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Keywords:

  • syntrophic degradation;
  • soil bacteria;
  • butachlor;
  • 2-chloro-N-(2,6-diethylphenyl) acetamide

Abstract

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

Two bacterial strains involved in syntrophic degradation of chloroacetamide herbicide butachlor were isolated from a rice paddy soil. Analysis of 16S rRNA gene sequences indicated that the two isolates were related to members of the genera Mycobacterium and Sphingobium, respectively. Thus, a pair consisted of Mycobacterium sp. J7A and Sphingobium sp. J7B could rapidly degrade butachlor (100 mg L−1) at 28 °C within 24 h, while each isolate alone was not able to completely degrade butachlor. The isolate Mycobacterium sp. J7A was observed to grow slightly on butachlor, possibly utilizing the alkyl side chain of butachlor as its carbon and energy source, but the isolate Sphingobium sp. J7B alone could not grow on butachlor at all. Gas chromatography–mass spectrometry on catabolic intermediates revealed that the strain J7A produced and accumulated 2-chloro-N-(2,6-diethylphenyl) acetamide (CDEPA) during growth on butachlor. This intermediate was not further degraded by strain J7A, but strain J7B was observed to be able to completely degrade and grow on it through 2,6-diethylaniline (DEA). The results showed that butachlor was completely degraded by the two isolates by syntrophic metabolism, in which strain Mycobacterium sp. J7A degraded butachlor to CDEPA, which was subsequently degraded by strain Sphingobium sp. J7B through DEA.


Introduction

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

The chloroacetanilide herbicide butachlor, N-(butoxymethyl)-2-chloro-N-(2′,6′-diethylphenyl)-acetamide, has been applied in agricultural soils to control pre-emergent weeds. Butachlor is one of the commonly applied herbicides in South America and Asia (Yu et al., 2003; Lo et al., 2008). Because of the extensive use of butachlor, this herbicide and its metabolites have been detected in various agricultural soil environments (Chiang et al., 2001). To identify the harmful effect of butachlor in the environments, extensive researches have been progressed based on toxicology. Several articles showed that butachlor exhibited toxicity to earthworm and affected microbial community structures and enzyme activities (Min et al., 2001; Muthukaruppan et al., 2004). Also, butachlor was observed to be harmful to some aquatic organisms (Tilak et al., 2007), which brought destruction of water environment. In addition, butachlor caused stomach tumors in rat and oxidative DNA damage in human (Coleman et al., 2000; Dwivedi et al., 2012). Therefore, cleanup of butachlor residues from the environment has been great concern. Several studies showed that butachlor could be degraded by both biotic and abiotic processes, and microbial degradation was the most important factor for degradation of butachlor in soil (Lin et al., 2000; Pal et al., 2006). Some bacteria and fungi have been reported to be able to use butachlor as the sole carbon source, and the formation of metabolites of butachlor were investigated (Chakraborty & Anjan, 1991; Dwivedi et al., 2010). Metabolic pathways of butachlor were proposed based on the results of metabolite analysis and enzyme study (Zhang et al., 2011; Zheng et al., 2012). However, there is only limited information on butachlor-degrading microorganisms, and there has been no report about syntrophic degradation of butachlor by soil microorganisms.

In this study, a syntrophic pair of bacterial strains J7A and J7B which rapidly and completely degrade butachlor were isolated from a rice paddy soil, and their degradation properties and metabolic pathway were identified using gas chromatography–mass spectrometry (GC-MS). To our knowledge, this is the first report on bacterial syntrophic degradation of butachlor.

Materials and methods

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

Chemicals

Butachlor (97.7% purity) was purchased from Sigma-Aldrich (St. Louis). 2-Chloro-N-(2,6-diethylphenyl) acetamide (CDEPA) (100% purity) was purchased from Accustandard (New Haven). All other chemicals used in this study were of analytical grade.

Media and culture conditions

The isolated bacterial pair was grown in minimal medium (MM) with 100 mg L−1 butachlor as the sole carbon source. Minimal medium contained Na2HPO4 1.42 g, K2HPO4 1.36 g, (NH4)2SO4 0.30 g, MgSO4·7H2O 0.05 g, CaCl2·H2O 5.80 mg, FeSO2·7H2O 2.75 mg, ZnSO4·H2O 1.20 mg, MnSO4·H2O 1.70 mg, Co(NO)·6H2O 0.38 mg, CuSO4·5H2O 0.24 mg, and (NH4)6Mo7O2·4H2O 0.13 mg per liter of distilled water. Peptone–Tryptone–Yeast extracts–Glucose medium (PTYG) was used for strain purification and maintenance. PTYG contained peptone (Difco Laboratories) 0.25 g, tryptone (Difco) 0.25 g, yeast extracts (Difco) 0.5 g, glucose 0.5 g, MgSO4 0.03 g, and CaCl2 0.003 g per liter of distilled water. All cultures were incubated at 28 °C, and liquid cultures were aerated by rotary shaking incubator (Vision Co., Korea) at 180 r.p.m.

Isolation and identification of butachlor-degrading bacteria

Rice paddy soil samples were taken at various locations in South Korea. These rice fields have been routinely treated with butachlor for several years. Samples from the top 15 cm of soil were taken, screened through a 2-mm pore-size sieve, and kept at 4 °C prior to use. A 20 g amount of each soil sample was transferred to each 100-mL sterile beakers, treated with butachlor dissolved in distilled water to a final concentration of 100 mg g−1 soil, and completely mixed. The treated soil was incubated with periodic mixing at room temperature. Four weeks after the butachlor application, a 1 g soil sample from each beaker was homo-genized with 9 mL of a sterilized 0.85% saline solution by shaking the preparation at 150 r.p.m. on a rotary shaker. Samples (0.1 mL) of appropriate 10-fold dilutions were inoculated into test tubes containing 3 mL of butachlor medium (100 mg L−1). The tubes were incubated at 28 °C for 4 weeks, and degradation of butachlor was analyzed by NanoDrop 2000c spectrophotometer (Thermo Fisher Scientific) and by high-performance liquid chromatography (HPLC; Shimadzu, Japan). The culture of the terminal positive tube showing substantial cell growth and < 20% of the butachlor remaining was enriched by two additional transfers into fresh medium. Each culture was streaked onto PTYG agar medium, and combinations of single colonies were then tested for butachlor degradation in fresh butachlor medium before strain purification.

The isolated butachlor-degrading bacteria were identified by the analysis of 16S rRNA gene sequences. Total genomic DNA was extracted from the isolates, and PCR amplification of 16S rRNA genes was performed with 27mf and 1492r primers, as previously described (Weiburg et al., 1991). The amplified 16S rRNA genes were sequenced using an ABI Prism BigDye Terminator Cycle Sequencing Ready Kit according to the manufacturer's instruction (Perkin-Elmer) with the sequencing primers 519r, 926f, and 1055r (Weiburg et al., 1991). Approximately 1400 obvious nucleotide positions were used for comparison with the data in GenBank using the Basic Local Alignment Search Tool (blast; Altschul et al., 1990). Sequences from nearest relatives were identified from the Ribosome Database Project (RDP) using the similarity-rank program of the RDP (Maidak et al., 1997). Phylogenetic tree was constructed by the neighbor-joining method using Jukes–Cantor matrix (Saitou & Nei, 1987). Bootstrap analysis was performed with 1000 replicates.

Analysis of growth and degradation patterns

The syntrophic pair was grown in PTYG medium for 24 h. Cells were collected by centrifugation at 19000 g for 10 min at room temperature and washed twice with mineral medium. Aliquots of resuspended cells were inoculated into flasks containing 200 mL of mineral medium supplemented with butachlor (100 mg L−1) as the sole carbon source at a final density of OD600 nm = 0.004. All cultures were incubated at 28 °C in the dark on a rotary shaker (150 r.p.m.). At specific intervals, aliquots of cultures were taken out and used to determine cell growth and the concentrations of butachlor. Cell growth was determined at OD600 nm. For the quantification of butachlor, 3 mL of methanol was added to a 3 mL culture of the flasks, mixed thoroughly, and filtered. After filtering, the culture was used for the measurement of optical densities using spectrophotometry and reverse-phase HPLC. The concentration of butachlor was calculated using standard curves prepared from the known concentrations of butachlor in the same medium.

Chemical analysis and identification of intermediates

Incubation conditions were same as above. The 5 mL of culture samples were centrifuged with swing rotor at 340 g for 10 min (Combi-514R; Hanil, Korea), and the supernatant was extracted with the same volume of dichloromethane or hexane/ethyl acetate (1 : 1) after saturation with NaCl. The organic solution was passed through sodium sulfate and evaporated. The residue was dissolved in 1 mL methanol and then filtered with PTFE syringe filters with a pore size of 0.2 μm (Pall Corporation). Filtered samples were analyzed by HPLC and GC-MS. Residual butachlor was analyzed by HPLC on a Luna 5u C18 column (4.6 mm × 250 mm). Butachlor was detected with SPD-10A VP UV–Vis detector (Shimadzu) at 230 nm. The mobile phase was methanol/water (80 : 20, v/v), and the flow rate was 1 mL min−1. Metabolites of butachlor were determined by GC-MS analysis. The GC-MS analyses were achieved in electron ionization mode (70 eV) with Perkin-Elmer clarus 680 GC equipped with DB-5MS column (30 m × 0.25 mm id; 0.25 μm film thickness). The column temperature system was programed from 100 °C (2 min hold) to 280 °C at 10 °C min−1 and then held for 20 min. The helium was used as the carrier gas at a constant flow of 1 mL min−1. The samples were analyzed in split mode (1 : 20) at an injection temperature of 280 °C and detected in the mass range from m/z 30 to 650.

Results and discussion

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

Isolation of butachlor-degrading bacteria

During isolation of butachlor-degrading bacteria from the enriched cultures, a few single colonies were observed to be able to degrade butachlor. However, some enriched cultures failed to produce purified strains able to degrade butachlor. When the different colony types from each of these cultures were combined on butachlor mineral medium, some mixed cultures were able to degrade and grow on butachlor. From these mixed cultures, seven pairs of presumably syntrophic bacteria (denoted by A and B) capable of degrading butachlor were isolated from seven different soils. Among the syntrophic pairs, J7 pair (J7A and J7B) degraded butachlor very rapidly and showed the highest cell growth. Thus, the J7 pair was chosen for further studies on syntrophic butachlor degradation. Butachlor in mineral medium was completely degraded by the J7 pair, and no dead-end products were accumulated during syntrophic biodegradation of butachlor when analyzed with HPLC.

Identification by 16S rRNA gene sequence analysis

Strain J7A could grow well on PTYG over a temperature range of 10–37 °C, and its colonies were irregular, raised, and white. This strain was Gram-positive and showed positive results for catalase and urease and negative for oxidase. The 16S rRNA gene sequence of strain J7A showed 99.3% similarity with Mycobacterium fluoranthenivorans FA-4T (AJ617741). Strain J7B grew well on PTYG over a temperature range of 10–37 °C, and its colonies were circular, convex, and yellow color. This strain was Gram-negative and, in contrast to a well-expressed catalase and oxidase activities, no urease activity was observed. J7B showed 98.7% degree of 16S rRNA gene sequence similarity with Sphingobium chlorophenolicum ATCC 33790T (X87161). Figure 1 shows phylogenetic trees of strains J7A, J7B, and their related species based on the 16S rRNA gene sequences. Many species of the genus Mycobacterium were reported to be able to degrade polycyclic aromatic hydrocarbons (Kleespies et al., 1996; Hormisch et al., 2004), and some species in the genus Sphingobium could degrade pesticides (Bao zhan et al., 2009; Bala et al., 2010). Several bacterial strains have been isolated as butachlor-degrading bacteria in previous studies, and these strains belonged to genera Pseudomonas, Streptrophomonas, Paracoccus, Rhodococcus, and Catellibacterium (Wang et al., 2007; Dwivedi et al., 2010; Zhang et al., 2011; Liu et al., 2012; Zheng et al., 2012). Strains J7A and J7B of this study are the first Mycobacterium and Sphingobium species reported that are involved in butachlor degradation.

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Figure 1. Phylogenetic trees of isolates J7A, J7B, and their related species based on the 16S rRNA gene sequences. Bootstrap values (%) are indicated at the nodes. (a) A tree of J7A and relative type strains. (b) A tree of J7B and relative type strains.

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Syntrophic biodegradation of butachlor by J7 pair

To understand growth pattern of syntrophic pair J7 on butachlor medium, the pair was inoculated into butachlor mineral medium (100 mg L−1 butachlor). The growth curve of the pair J7 is shown in Fig. 2. The syntrophic pair J7 grew very quickly on butachlor medium, taking only 1 day for degrading butachlor completely. The J7 pair was observed to degrade butachlor at relatively broad temperature range of 10–37 °C without producing any residual compounds. Previously reported butachlor-degrading bacteria left residual compounds or took long time to degrade butachlor. Paracoccus sp. FLY-8 left 35.02% of 100 mg L−1 butachlor 5 days after inoculation and Catelibacterium caeni sp. DCA-1T left 19.8% of 50 mg L−1 butachlor even after 4 days of inoculation (Zhang et al., 2011; Zheng et al., 2012). Rhodococcus sp. strain B1 also utilized butachlor as a sole carbon and energy source, but it took 5 days to completely degrade 100 mg L−1 butachlor (Liu et al., 2012). Thus, in view of its rapid degradation and complete mineralization of butachlor, the J7 pair could be an effective bioresource for removal of butachlor from contaminated soil environments.

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Figure 2. Degradation of butachlor and cell growth of J7 syntrophic pair. (◆) residual butachlor concentrations; (■) OD600 nm values of J7 pair in butachlor mineral medium.

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When each strain of the J7 pair was separately inoculated into butachlor medium, strain J7B alone was not able to degrade and grow on butachlor, but strain J7A alone could grow slightly on butachlor (Fig. 3a). Spectrophotometry data showed that strain J7A transformed butachlor to an intermediate, which was subsequently degraded by strain J7B (Fig. 3b), indicating that butachlor was degraded completely by syntrophic metabolism of the two strains. There have been some reports on the syntrophic biodegradation of environmental pollutants. A Gram-negative pair of Sphingomonas sp. TFEE and Burkholderia sp. MN1 showed syntrophic association in degrading organophosphate insecticide (Katsuyama et al., 2009). Pseudomonas palleronii S1 and Agrobacterium radiobacter S2 degraded 4-aminobenzenesulfonic acid in a syntrophic way (Feigel & Hans, 1993). A Gram-positive pair of Arthrobacter sp. and Streptomyces sp. also has been shown to degrade diazinon synergistically (Gunner & Zuckerman, 1968). In our study, it is interesting that the syntrophic pair J7 was consisted of a Gram-positive strain and a Gram-negative strain. Moreover, as this kind of syntrophic pairs were isolated more frequently than single microorganisms, it appeared that syntrophic biodegradation of butachlor could be ubiquitous in the rice paddy soils tested in this study.

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Figure 3. (a) Growth profiles of strains J7A and J7B in butachlor mineral medium. (▲) syntrophic growth of J7A and J7B; (◆) slight growth of J7A; (■) no growth of J7B. (b) Absorption spectrograms of bacterial cultures after 7 days of incubation in butachlor mineral medium. (a) butachlor mineral medium; (b) J7A culture; (c) J7B culture; (d) syntrophic culture of strains J7A and J7B.

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Analysis of intermediates and pathway during syntrophic butachlor biodegradation

To investigate the syntrophic degradation pathway of butachlor, strain J7A that alone was able to attack butachlor was inoculated into butachlor mineral medium, and then, the intermediate compound formed was purified and identified with GC-MS analysis. Initially, only one large peak A was detected with GC-MS in the butachlor medium, but after 12-h incubation with strain J7A, two large peaks A and B were detected (Fig. 4a). Peak A [Rt (min) = 16.29] was identified as butachlor, which originally was supplemented in butachlor medium. Peak B [Rt (min) = 12.12] appeared to be an intermediate produced by J7A during butachlor degradation, and this peak was identified as CDEPA (Fig. 4c). The result indicated that strain J7A transformed butachlor to CDEPA. To test whether the intermediate CDEPA could be degraded by strain J7B, the intermediate was purified from J7A culture after 7 days of incubation in butachlor medium. All butachlor in butachlor mineral medium (100 mg L−1 butachlor) was observed to be almost completely transformed to CDEPA after 7 days of incubation with J7A (Fig. 4b). When J7B was inoculated into CDEPA medium, the intermediate CDEPA began to be degraded by J7B. After 3 days of incubation with J7B, a new peak (peak C) was detected (Fig. 4b). The peak C with retention time of 6.30 min was identified as 2,6-diethylaniline (DEA; Fig. 4c). In subsequent experiments, the intermediate DEA was observed to be degraded by J7B but not by J7A. From the results described above, the most likely syntrophic pathway of butachlor by J7 pair was proposed as in Fig. 5. It was assumed that strain J7A transformed butachlor to CDEPA by N-dealkylation, and then, strain J7B completely degraded CDEPA through DEA. Regarding the small metabolite peak near by the CDEPA peak in Fig. 4b, we attempted to search the mass spectra of this peak in NIST mass spectrometry database. But we could not obtain reasonable mass spectrum for this metabolite peak in database. The structure of this metabolite could be determined with NMR spectroscopy in further research.

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Figure 4. (a) GC profiles of butachlor control and its metabolite produced during degradation by J7A. (b) GC profiles of CDEPA and its metabolite produced during degradation by strain J7B. (c) GC-MS spectra of butachlor control and its metabolites produced during degradation.

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Figure 5. Proposed metabolic pathway of butachlor by syntrophic pair J7.

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Zhang et al. (2011) isolated a butachlor-degrading bacterium, Paracoccus sp. FLY-8, from a rice field soil. They found that this strain degraded butachlor to DEA when its degradation products of butachlor were identified by GC-MS. As this strain mineralized butachlor, they analyzed some possible enzyme activities such as aniline dioxygenase and catechol 1,2-dioxygenase to infer the mineralization pathway. The cellular lysates of Paracoccus sp. FLY-8 exhibited these two enzyme activities, and thus, they proposed the plausible mineralization pathway of butachlor through alachlor, 2-chloro-N-(2,6-dimethylphenyl) acetamide, DEA, aniline, and catechol. When, based on the results of Zhang et al., we analyzed degradability of aniline and catechol by J7A and J7B, only strain J7B was able to degrade and grow on aniline and catechol. Thus, DEA appeared to be further mineralized by J7B through aniline and catechol in our syntrophic mineralization pathway of butachlor (Fig. 5). In contrast to Paracoccus sp. FLY-8 of Zhang et al. (2011), which converted butachlor to alachlor by C-dealkylation, our strain J7A did not produced alachlor as an intermediate during degradation of butachlor, and also, alachor was not degraded by our syntrophic pair. In the study of Liu et al. (2012), a Gram-positive bacterium Rhodococcus sp. strain B1 transformed butachlor to CDEPA by hydrolase (ChlH) responsible for N-dealkylation without producing alachlor. Thus, our strain J7A appeared to transform butachlor to CDEPA by hydrolase activity without going through alachlor. Further investigation on the hydrolase genes would ascertain the divergence of sequences of butachlor hydrolase genes among the butachlor-degrading bacteria.

Acknowledgements

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

This study was supported by the RDA Genebank Management Program from the Genetic Resources Division, National Institute of Agricultural Biotechnology.

References

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