron mineralization in the environment carrying thatcapacity, potentially in accordance with dcaQ-containing pop-ulations that clearly proliferated upon linuron addition in thelinuron-amended SMs containing nonprimed soil.

Conclusions. Our results provide further insight into the mi-crobial ecology of linuron biodegradation in agricultural environ-ments. Both organisms containing libA and hylA coexist and con-

FIG 5 Dynamics of libA, hylA, dcaQI, and dcaQII abundances and of the linuron mineralization lag phase (as a measure for linuron mineralization capacity) inBMs inoculated with linuron-primed soil (top) (approximately 109 to 1010 bacterial 16S rRNA gene copies/g soil) and non-linuron-primed soil (bottom)(approximately 108 to 109 bacterial 16S rRNA gene copies/g soil) treated either with water (left) or with linuron (right). Abundances of the hydrolase genes (Œ)(libA [solid line]; hylA [dashed line]) and dcaQ genes (o) (dcaQI [solid line]; dcaQII [dashed line]) are expressed as log10 values of the average ratio of therespective gene copy number to the bacterial 16S rRNA gene copy number (n � 6, 3 replicates; 2 qPCRs for each replicate) with the standard deviations shownby the error bars. Values of libA for the linuron-treated BM containing nonprimed soil are from only one replicate BM (BM3) since libA was not detected for theother two BMs. Values of libA abundance were taken from the work of Bers et al. (21). Values below the x axis are below the detection limit. Values of the lag time(days) recorded in linuron mineralization assays as a measure for the linuron degradation capacity were taken from the work of Bers et al. (21). Gray bars areaverage values with standard deviations (n � 3; 3 replicate microcosms) for the water- and the linuron-treated BM setups.

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tribute to linuron biodegradation. However, situations occur inwhich one is privileged over the other as observed under liquidenrichment conditions in minimal medium and in the BM exper-iment after the drought-wetting period. This might be related tothe particular bacterial host which carries the genes or otherwiseto the enzymatic kinetic parameters of the corresponding en-zymes. LibA is known to have a 2-fold-higher affinity than HylAfor linuron. Due to a lack of data on the maximal specific conver-sion rates for LibA, it is difficult, however, to draw conclusionsfrom the role of the linuron hydrolysis kinetic parameters for us-age of linuron as a growth substrate. Our study further shows thatboth hydrolysis genes and downstream catabolic genes should beused for assessing linuron mineralization capacity in environmen-tal samples. However, other (unknown) enzymes initiating linu-ron mineralization apparently exist in the environment. Further-more, our data strongly suggest that consortia involving bothlinuron-hydrolyzing organisms and organisms that further de-grade the produced 3,4-DCA cooperate in complete linuron min-eralization in the examined ecosystems. Moreover, complex dy-namics and interactions of hydrolysis genes and downstreamcatabolic genes exist in the examined ecosystems, potentially in-volving horizontal gene transfer. This study proves the presenceand activity of pesticide degraders functioning as synergistic con-sortia in natural environments.

ACKNOWLEDGMENTS

Many thanks go to K. Simoens and D. Grauwels for real-time qPCR anal-ysis.

FUNDING INFORMATIONThis work, including the efforts of Dirk Springael, was funded by KULeuven (Katholieke Universiteit Leuven) (OT10/03). This work, includ-ing the efforts of Benjamin Horemans, was funded by Fonds Wetenschap-pelijk Onderzoek (FWO) (12Q0215N). This work, including the efforts ofDirk Springael, was funded by Fonds Wetenschappelijk Onderzoek(FWO) (G.0371.06). This work, including the efforts of Karolien Bers, wasfunded by Agentschap voor Innovatie door Wetenschap en Technologie(IWT) (SB/73381). This work, including the efforts of Dirk Springael, wasfunded by Federaal Wetenschapsbeleid (BELSPO) (P7/25).

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FIG 6 Correlation between the lag time of linuron mineralization and the abundance of libA, hylA, dcaQI, and dcaQII as the log10 value of the percentage of thebacterial 16S rRNA gene copy number in BM setups with linuron-primed soil treated with linuron (black) and water (white) of the BPS experiment. Values areaverage values (lag time [n � 3]; qPCR, n � 6). Linear regressions for libA and dcaQII and exponential regressions for hylA and dcaQII are shown as a dotted line.R2 values and equation parameters are shown on the right. Symbols related to day 0 and day 491 are indicated.

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