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Keywords:

  • biopurification system;
  • pesticide-primed soil;
  • bioaugmentation;
  • linuron mineralization;
  • Variovorax

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Microcosms were used to examine whether pesticide-primed soils could be preferentially used over nonprimed soils for bioaugmentation of on-farm biopurification systems (BPS) to improve pesticide mineralization. Microcosms containing a mixture of peat, straw and either linuron-primed soil or nonprimed soil were irrigated with clean or linuron-contaminated water. The lag time of linuron mineralization, recorded for microcosm samples, was indicative of the dynamics of the linuron-mineralizing biomass in the system. Bioaugmentation with linuron-primed soil immediately resulted in the establishment of a linuron-mineralizing capacity, which increased in size when fed with the pesticide. Also, microcosms containing nonprimed soil developed a linuron-mineralizing population, but after extended linuron feeding. Additional experiments showed that linuron-mineralization only developed with some nonprimed soils. Concomitant with the increase in linuron degradation capacity, targeted PCR-denaturing gradient gel electrophoresis showed the proliferation of a Variovorax phylotype related to the linuron-degrading Variovorax sp. SRS16 in microcosms containing linuron-primed soil, suggesting the involvement of Variovorax in linuron degradation. The correlation between the appearance of specific Variovorax phylotypes and linuron mineralization capacity was less clear in microcosms containing nonprimed soil. The data indicate that supplementation of pesticide-primed soil results in the establishment of pesticide-mineralizing populations in a BPS matrix with more certainty and more rapidly than the addition of nonprimed soil.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Since 1940, pesticides are intensively used worldwide. An important environmental issue of pesticide use is the pollution of ground and surface water as a result of either diffuse (run-off, percolation and spray drift) or point contamination (direct losses through spillage and leakages). Recent studies showed that direct losses account for 40–90% of the surface water pollution (De Wilde et al., 2007). To minimize direct pesticide losses, the installation of biopurification systems (BPS) to treat pesticide-contaminated wastewater on the farm yard has been proposed (Torstensson & del Pilar Castillo, 1997; Vischetti et al., 2004; De Wilde et al., 2007). In on-farm BPS, the contaminated water is conducted over a solid matrix, called a biomix, which is composed of a mixture of various materials, for example straw, peat and soil, in which biodegradation and sorption result in pesticide removal. BPS are considered a simple, low-cost, practical and labor-extensive approach for farmers to treat pesticide-contaminated wastewater on the farm. Despite the high pesticide removal percentage observed in BPS (Fogg et al., 2003a, b, 2004; Pigeon et al., 2005), degradation remains poor for some pesticides (Fogg et al., 2003a, 2004). Moreover, a rapid complete degradation is advised to avoid possible toxicity effects of accumulated contaminants (Henriksen et al., 2003), aging (Johannesen et al., 2003; Zhao et al., 2003) and the occurrence of mobile and toxic metabolites (Coppola et al., 2007). In addition, degradation during start-up of the system is poor (Fogg et al., 2004) because the appropriate microorganisms need to proliferate in the biomix before maximum degradation rates are obtained. To ensure rapid and complete degradation of the pesticides in BPS, bioaugmentation of the biomix with microorganisms containing catabolic pathways enabling complete mineralization of pesticides is suggested. Pesticide-mineralizing microorganisms can often be found in soils or other ecosystems with a long history of pesticide contamination, designated as pesticide-primed materials. The organisms can be inoculated either as (formulated) cultured strains or along with the pesticide-primed material (e.g. soil) in which they have developed. In comparison with pure cultures, the latter is expected to contain (1) a larger gene pool and higher diversity in microorganisms that contribute to pesticide degradation and (2) populations better adapted to in situ conditions. In addition, this approach does not require the isolation of the appropriate organisms. This is important since, to date, pure strains able to mineralize a pesticide have been reported only for a few pesticide compounds. Moreover, bioaugmentation using lab-cultured pollutant-degrading isolates often had limited success (Chatterjee et al., 1982; Grigg et al., 1997; Struthers et al., 1998; Topp, 2001; Mertens et al., 2006; Moran et al., 2006; Singh et al., 2006; Bazot & Lebeau, 2008). On the other hand, the few reports on bioaugmentation of contaminated matrices with pollutant-primed materials showed promising results (Barbeau et al., 1997; Runes et al., 2001; Grundmann et al., 2007). Despite the apparent high potential of applying primed materials for bioaugmentation of contaminated ecosystems, little research has been performed on this topic. Moreover, no reports exist of using pesticide-primed materials for bioaugmenting BPS despite the low associated cost of such an approach for farmers. However, BPS contain a complex matrix prepared of components from different ecosystems harboring different microbial communities and provide a high nutrient content with multiple substrates for microbial growth. It has to be examined whether pesticide-mineralizing populations introduced by inoculating pesticide-primed materials can compete and proliferate in such a complex biotechnological matrix.

Therefore, in this study, the dynamics of the pesticide mineralization capacity of lab-scale BPS microcosms (BM), containing a mixture of soil, straw and peat, were compared when inoculated with either a pesticide-primed soil possessing a pesticide mineralization capacity or a nonprimed soil without apparent pesticide mineralization capacity and irrigated or not with a pesticide-containing solution. The phenylurea herbicide linuron was used as model pesticide and soil originating from an agricultural field that had been treated annually with linuron and that contains linuron-mineralizing organisms (Breugelmans et al., 2007) was used as the model pesticide-primed material. The linuron mineralization capacity of the BMs was monitored by means of 14C-linuron mineralization assays. Because linuron-degrading isolates originating from linuron-treated soils, including the linuron-primed soil used in this study, almost exclusively belong to the genus Variovorax (Dejonghe et al., 2003; Sorensen et al., 2005; Breugelmans et al., 2007), the number of Variovorax and composition of the Variovorax community within the BMs was monitored by means of targeted molecular techniques.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Pesticides used

Linuron [3-(3,4-dichlorophenyl)-1-methoxy-1-methyl urea] (purity, 99.5%) was purchased from Sigma Aldrich (Belgium). [phenyl-U-14C] linuron (16.93 mCi mmol−1, radiochemical purity >95%) was obtained from Izotop, Hungary.

BM set-ups

BM were set up in glass cylinders (height 10 cm; diameter 4 cm) containing a glass filter positioned at 8 cm depth and filled with the appropriate mixture of soil, peat and straw (Table 1). The linuron-primed soil (soil L) was sampled from the A-horizon of a potato field in Halen, Belgium, in April 2005. The field had been treated for several years with linuron and was shown to contain a linuron-mineralizing microbial community (Breugelmans et al., 2007). Non-linuron-primed soils were obtained from seven different locations representing four different ecosystems, i.e. garden (soils G1 and G2), agriculture field (soils A1 and A2), forest (soils F1 and F2) and a construction site (soil C). To the best of our knowledge, these soils were never treated with linuron and, in addition, did not show any capacity to mineralize linuron for a period of 70 days in 14C-linuron mineralization assays (data not shown). All soils were top soils sampled from the upper 20 cm. The soils were stored at 4 °C in the dark and sieved before use. After filling, the BMs were positioned on a dish to collect drainage water and placed in a glass jar closed with an air-open lid to avoid desiccation and contamination. No leaching of linuron was observed.

Table 1.   Overview of the different BM set-ups operated in this study
Set-up*Origin soilLinuron- primed soilLinuron treatmentMoisture content (w/w%)pH (± SD)
  • *

    + and − indicate treatment with and without linuron, respectively.

  • Straw: 50 vol%, peat: 50 vol%, soil: 0 vol%.

Experiment 1: mixture: straw (25 vol%); peat (25 vol%); soil (50 vol%)
LAgriculture+56.126.20 (± 0.12)
L+Agriculture++60.526.20 (± 0.12)
CConstruction site48.006.05 (± 0.22)
C+Construction site+48.146.05 (± 0.22)
Experiment 2: mixture: straw (37.5 vol%); peat (37.5 vol%); soil (25 vol%)
O+No soil+217.785.11 (± 0.10)
L+Agriculture++89.525.44 (± 0.26)
C+Construction site+71.035.49 (± 0.21)
A1+Agriculture+42.675.56 (± 0.55)
A2+Agriculture+34.645.09 (± 0.15)
F1+Forest+82.335.61 (± 0.10)
F2+Forest+57.544.87 (± 0.11)
G1+Garden+74.195.43 (± 0.05)
G2+Garden+42.255.38 (± 0.30)

Two BM experiments were set up as outlined in Table 1, which provides an overview of the compositions, moisture contents and pHs of the biomix in the various BM set-ups. The moisture content, based on the weight of a sample (± 0.500 g) taken from the upper layer of the BMs before and after 2 days incubation at 60 °C, was determined at each sampling time. The pH was measured at the start and the end of both experiments in a 0.01 M CaCl2 extract (soil : liquid ratio 1 : 5). The pHs determined at the start of the experiments did not differ significantly within each set-up. In addition, neither pH nor moisture content changed significantly during the incubation period (data not shown).

In the first experiment, BMs contained a mixture of 25 vol% cut straw (± 0.5 cm2), 25 vol% peat and 50 vol% soil. The physico-chemical characteristics of the used substrata are shown in Table 2. BMs of set-ups L and L+ were inoculated with the linuron-primed soil L and packed to a density of 0.94 g cm−3, while BMs of set-ups C and C+ were inoculated with nonprimed soil C packed to a density of 0.78 g cm−3. BMs of set-ups C and L were irrigated with sterile tap water, while BMs of set-ups C+ and L+ received sterile tap water containing 60 mg L−1 linuron. Both solutions were manually spread evenly over the surface of the matrix using a 1-mL pipette during 12 weeks. Each week, the solutions were applied on Monday (1 mL), Wednesday (1 mL) and Friday (1.5 mL), resulting in an average added volume of 3.18 L m−3 day−1. Each set-up included triplicate BMs. All BMs were incubated in the dark at 25 °C. The upper 1 cm of the matrix in the BMs was mixed with a sterile spatula, before taking samples to examine the linuron mineralization capacity. The samples were taken exactly 0, 2, 4, 8 and 12 weeks after starting the treatments. Additional samples were taken at weeks 0, 2 and 12 for analysis of the Variovorax community.

Table 2.   Physicochemical characteristics of the substrata used in the biomix of the BMs
SubstratumMoisture content % (w/w)pHSpecific density (g cm−3)Total C (%)Total N (%)C/N (%)
  1. ND, not determined.

Soil C2.836.92.810.950.0812.1
Soil L11.534.9ND0.800.0613.3
Straw8.406.61.5643.250.4695.1
Peat49.676.41.5843.970.9347.1

In the second experiment, BMs contained eight different mixtures consisting of 37.5 vol% cut straw, 37.5 vol% peat and 25 vol% of soil and were operated in triplicate. The straw and peat used were the same as those used in the first experiment. BMs of set-up L+ were inoculated with the linuron-primed soil, while BMs of the other set-ups were inoculated with either one of the seven nonprimed soils (G1, G2, A1, A2, F1, F2, and C). As a control, a ninth set-up was included in which the BMs contained a mixture of 50 vol% straw and 50 vol% peat only. Each of these BM set-ups received a linuron-containing solution for a period of 20 weeks as described above. Samples were taken after 0, 15 and 20 weeks of incubation at 25 °C for determination of the 14C-linuron mineralization capacity as described for the first set-up. Additional samples were taken at weeks 0 and 15 for analysis of the Variovorax community.

14 C-linuron mineralization assays

14C-linuron mineralization assays were performed as described by Breugelmans et al. (2007). The matrix sample (200 mg) was added to a volume of 5 mL MMN minimal medium (Breugelmans et al., 2007), pH 5.8, containing both unlabeled (20 mg L−1) and 14C-labeled (31 μg L−1) linuron as the only carbon and nitrogen source (a final radioactivity of 213 Bq mL−1) in 20-mL pyrex tubes containing NaOH traps. During incubation at 20 °C on a rotary shaker at 150 r.p.m., the amount of 14CO2 produced with reference to the initial added amount of 14C-linuron was measured and cumulative mineralization curves were established. As a negative control, 14CO2 production of the medium without inoculation of a biomix sample was monitored. The lag phase, defined as the period between initiating the mineralization assay and start of mineralization, was calculated as the intersection of the x-axis with the linear regression line between two successive points of the mineralization curve, where the amount of 14CO2 showed the largest increase. The slope of the linear regression line of maximum mineralization determined the maximum mineralization rate (Broos et al., 2005).

Most probable number (MPN) mineralization method

An estimation of the size of the active linuron-mineralizing biomass in samples of the BM matrix was performed using an MPN approach. A sample (10 g, wet weight) taken from the biomix of the BM was added to 25 mL MgSO4 (10−2 M) and incubated overnight on a shaker to remove the cells from the matrix. After settling of the matrix material for 2 h, a decimal serial dilution was made with 2 mL of the aqueous extract. Aliquots of 0.5 mL of these dilutions were used as inoculum for 14C-linuron mineralization assays in triplicate, operated as described above. Positive tubes for MPN calculation were those vials where >10% total 14CO2 was produced within 60 days. MPN calculations were performed using the computer-assisted method developed by Briones & Reichardt (1999) and the MPN number was expressed as active mineralizing units per gram dry weight (dw) of the sample (AMU g−1 dw).

Molecular techniques

Total DNA was extracted from 400-mg samples of the biomix as described by Uyttebroek et al. (2006). The copy number of the Variovorax 16S rRNA gene was determined by real-time PCR, performed in a Rotor Gene (RG 3000) apparatus from Westburg using the Variovorax-specific 16S rRNA gene primers VarF and VarR. Details of the method are described in Bers et al. (2011). Real-time PCR determined the bacterial 16S rRNA gene copy number as described by Haest et al. (2010) using primers Eub341F and Eub534R (Muyzer et al., 1993). The detection limit of both methods was 105 copies g−1 biomix dw.

Denaturing gradient gel electrophoresis (DGGE) fingerprints of the Variovorax community were performed using 16S rRNA gene fragments amplified with primers VarF-GC and VarR in either a single or a double PCR as described by Bers et al. (2011) on a polyacrylamide gel (10%) with a denaturating gradient from 45% to 75% as described by Uyttebroek et al. (2006). The 16S rRNA gene fragments in the gel were mobilized through an electric field set at 120 V for 15 h. On each gel, a reference DGGE migration Variovorax marker sample (ref-Var) was loaded, containing 16S rRNA gene fragments PCR amplified with primers GC-VarF and VarR from Variovorax sp. SRS16 (Sorensen et al., 2005), Variovorax sp. DSM66 and Variovorax sp. WDL1 (Dejonghe et al., 2003) representing three major phylotypes within the Variovorax genus (Breugelmans et al., 2007). In addition, reference DGGE migration marker samples (ref-nonVar) containing 16S rRNA gene fragments with different GC content from various bacteria (not belonging to the Variovorax genus) amplified with bacterial primers 63F and 518R (El Fantroussi et al., 1999) were loaded on each gel. The detection limit of the Variovorax-specific PCR-DGGE method was approximately 105 copies g−1 biomix dw. The significance of differences between averages of the 16S rRNA gene copy numbers of triplicate BM samples were analyzed by anova (P<0.05).

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

Linuron mineralization capacity of BM bioaugmented with linuron-primed soil and nonprimed soil

In the first experiment, bioaugmentation of BPS with linuron-primed soil L and nonprimed soil C was compared. Initially, only samples from BMs inoculated with linuron-primed soil L, i.e. BMs of set-ups L and L+, showed linuron mineralization with a lag time of approximately 9.9 ± 0.4 days and a maximum mineralization rate of 14.2 ± 1.5% day−1. After 2 weeks of linuron treatment, the lag time of linuron mineralization recorded for samples of BMs of set-up L+ was reduced to 2.6 ± 1.4 days (Fig. 1). At week 12, the lag time further decreased to 0.46 ± 0.56 days. MPN counting showed that the linuron-mineralizing population in BMs of set up L+ increased from 4.5 × 102 to 1.8 × 104 AMU g−1 during the 12 weeks of linuron treatment. The lag time of the mineralization curves performed with the dilution series increased with higher dilution. Therefore, it was concluded that the reduction of the lag time observed in the BMs of set-up L+ corresponded to an increase in the linuron-mineralizing biomass (Sniegowski et al., 2009). With the samples taken from BMs of set-up L, the lag time initially decreased to 6.1 ± 0.1 days at week 2, but again increased to 8.9 ± 0.9 days at week 12.

image

Figure 1.  Linuron mineralization kinetics recorded for samples taken from the BMs of (a) set-up L+ and (b) set-up L in the first experiment. BMs of set-up L+ were continuously irrigated with tap water containing linuron, while BMs of set-up L received noncontaminated water. The arrow indicates the changes in lag time. The data are average values with indicated SDs of samples taken from three replicate BMs.

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BMs from set-up C+ and set-up C did not show a linuron-mineralizing capacity initially. However, BMs from set-up C+ clearly acquired the capacity to mineralize linuron upon continuous irrigation with linuron. This capacity developed in the three replicate BMs, after different periods of linuron feeding (Fig. 2). Moreover, for one replicate BM, a significantly lower mineralization rate than that for the other two was recorded. In addition, the mineralization curve recorded with this replicate demonstrated a rather linear mineralization curve, indicating that mineralization was not linked to growth. At week 12, the average recorded lag time was 4.5 ± 1.8 days. BMs from set-up C did not develop an observable linuron mineralization capacity during the 12-week incubation period (data not shown).

image

Figure 2.  Evolution of the linuron mineralization kinetics recorded with samples taken from the three replicate BMs designated as BM1, BM2 and BM3 of set-up C+ inoculated with the non-linuron-primed soil in the first experiment.

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Adaptability of BPS bioaugmented with nonprimed soils

Because BMs of set-up C+ developed a linuron mineralization capacity upon the continuous supply of a linuron-containing solution, it was tested whether inoculation with other nonprimed soils led to a similar development. Therefore, a second experiment was performed in which nonprimed soils of different origins were used to bioaugment BMs. To ensure that the acquisition of a linuron mineralization capacity was related to the inoculation of the soil, a set-up containing control BMs without soil addition was included. BMs containing linuron-primed soil L and nonprimed soil C were included in the experimental set-up. All systems were irrigated with the linuron-containing solution. Because in the first experiment, the development of a linuron-mineralizing capacity in BMs containing nonprimed soil was observed only after 12 weeks of linuron irrigation, matrix samples in the current experiment were taken for linuron mineralization assays after 15 and 20 weeks of incubation. BMs of set-ups L+ and C+ behaved identical to those in the first experiment (data not shown). However, the development of a linuron mineralization capacity was observed with only one of the other tested nonprimed soils, i.e. soil A1, and in only one replicate BM (lag time 4.9 days). After 20 weeks of linuron treatment, the recorded linuron mineralization curves of the BMs were similar to those recorded after 15 weeks of treatment (data not shown).

Size and composition of the Variovorax community in bioaugmented BPS

In the first experiment, the Variovorax 16S rRNA gene copy number increased both in BMs of set-up L+ and in BMs of set-up L between the start of the experiment and week 12 while the number of bacterial 16S rRNA gene copies remained stable (Table 3). The increases were, however, insignificant. In BMs of set-ups C and C+, the Variovorax 16S rRNA gene copy number decreased after 12 weeks of incubation, but only the decrease observed in set-up C was significant (Table 3). Double PCR-DGGE analysis of the Variovorax community in BMs of set-ups L, L+, C and C+ showed similar fingerprints at week 0 for all set-ups with bands at positions V1, V2 and V4 (Fig. 3a). Position V1 corresponds to the migration position of the 16S rRNA gene fragment amplified from the linuron-degrading Variovorax sp. WDL-1, which belongs to Variovorax phylotype B (Bers et al., 2011). 16S rRNA gene fragments at position V2 correspond to Variovorax members of phylotype C, which does not contain Variovorax strains with a linuron degradation capacity (Bers et al., 2011). The fingerprints of week 2 and week 12 showed the same bands at positions V1, V2 and V4. However, in set-ups L and L+ an additional band appeared at position V3. This band especially dominated in set-up L+ at week 12 and as such can be associated with the observed increased capacity to mineralize linuron. Position V3 corresponds to the migration position of the 16S rRNA gene fragment amplified from the linuron-degrading Variovorax sp. SRS16, which is a member of the Variovorax phylotype A (Bers et al., 2011). A band at position V3 also appeared in the profiles of set-ups C and C+ at week 12, but no clear differences in intensity were found between C and C+.

Table 3.   Number of bacterial and Variovorax 16S rRNA gene copies in BMs of experiment 1 (set-ups L, L+, C and C+) at week 0 and after 2 and 12 weeks of irrigation with tap water with or without linuron and in BMs of experiment 2 (set-ups A1+, L+, O+ and C+) at week 0 and after 15 weeks of irrigation with linuron
Experiment 1Set-up LSet-up L+Set-up CSet-up C+
VariovoraxBacteriaVar./bact. (%)VariovoraxBacteriaVar./bact. (%)VariovoraxBacteriaVar./bact. (%)VariovoraxBacteriaVar./bact. (%)
Week 03.4 ± 1.4 × 1072.6 ± 0,1 × 10100.13 ± 0.066.7 ± 4.6 × 1071.6 ± 0.8 × 10100.42 ± 0.355.4 ± 3.8 × 1073.3 ± 1.4 × 1091.66 ± 1.114.1 ± 3.8 × 1074.9 ± 1.4 × 1090.84 ± 1.11
Week 28.7 ± 3.4 × 1073.7 ± 4.6 × 10100.23 ± 0.107.5 ± 1.1 × 1072.8 ± 0.3 × 10100.27 ± 0.042.1 ± 0.4 × 1074.9 ± 0.5 × 1090.43 ± 0.082.3 ± 2.4 × 1074.1 ± 1.9 × 1090.55 ± 0.06
Week 121.2 ± 0.7 × 1083.7 ± 3.5 × 10100.31 ± 0.223.1 ± 1.4 × 1083.1 ± 0.1 × 10100.72 ± 0.161.3 ± 1.1 × 1062.5 ± 1.1 × 1080.54 ± 0.499.0 ± 0.8 × 1063.3 ± 3.0 × 1090.28 ± 0.37
Experiment 2Set-up A1+Set-up L+Set-up O+Set-up C+
VariovoraxBacteriaVar./bact. (%)VariovoraxBacteriaVar./bact. (%)VariovoraxBacteriaVar./bact. (%)VariovoraxBacteriaVar./bact. (%)
  1. The data are average values with indicated SDs.

Week 04.5 ± 1.4 × 1074.4 ± 0.3 × 1081.06 ± 0.423.0 ± 1.5 × 1072.8 ± 0.5 × 1091.10 ± 0.571.7 ± 0.4 × 1081.8 ± 0.2 × 10100.19 ± 0.041.7 ± 0.8 × 1075.8 ± 5.7 × 1080.69 ± 0.76
Week 151.7 ± 0.8 × 1083.4 ± 3.3 × 1093.06 ± 2.001.0 ± 0.5 × 1092.2 ± 1.2 × 10105.04 ± 1.701.2 ± 0.3 × 1081.1 ± 1.1 × 10110.005 ± 0.051.2 ± 4.3 × 1088.0 ± 8.3 × 1093.31 ± 3.27
image

Figure 3.  16S rRNA gene-based DGGE fingerprints of the Variovorax community obtained from (a) BM samples taken at weeks 0, 2 and 12 from BMs of set-ups C, C+, L and L+ in the first experiment and (b) BM samples taken at weeks 0 and 15 from set-ups A1+, C+, L+ and O+ in the second experiment. (a) Only DGGE profiles obtained with 16S rRNA gene fragments produced by double PCR are shown. (b) DGGE profiles obtained with 16S rRNA gene fragments produced by double PCR (upper gel) and single PCR (lower gel). DGGE fingerprints of reference DGGE migration markers ref-nonVar and ref-Var are indicated with ‘R’ and ‘V’, respectively. Band V3 corresponds to phylotype A, band V2 with phylotype C and band V1 with phylotype B.

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Real-time PCR results performed with DNA extracted from the samples of set-ups L+, C+, A1+ and O+, taken at weeks 0 and 15 in the second experiment, showed that the number of bacterial and Variovorax 16S rRNA gene copies were initially not significantly different between the different set-ups (Table 3). The number of Variovorax 16S rRNA gene copies in BMs of set-ups A1+, C+ and L+ increased after 15 weeks of linuron treatment. However, the increase was significant only for set-ups C+ and L+. In set-up A1+, no significant increase was recorded in the Variovorax 16S rRNA gene copy number in the BM replicate that had developed a linuron mineralization capacity compared with the other two replicate BMs (data not shown). The Variovorax population in BMs of set-up O+ remained constant in size, while the total number of bacteria increased six times. Variovorax 16S rRNA gene double PCR-DGGE fingerprints in samples taken from BMs of set-ups L+, C+, A1+ and O+ of the second bioaugmentation experiment are shown in Fig. 3b. DGGE fingerprints of all set-ups at week 0 displayed profiles similar to those observed at week 0 in experiment 1, but with additional bands such as at position V3. This also accounted for the set-up O+ where no soil was added, indicating that the observed Variovorax populations originated from the biomix substrata peat or straw. As in the Variovorax profiles of set-up L+ in the first experiment, an additional band at position V3 became clearly dominant in BMs of set-up L+. Interestingly, at week 15, in set-up A1+, only the replica BM showing enhanced linuron-mineralizing capacity displayed a dominant band at position V1 in the Variovorax DGGE profiles, although this was clearly observed only after a single PCR approach (Fig. 3b lower gel), which associates this band with the observed linuron mineralization activity. Position V1 corresponds to the Variovorax phylotype B, which, as phylotype A, includes predominantly linuron-degrading Variovorax strains. The same band also appeared in all profiles of set-up C+, but was also observed at week 0.

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

This study demonstrates for the first time the potential of bioaugmenting BPS by means of pesticide-primed soil. Although this bioaugmentation approach has been previously applied with success in polluted soil and wetlands, it has never been tested in a complex biotechnological matrix such as the biomix of a BPS with 8 and 2.5 times higher total carbon and nitrogen percentages than an average soil biotope, and sustaining a high biomass and rich biodiversity. Moreover, in contrast with previous studies, a comparison was made between systems bioaugmented with pesticide-primed soil and nonprimed soil. Clear differences in initial linuron mineralization capacity and the time to establish a maximum mineralization capacity were observed between the linuron-primed soil and any nonprimed control soil. The nonprimed perfect control would have been a non-linuron-treated soil originating from the same field site as the linuron-primed soil but such a soil was not available. Nevertheless, the effect of linuron priming is accentuated in the second experiment where only two of the tested nonprimed soils resulted into the development of a linuron mineralization capacity.

In contrast to nonprimed soil, bioaugmentation with linuron-primed soil immediately enhanced the linuron mineralization capacity of the BMs. Moreover, feeding with linuron further increased the linuron-mineralizing capacity in the matrix. MPN counting demonstrated that the observed enhancement in linuron mineralization capacity could be related to an increase in size of the linuron-mineralizing population. Therefore, it can be concluded that despite the complex background, the high biomass and high biodiversity, the community or at least the linuron-degrading fraction in the added soil could proliferate in the biomix in case fed with linuron. Various studies, mostly performed with suspended cultures inoculated with pure bacterial strains, reported on the effect of a highly degradable carbon and nitrogen content on mineralization/degradation of pesticides or other pollutants. Positive effects of additional C and/or N sources on pesticide degradation were reported by Cullington & Walker (1999), Aslan & Türkman (2005) and Fogg et al. (2003b), while Breugelmans et al. (2010) noticed negative effects on degradation of linuron. On the other hand, the maintenance of the linuron-mineralizing capacity in BMs of set-up L throughout the experiment indicate that the linuron-degrading population present in the primed soil is sufficiently competitive in its new biotope and can establish even without apparent selective conditions. Similarly, Johnsen et al. (2007) reported the long-term survival of a polyaromatic hydrocarbon (PAH)-degrading community present in inoculum PAH-primed soil, in soils without PAH pollution.

BMs of set-up C+, inoculated with nonprimed soil C, started to mineralize linuron after an extended period of linuron supply. A similar result was obtained with one soil of the other tested non-linuron-primed soils. Development of a linuron mineralization capacity in those BMs occured only when the pesticide was supplied, showing the selective nature of the process. Because the set-ups showing this outcome had the maximum moisture contents and because the pH did not significantly differ between set-ups, it is very unlikely that such differences in environmental conditions were responsible for the observed differences in linuron mineralization development. Instead, it can be hypothesized that either the microbial community in those BMs genetically adapted to degrade linuron or a linuron-mineralizing bacterial population initially present in the soil at undetectably low numbers proliferated when linuron was added. Interestingly, different replicate BMs developed a linuron-mineralizing capacity either after different treatment periods (in case of bioaugmentation with soil C) or only in one BM replicate (in case of bioaugmentation with soil A1). This can be explained by the occurrence of genetic adaptation at different time points or in one replicate, or by the fact that low initial numbers of mineralizing biomass were unevenly distributed in the soil sample used for inoculation. A similar observation was reported by Cullington & Walker (1999) with diuron.

The number of Variovorax in BMs containing linuron-primed soil L and receiving linuron (set-up L+) increased concomitantly with the increase in linuron mineralization capacity, but this was statistically significant only in BMs operated in the second experiment. The increase in Variovorax 16S rRNA gene copy number was less pronounced compared with the 100-fold increase observed in soil L when it was supplied linuron on a long-term base (Bers et al., 2011). The reason for the observed difference in proliferation of Variovorax between set-ups L+ in the first and second experiment might be due to the longer incubation period (3 weeks) and/or due to the higher amount of nutrients because of the higher fraction of straw and peat (only 25 vol% soil inoculum) in the biomix used in the second experiment. On the other hand, in both experiments, Variovorax DGGE fingerprinting clearly demonstrated the proliferation of a particular Variovorax population in BMs bioaugmented with linuron-primed soil L and fed with linuron. This Variovorax population belonged to phylotype A, which includes only linuron-degrading Variovorax strains such as Variovorax SRS16 (Breugelmans et al., 2007). The same phylotype A band proliferated in soil L when continuously fed with a linuron-containing solution (Bers et al., 2011). The Variovorax 16S rRNA gene PCR products recovered from these BMs were cloned but only some randomly chosen clones were sequenced because the association of bands with specific phylotypes was demonstrated previously (Bers et al., 2011). Nevertheless, blast analysis of these sequences identified them all as Variovorax 16S rRNA genes (E-value <10−18) with sequences 100% identical to those reported by Bers et al. (2011). The enrichment of phylotype A Variovorax in these BMs was probably not detected by the Variovorax real-time PCR due to masking by the large number of Variovorax initially present in the biomix substrata. These data suggest that linuron-degrading Variovorax populations endogenous to the primed soil were successfully transferred to the BM biomix. In addition they suggest that strains belonging to phylotype A proliferated in the BM upon linuron supply, thereby enhancing the linuron degradation capacity. Because those populations only proliferated in case linuron was supplied and because they could be associated with a phylotype containing only linuron-degrading Variovorax, it can be suggested that these populations are involved in linuron mineralization in the L+ set-up. Conclusive evidence for this link can be provided using techniques such as DNA/RNA-stable isotope probing (Dumont & Murrell, 2005). It cannot be excluded that bacteria other than Variovorax were also involved in linuron degradation. DGGE analysis of bacterial 16S rRNA genes was performed but no differences were observed between set-ups L+ and L (data not shown).

Linuron is degraded through 3,4-dichloroaniline (3,4-DCA). Several 3,4-DCA-degrading strains have been isolated previously from soil L in co-culture with linuron-degrading Variovorax strains and it was suggested that the former proliferated by growth on 3,4-DCA leaking from the linuron-degrading Variovorax strains (Breugelmans et al., 2007). Some of these 3,4-DCA-degrading strains belonged to Variovorax phylotype C. The band associated with Variovorax phylotype C in the Variovorax DGGE profiles migrates at position V2, but its occurrence in BMs of set-up L+ could not be correlated with the appearance of the linuron mineralization capacity. An explanation is that the release of 3,4-DCA was limited during linuron degradation in the biomix, hindering the proliferation of 3,4-DCA-degrading bacteria or that, alternatively, 3,4-DCA degradation in the biomix was performed by other genera such as Comamonas as observed in other linuron-mineralizing bacteria consortia (Dejonghe et al., 2003).

No relationship was found between the linuron-mineralization capacity dynamics and Variovorax community dynamics (neither in size nor in structure) in BMs of set-up C+ in the first experiment. On the other hand, in experiment 2, a Variovorax population related to phylotype B (at position V1) appeared in the DGGE profile concomitantly with the appearance of a linuron mineralization capacity. Moreover, the Variovorax number increased concomitantly with the linuron mineralization capacity. Phylotype B includes the linuron-degrading Variovorax sp. WDL1. However, in experiment 2, no set-up was included without linuron feed and therefore the appearance of this phylotype B cannot be linked to the observed increased linuron mineralization capacity. Moreover, the presence of a band at position V1 was not correlated with an increase in linuron mineralization capacity in the first experiment. On the other hand, based on the single PCR-DGGE results, the proliferation of a Variovorax phylotype B and hence a WDL1-related Variovorax population could be clearly associated with the increased linuron mineralization capacity in BM1 of set-up A1+ because neither of the other two BM replicas in this set-up showed enhanced linuron mineralization capacity nor the DGGE band associated with Variovorax phylotype B. However, this association was less clear in the DGGE profiles obtained with the product of the double-PCR approach. Because a WDL1-related population was also detected in BMs without inoculum and in BMs with nonprimed soil A1 (as shown in the double PCR-DGGE approach) that did not develop a linuron-degrading capacity, it can be suggested that different WDL1-related populations were initially present in the straw and peat mix on the one hand and in the soil inoculum on the other.

In conclusion, the results suggest that it is preferable to use pesticide-primed soils containing endogenous pesticide-mineralizing microorganisms over nonprimed soil for bioaugmentation of on-farm BPS for several reasons. First, although microorganisms in soils without previous pesticide treatment can apparently develop a capacity to mineralize the pesticide used, this is not always the case. Second, the initiation of mineralization of the target pesticide, in case nonprimed soil is used, takes much more time compared with a system inoculated with pesticide-primed soil. This is important because mineralization should start as fast as possible in a BPS to minimize leaching of pesticides during the start-up period of the BPS. Furthermore, the results show that Variovorax populations originating from the added soil contribute to linuron degradation in the BPS at least in the systems containing linuron-primed soil.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References

This research was supported by IWT-Vlaanderen Strategic Basic Research project 73352 and IWT-Vlaanderen Agricultural Research project LBO 040272.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. References