Isolation of an Indigenous Imidacloprid-Degrading Bacterium and Imidacloprid Bioremediation Under Simulated In Situ and Ex Situ Conditions

Abstract

The Bacterial community structure and its complexity of the enrichment culture during the isolation and screening of imidacloprid-degrading strain were studied using denaturating gradient gel electrophoresis analysis. The dominant bacteria in the original tea rhizosphere soil were uncultured bacteria, Rhizobium sp., Sinorhizobium, Ochrobactrum sp., Alcaligenes, Bacillus sp., Bacterium, Klebsiella sp., and Ensifer adhaerens. The bacterial community structure was altered extensively and its complexity reduced during the enrichment process, and four culturable bacteria, Ochrobactrum sp., Rhizobium sp., Geobacillus stearothermophilus, and Alcaligenes faecalis, remained in the final enrichment. Only one indigenous strain, BCL-1, with imidacloprid-degrading potential, was isolated from the sixth enrichment culture. This isolate was a gram-negative rod-shaped bacterium and identified as the genus Ochrobactrum based on its morphological, physiological, and biochemical properties and its 16S rRNA gene sequence. The degradation test showed that approximately 67.67% of the imidacloprid (50 mg/l) was degraded within 48 h by strain BCL-1. The optimum conditions for degradation were a pH of 8 and $30^{\circ}C$. The simulation of imidacloprid bioremediation by strain BCL-1 in soil demonstrated that the best performance in situ (tea soil) resulted in the degradation of 92.44% of the imidacloprid (100 mg/g) within 20 days, which was better than those observed in the ex situ simulations that were 64.66% (cabbage soil), 41.15% (potato soil), and 54.15% (tomato soil).

Introduction

Imidacloprid (1-((6-chloro-3-pyridinyl) methyl)-N-nitro- 2-imidazolidinimine) belongs to the family of neonicotinoids, which is widely used for the control of defoliating insects. Its function was developed by interfering with the transmission of stimuli in the insect nervous system [2, 13, 25, 33]. Although imidacloprid has the advantages of high insecticidal activity and low mammalian toxicity, it is extremely toxic to birds, bees, and aquatic animals even at low concentrations [11, 12, 18, 28]. Furthormore, imidacloprid can easily be released into diverse environments and persists for a long time. In soils, this compound can persist for 48-190 days [3]. In addition, imidacloprid can be taken up by crops and thus enter the food chain [10, 16, 24], which may result in harm to aquatic organisms and humans.

Previous studies have revealed that imidacloprid can be removed by natural processes in natural environments, such as hydrolysis, photodegradation, and biodegradation [14,17,32,36]. Biodegradation is an attractive approach to clean up contaminants because of its easily operation, high applicability, low cost and complete destruction of the contaminant. Anhalt et al. [1] isolated a bacterium belonging to the genus Leifsonia that was capable of degrading imidacloprid in agricultural soils. Gopal et al. [14] also reported that 50 µg/ml of imidacloprid could be degraded to 69% within 20 days by Burkholderia cepacia from agricultural soils [14]. However, many researchers all faced the situation that pollutant-degrading microbes with high degradation capacities in the laboratory performed poorly in the field [6]. Exogenous pollutant-degrading microbes could easily be outcompeted by indigenous microbes when applying into the environments [34], which was caused by the fact that the plants, soil properties, and microecology were different between in situ and ex situ environments. Therefore, biodegradation capacity depends on not the degrading-microbe effectiveness but its compatibility with the environment [35]. It is very likely that a pollutant-degrading bacterium would be a more effective bioremediation tool in situ than ex situ because the bacterium has already adapted to the environment in situ. Previous reports showed that several bacterial strains indigenous to a particular habitat had shown to outcompete other artificially introduced strains in the oil-contaminated bioremediation investigations [5,15]. Importantly, the biological activity of pollutants and the intermediate metabolites become lost by microbial degradation [26]. Therefore, the isolation of indigenous pollutant-degrading bacteria and a detailed knowledge of their ecology are important prerequisites in the design and implementation of any bioremediation scheme.

Oolong tea is an important beverage crop in China. Imidacloprid was detected at levels outside of the acceptable range in tea leaves and soils, resulting in serious food safety problems and reduced amounts of tea for export, and was banned for pest control on tea in 2010 in China. Therefore, the identification of an effective pollutant-degrading microbe, especially an indigenous strain, is necessary for the proper bioremediation of imidacloprid-polluted soils in tea-growing areas. In the present study, we isolated an indigenous imidacloprid-degrading bacterium from the tea rhizosphere soil using enrichment cultures, and then identified and characterized its imidacloprid-degrading capabilities. In addition, we analyzed the microbial diversity during the enrichment process using DGGE to explore the ecological significance of the isolates. Finally, ex situ and in situ bioremediation experiments were performed using tea rhizosphere soils and cabbage, potato, and tomato soils, respectively, to investigate the effectiveness of the bioremediation in soil.

 

Materials and Methods

Chemicals and Media

Imidacloprid (N-[1-[(6-chloro-3-pyridyl)methyl]-4,5-dihydroimidazol-2-yl]nitramide) (99.9% purity) was obtained from the Fujian Inspection and Testing Center for Agricultural Product Quality and Safety (TCAPQS), Fuzhou, Fujian, China. All other chemicals and solvents used were analytical or HPLC grade.

The enrichment medium was mineral salt medium (MSM, g/l) composed of 1.0 NH4NO3, 1.0 NaCl, 1.5 K2HPO4, 0.5 KH2PO4, and 0.2 MgSO4·7H2O, and adjusted to pH 8.0 with 10 M NaOH. The Luria-Bertani medium (LB, g/l) contained 10.0 tryptone, 10.0 yeast extract, and 5.0 NaCl. For solid plates, 1.5% (w/v) agar was added. The medium was sterilized by autoclaving at 121℃ for 25 min.

Soils

The tea rhizosphere soil used for the isolation of imidaclopriddegrading bacteria and the in situ bioremediation experiments was collected from the 20 cm top layer of a tea plantation located in Anxi, Fujian (27°03’N, 118°20’E), southeastern China. The cabbage, potato, and tomato soils used for the ex situ bioremediation experiments were collected from trial fields at the Fujian Agriculture and Forestry University, China that had no history of imidacloprid application. The properties of the two soil types were as follows: tea rhizosphere soil: nitrogen 48.90 mg/kg, available phosphorus 18.56 mg/kg, available potassium 12.00 mg/kg, pH 4.5 (measured in water), and organic matter 69.16 mg/kg; cabbage, potato, and tomato soils: nitrogen 34.52, 52.74, and 21.94 mg/kg, respectively; available phosphorus 26.15, 34.57, and 30.54 mg/kg, respectively; available potassium 14.52, 12.57, and 13.97 mg/kg, respectively; pH of 7.6, 8.4, and 6.8, respectively (measured in water); and organic matter 105.73, 83.74, and 129.40 mg/kg, respectively. For each type of soil, ten samples were collected randomly, mixed together, and then transported to the laboratory in a plastic bag.

Enrichment, Isolation, and Analysis of an Imidacloprid-Degrading Strain

Five grams of tea rhizosphere soil was transferred to a 250 ml Erlenmeyer flask containing 50 ml of sterilized minimal salts medium (MSM) to create enrichment cultures for the isolation of imidacloprid-degrading microorganisms. Imidacloprid dissolved in acetone solution was added at a final concentration of 50 mg/l. The enrichment culture was incubated at 30℃ on a rotary shaker at 170 rpm for 7 days. Five milliliters of the enrichment culture was transferred into 50 ml of fresh enrichment medium containing 50 mg/l of imidacloprid for another 7 days, and three additional successive transfers were made. The final cultures were serially diluted and spread on MSM plates to isolate individual colonies. The plates were incubated at 30℃ for 2 days, and the colonies were picked and purified [9]. Their abilities to degrade imidacloprid were determined by high-performance liquid chromatography (HPLC) as described by Blasco et al. [4].

DGGE Analysis of the Microbial Community During the Enrichment Process

DGGE was used to analyze the bacterial communities in the tea rhizosphere soil and the different enrichment cycles (second to sixth). Whole-community DNA of the tea rhizosphere soil was extracted using the FastDNA SPIN Kit for Soil (Qbiogene, Inc., USA). To extract DNA from the enrichment cultures, the cultures were centrifuged (10,000 ×g) for 25 min at 4℃, and the pellet was redissolved in sterilized ddH2O (0.6 ml). The redissolved pellet was utilized for DNA extraction using a kit. The DNA was finally eluted in 100 µl of DNase/RNase-free water (Qbiogene) and stored at -80℃. The DNA was amplified using eubacteria primers for the 16S rRNA gene (F338GC: 5’-CGC CCG CCG CGC GCG GCG GGC GGG GCG GGG GCA CGG GGG GAC TCC TAC GGG AGG CAG ACG-3’; R518: 5’-ATT ACC GCG GCT GCT GG-3’) in an iCycler iQ (Bio-Rad, Hercules, CA, USA). All reactions were performed in a final volume of 25 µl containing 2.5 µl of buffer (160 mmol/l )undefined(NH4)2SO4, 670 mmol/l Tris-HCl (pH8.8), 0.1% Tween-20, and 25 mmol/l MgCl2) (BIORON, Germany), 400 mmol/l of each primer, 200 µmol/l dNTPs, 0.5 U of DFS-Taq polymerase (BIORON), and 1 µl of template DNA. For the second PCR, 1 µl of the first PCR product was used as the template. The following amplification conditions were used for the two PCRs: 94℃ for 3 min; 30 cycles of denaturation at 94℃ for 1 min, annealing for 1 min at 55℃ for the first PCR and at 48℃ for the second PCR, and primer extension at 72℃ for 2 min; and a final extension at 72℃ for 5 min [21].

The DGGE analysis was performed with a DGGE-2001 system from CBS Scientific (Del Mar, USA). The PCR products (20-30 µl) were used in the analysis and loaded onto 8% (w/v) polyacrylamidebisacrylamide (37.5:1) (Amresco, USA) gels with denaturation gradients from 45% to 70%, where 100% represents 7 mol/l of urea and 40% (v/v) of deionized formamide in 1× TAE electrophoresis buffer. The gels (22 cm × 17 cm) were run at 20 V for 15 min and then at 70 V for 16 h. They were maintained at a constant temperature of 60℃. The gels were stained for 20 min in 1.0× GelStar and destained for 30 min in distilled water prior to visualization.

The representative bands were excised with a sterile scalpel, and immersed into 20 µl of TE buffer, and then incubated overnight at 4℃. Two microliter of the DNA extract was used as PCR template for PCR performence with the primers F341 (CC TAC GGG AGG CAG CAG) and R534 (ATT ACC GCG GCT GCT GG). The PCR products were ligated into PMD-19T vector, and then transformed into E. coli DH5α. White transformants were selected on LB medium supplemented with X-gal (20 mg/ml), IPTG (50 mg/ml), and ampicillin (100 mg/ml), and cultured in LB broth with ampicillin at 37℃ for 12 h. Positive clones were detected by plasmid PCR and liquid cultures were used to determine the plasmid sequence. DNA sequencing was conducted by Shanghai Personal Biotechnology Co., Ltd and the resulting sequence was compared with gene sequences in GenBank using BLAST (http://www.ncbi.nlm.nih.gov/BLAST).

Identification and Characterization of the Imidacloprid-Degrading Strain

Isolates were characterized and identified based on morphological and physiobiochemical characteristics and 16S rRNA gene sequences. The cells’ morphological properties were examined by light microscopy (Leica, Inc. Wetzlar, Germany) and scanning electron microscopy (FEI, Inc. Oregon, USA).

Physiobiochemical tests were performed using a physiological and biochemical kit purchased from Beijing Luqiao Technology Co., LTD. A series of characteristics were assessed, including carbon source utilization, hydrogen sulfide production, nitrate reduction, gelatin liquefaction, starch hydrolysis, Voges-Proskauer (V-P) test results, and indole test results. In the tests for carbon source utilization, Simmons citrate, lactose, glucose, maltose, amylum, D-galactose, mannose, D-fructose, and D-xylose were selected as the carbon sources.

Genomic DNA was prepared according to rapid methods [23]. The 16S rRNA gene was amplified by PCR with intF (AGAGTT TGATCCTGGCTCAG) and intR (GGCTACCTTGTTACGACT) as universal primers. PCR was performed with an iCycler thermocycler (BioRad, Sydney, Australia) using the following program: 5 min of pre-heating at 95 ℃; 30 cycles of 30 sec of denaturation at 95℃, 30 sec of primer annealing at 55℃, and 2 min of elongation at 72℃; and a final extension step of 10 min at 72℃. Then, the PCR products were cloned into a pMD 18-T vector (TaKaRa), and the plasmid was transformed into E. coli DH5α cells. Positive clones were identified and sent to Invitrogen Biotechnology Co., Ltd., for sequencing. The resulting sequence was compared with gene sequences in GenBank using BLAST (http://www.ncbi.nlm.nih.gov/BLAST). The sequences with the highest partial 16S rRNA sequence similarities were selected and compared using Cluster analysis. Phylogenetic and molecular evolutionary analyses were conducted using MEGA 5.0 software. Distances were calculated using the Kimura 2-parameter model. Unrooted trees were constructed using the neighbor-joining algorithm. The data were bootstrapped1,000 times [37].

Biodegradation of Imidacloprid by Strain BCL-1

Imidacloprid degradation in liquid culture was performed using a washed cell suspension pre-grown in MSM. Each 250 ml flask contained 100 ml of MSM inoculated with 1.0% (v/v) imidacloprid-degrading bacterium (OD600nm=1.0), and the initial pH was adjusted to 8.0. The degradation rate of imidacloprid in the culture over 48 h was measured at 2 h intervals. The effects of different temperatures (20℃, 25℃, 30℃, 35℃, and 40℃) and medium pHs (5, 6, 7, 8, and 9) on the imidacloprid degradation rate were investigated. All flasks were incubated in triplicate at 30℃ and 170 rpm on a rotary shaker. Uninoculated medium was used as a control.

Five milliliters of culture was collected from each flask at 24, 48, and 96 h and transferred to a 50 ml glass-stoppered flask. The cultures were extracted twice with 10 ml of dichloromethane using ultrasonic-assisted extraction in an ultrasonic crusher (SCIENTA JY99, China). Then, the organic liquid phases were combined and concentrated in a rotary evaporator (BUCHI, Inc., Switzerland). The residues were dissolved in 2 ml of methanol and filtered with a 0.45 µm membrane. The imidacloprid concentration was measured using a Hitachi 2000 HPLC system equipped with a Symmetry C18 reversed-phase column (0.5 µm × 4.5 mm × 250 mm) at a detection wavelength of 270 nm. A mixture of methanol and water (90:10)undefined(v/v) was used as the mobile phase at a flow rate of 1.0 ml/min. The injection volume was 10 μl.

Simulation of the Bioremediation of Imidacloprid in Soil

To investigate the effectiveness of the imidacloprid-degrading strain in the bioremediation of imidacloprid-contaminated soils, we performed the experiments in situ and ex situ using tea rhizosphere soil, and cabbage, potato, and tomato soils, respectively. Four thousand grams of each type of soil was placed in a 5 L Erlenmeyer flask, and the moisture content was adjusted to 40%. The soil moisture content was maintained at a constant level throughout the experiment by the addition of distilled water when necessary. Imidacloprid was added to a final concentration of 100 mg/kg. After mixing, a suspension of the imidaclopriddegrading strain was used to inoculate the soil (in triplicate) at a final concentration of 1.0 × 106 CFU/g . After inoculation, the soil was incubated at 35℃. Soil treated with the same amount of imidacloprid without bacteria served as uninoculated control samples. Twenty grams of soil was collected from each sample on days 5, 10, 15, and 20 for analysis of the residual imidacloprid concentration by HPLC (HPLC-1100; Agilent Technologies, USA). The extraction and analysis protocols were based on the methods outlined by Blasco et al. [4] and Sabale et al. [27], respectively.

 

Results

Microbial Community Change During the Enrichment Process Using DGGE

As shown in Fig. 1, the bacterial community during the enrichment process was substantially altered compared with the one in the original tea rhizosphere soil. It was obvious that the complexity of the bacterial community was reduced during the latter enrichment steps (Table 1). Several bands presented in the fingerprint of the tea rhizosphere soil bacterial community became less dominant or disappeared completely during the enrichment process.

Twenty-six bands that were dominant in the fingerprint of the tea rhizosphere soil bacterial community (Fig. 1, Table 1) were identified as different uncultured bacteria (band 1, HQ121331.1; band 4, EF196941.1; band 6, JX133525.1; band 9, HE819608.1; band 11, JQ957842.1; band 16, JN172809.1; band 23, JN172788.1), Rhizobium sp. (band 1, JX174273.1; band 18, JN819573.1; band 24, JN819573.1), Sinorhizobium (band 7, JX133181.1), Ochrobactrum spp. (band 5, KC153015.1; band 10, KC252620.1; band 13, KC146415.1), Alcaligenes (band 14, JX849036.1; band 19, JX975452.1), Bacillus sp. (band 12, HQ202555.1; band 15, GU566326.1; band 17, HM451429.1; band 20, FJ581462.1; band 21, JN700141.1; band 25, AY484507.1), Bacterium (band 3, JN792200.1), Klebsiella sp. (band 22, KC455419.1; band 26, GU997596.1), and Ensifer adhaerens (band 8, JX298811.1). However, bands 3, 6, 7, 8, 11, and 17 disappeared at the first and second enrichment, and then more bands were absent including bands 4, 14, 15, and 24 after the third and fourth enrichment. Along with the continual enrichment, only eight bands (2, 5, 9, 10, 13, 16, 19, and 20), which included four culturable bacteria, Ochrobactrum sp., Rhizobium sp., Geobacillus stearothermophilus, and Alcaligenes faecalis, appeared in the final enrichment and exhibited to be favored by the enrichment process (Fig. 1, Table 1).

Fig. 1.Denaturing gradient gel electrophoresis profiles of 16S rRNA gene fragments from bacteria within the tea rhizosphere soil (right panel, S) and various stages of enrichment culture (left panel, from M-1 to M-6). S stands for the tea rhizosphere soil; M stands for sample from the enrichment. Numbers stand for the time in the enrichment process.

Table 1.+: The bacteria was detected in the sample; -: the bacteria was undetected in the sample.

Isolation and Identification of an Imidacloprid-Degrading Bacterium

After the sixth enrichment, only one strain isolated from tea rhizosphere soil, named BCL-1, was found to be capable of growing on minimal salts medium in the presence of imidacloprid at a concentration of 20 mg/l. The degradation test demonstrated that the bacterium could degrade 67.67% of 50 mg/l imidacloprid within 48 h (Fig. 2). Strain BCL-1 was aerobic and rod-shaped, with a length of 2.48 µm and a width of 1.34 µm (Fig. 3). The colonies ranged from yellow to creamy white in color on MSM plates (Fig. 3). BCL-1 had positive results for the starch hydrolysis, nitrate reduction, and hydrogen sulfide production tests and was able to utilize Simmons citrate, lactose, glucose, maltose, amylum, D-galactose, D-fructose, and D-xylose. This strain had negative results for gram staining, the Voges-Proskauer (V-P) test, the indole test, gelatin liquefaction, and mannose utilization. PCR amplification of the 16S rRNA yielded a single fragment of 1,337 bp. BLAST analysis of the partial 16S rRNA sequence of BCL-1 showed that it was closely related to Ochrobactrum anthropic strain W-7 (GenBank Accession No. EU187487.1), with sequence identities of 99%. A phylogenetic tree was constructed based on the 16S rRNA gene sequence of strain BCL-1 and related strains using MEGA 5.0 (Fig. 4). Based on its morphological and physiobiochemical characteristics and its 16S rRNA gene sequence, BCL-1 was tentatively identified as O. anthropic with the accession number KC920738. It was interesting that O. anthropic was the predominant bacteria during the final enrichment.

Fig. 2.Degradation dynamics of imidacloprid in minimal salts medium within 48 h by inoculating strain BCL-1 at 30℃. The degradation rate was measured at 2 h intervals. Error bars represent the standard error of three replicates.

Degradation of Imidacloprid

The effect of pH on the degradation of imidacloprid is shown in Fig. 5. The imidacloprid degradation rate by O. anthropic strain BCL-1 increased to 46.58% at pH 5.0 and reached the maximum at pH 8, but reduced to 37% at pH 9 after 96 h of inoculation. The degradation efficiency increased from 34.41% over 24 h to 84.25% over 96 h at pH 8, which suggested that imidacloprid was easily hydrolyzed in neutral to alkali solutions (pH 7.0-8.0).

Fig. 3.Morphology of strain BCL-1 assessed using scanning electron microscopy (A) (×25,000) and light microscopy (B) (×1,000), and the characteristics of the colonies of this strain grown on minimal salts medium (C).

Data in Fig. 6 show that the incubation temperature greatly influenced the degradation of imidacloprid by strain BCL-1. The maximum degradation rate of 76.8% was observed at 30℃ within 96 h, but this rate decreased markedly as the temperature increased above or dropped below 30℃, and the degradation rate was only 29.4% at 20℃. The results displayed that 30oC was the optimal temperature for the degradation of imidacloprid.

Bioaugmentation Potentials In Situ and Ex Situ

The results showed that the degradation of imidacloprid by BCL-1 in situ (tea soil) preceded to the one ex situ (cabbage, potato, and tomato soils) (Fig. 7). After 10 days of testing, the degradation percentage of imidacloprid was 40.00 in tea soil, whereas the levels of degradation in the cabbage, potato, and tomato soils were 21.63%, 26.50%, and 31.65%, respectively. In the case of uninoculation soils, similar rates of degradation were seen; 16.28% (tea soil), 18.41% (cabbage soil), 19.33% (tomato soil), 18.64% (potato soil). On the 20th day, 92.44% of the applied dose of imidacloprid was removed from the tea soil by the introduced strain BCL-1, compared with 64.66%, 41.15%, and 54.15% in the cabbage, potato, and tomato soils, respectively. In contrast, the imidacloprid degradation rates in the uninoculated treatment were lower, at 38.21% (tea soil), 34.85% (cabbage soil), 36.68% (tomato soil), and 34.86% (potato soil). The results demonstrated that the imidacloprid degradation should be enhanced by inoculating the degrading strain BCL-1, and also further confirmed the finding that bioremediation in situ was much better than ex situ, with the degradation rate in tea soil by inoculating BCL-1 being much faster than that in other soils.

Fig. 4.Phylogenetic tree resulting from the analysis of 16S rRNA gene from imidacloprid-degrading bacterium BCL-1 and related species. The tree was constructed using the neighbor-joining algorithm and ñ-distance estimation method of the software MEGA 5.0. Bootstrap values (> 50%) generated from 1,000 replicates are shown as percent values at the nodes.

Fig. 5.The effect of initial pHs of 5, 6, 7, 8, or 9 on the imidacloprid degradation rate by strain BCL-1 at 30℃ in sterilized minimal salts medium. Error bars represent the standard error of three replicates.

Fig. 6.The effect of incubation temperatures of 20℃, 25℃, 30℃, 35℃, or 40℃on the imidacloprid degradation rate by strain BCL-1 in sterilized minimal salts medium. Error bars represent the standard error of three replicates.

Fig. 7.Simulation experiments of imidacloprid bioremediation in situ and ex situ, by inoculating strain BCL-1. The in situ experiment was conducted in tea soil and one without inoculation was considered as control (CK); ex situ experiments were performed in cabbage soil, potato soil, and tomato soils with the uninoculated as control (CK). Error bars represent the standard error of three replicates.

 

Discussion

Traditionally, pesticide-degrading microorganisms are isolated via enrichment cultures [22]. However, the interactions of pesticide-degrading bacteria with other microorganisms during the enrichment process are unclear. We analyzed the alteration of the microorganism community during the enrichment process using DGGE. While selection for imidacloprid degradation was maintained, the dominant species decreased in abundance, and many nondegrading bacteria were displaced by other potential imidaclopriddegrading bacteria. The bands favored during the enrichment process were dominant in the bacterial community of the enrichment cultures and later were isolated for imidacloprid degradation. We isolated strain BCL-1, capable of degrading imidacloprid, as the predominant bacterium in the enrichment. Similar results were obtained by Sorensen et al. [31], who reported that several linuron-degrading Variovorax strains isolated from the soils were dominant in the bacterial community of the enrichment cultures [31]. The study conducted by Breugelmans et al. [7] demonstrated the same phenomenon. Chanika et al. [8]also revealed that two isolates capable of degrading both organophosphate and fenamiphos were dominant members of the enrichment culture. However, Simonsen et al. [30] noted that a 2,6-dichlorobenzamide-degrading Aminobacter strain was not among the dominant bacteria in the enrichment culture from which it was isolated [30].

In the present study, strain BCL-1, isolated from tea rhizosphere soil, was identified as Ochrobactrum sp. and was shown to degrade 50% of the imidacloprid (50 mg/l) present in the culture within 26 h and approximately 70% within 48 h. In addition to species of the Ochrobactrum genus, many individual isolates have been found to be able to degrade imidacloprid, such as Leifsonia sp. [1], Burkholderia cepacia [14], Enterobacteriaceae, Pseudomonas, Bacillaceae [29], and Stenotrophomonas maltophilia [19]. Liu et al. [20] reported that imidacloprid could be degraded by soil microorganisms but with poor degradation efficiency.

In a new bioaugmentation approach, achievable bioremediation requires the creation of unique niches or microhabitats for the desired pollutant-degrading microbes, and therefore, the selection and culturing of microbes directly from the local site is necessary. Bioaugmentation of contaminated soil with commercial preparations has been less promising. When commercial products are used, microbial species that may be much different from those of the local environment are often applied. The best bioaugmentation performance can be achieved by using microorganisms that are already present in the soil and increasing their abundance. Indigenous microorganisms are well adjusted to their own environments. An immediate increase in the population density of the microbes could ensure the rapid degradation of the pollutant. The results of our experiment comparing the bioaugmentation potentials of strain BCL-1 in in situ and ex situ environments support this hypothesis. We determined the bioremediation rate of the imidaclopriddegrading bacterium BCL-1 in situ (in tea soil) and ex situ (in cabbage, potato, and tomato soils) and found that the imidacloprid degradation rate was significantly higher in situ than ex situ. Obviously, strain BCL-1 from the tea rhizosphere had best bioremediation performance in the tea soil. However, more evidence is needed to support the hypothesis that strains with native ecological niches are more effective tools for bioremediation. Further research into the chemical metabolism mechanisms, physiological regulation, energy consumption, cellular respiration and nutrition circulation of the pollutant-degrading microbes used in bioremediation should also be conducted.