Abstract
Acetochlor [2-chloro-N-(ethoxymethyl)-N-(2-ethyl-6-methylphenyl)-acetamide] is a widely applied herbicide with potential carcinogenic properties. N-Deethoxymethylation is the key step in acetochlor biodegradation. N-Deethoxymethylase is a multicomponent enzyme that catalyzes the conversion of acetochlor to 2′-methyl-6′-ethyl-2-chloroacetanilide (CMEPA). Fast detection of CMEPA by a two-enzyme (N-deethoxymethylase–amide hydrolase) system was established in this research. Based on the fast detection method, a three-component enzyme was purified from Rhodococcus sp. strain T3-1 using ammonium sulfate precipitation and hydrophobic interaction chromatography. The molecular masses of the components of the purified enzyme were estimated to be 45, 43, and 11 kDa by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Based on the results of peptide mass fingerprint analysis, acetochlor N-deethoxymethylase was identified as a cytochrome P450 system, composed of a cytochrome P450 oxygenase (43-kDa component; EthB), a ferredoxin (45 kDa; EthA), and a reductase (11 kDa; EthD), that is involved in the degradation of methyl tert-butyl ether. The gene cluster ethABCD was cloned by PCR amplification and expressed in Escherichia coli BL21(DE3). Resting cells of a recombinant E. coli strain showed deethoxymethylation activity against acetochlor. Subcloning of ethABCD showed that ethABD expressed in E. coli BL21(DE3) has the activity of acetochlor N-deethoxymethylase and is capable of converting acetochlor to CMEPA.
INTRODUCTION
Chloroacetanilide herbicides are a class of important herbicides used worldwide. The chloroacetanilide herbicide acetochlor [2-chloro-N-(ethoxymethyl)-N-(2-ethyl-6-methylphenyl)-acetamide] is a selective preemergent herbicide that has been used to effectively control broadleaf weeds and annual grasses in corn fields for almost 40 years (1). The excessive and frequent application of acetochlor may result in high levels of acetochlor residues, which have been detected in ground and surface waters and negatively impact both the environment and agricultural ecosystems (2). The U.S. Environmental Protection Agency (EPA) has classified acetochlor as a B-2 carcinogen and a probable human carcinogen (3). Studies of different soil types treated with acetochlor have demonstrated that 2′-methyl-6′-ethyl-2-chloroacetanilide (CMEPA), a compound derived from acetochlor N-deethoxymethylation, is a major product of the microbial degradation process (1, 3–5).
We previously isolated an efficient acetochlor N-deethoxymethylation strain, T3-1, from a microbial consortium that could mineralize acetochlor completely. The strain was identified as a Rhodococcus sp. (1). The biochemical pathway of acetochlor degradation by the three bacteria in the consortium was proposed to be as follows: acetochlor to CMEPA by Rhodococcus sp. strain T3-1, CMEPA to 2-methyl-6-ethyl aniline (MEA) by Delftia sp. strain T3-6, and MEA by Sphingobium sp. strain MEA3-1, based on the identified degradation intermediates (1). In this study, a three-component cytochrome P450 system that deethoxymethylated acetochlor into CMEPA was purified from Rhodococcus sp. T3-1, and its encoding gene was cloned and expressed in Escherichia coli BL21(DE3). The activity of N-deethoxymethylase that converts acetochlor to CMEPA has been detected by subcloning of the gene cluster into E. coli BL21(DE3).
MATERIALS AND METHODS
Chemicals and media.
Acetochlor was purchased from Shanghai Jingchun Co. Ltd. CMEPA (96.5% purity) was purchased from Qingdao Vochem Co. Ltd. (Shandong, China). All other chemicals used in this study were purchased from Sigma (St. Louis, MO, USA) and were analytical grade or higher purity.
Minimal salts medium (MSM) contained (liter−1) 1.5 g K2HPO4, 0.5 g KH2PO4, 1.0 g NH4NO3, 0.10 g MgSO4 · 7H2O, and 1.0 g NaCl, with 25 mg acetochlor added to the medium as the sole carbon source. Stock solutions of different chloroacetamide herbicides with methanol at 10 g/liter were prepared by membrane filtration with a pore size of 0.22 μm. The solutions were added to the sterilized MSM or the reaction solution to determine the activities of strain T3-1 and the spontaneous mutants.
Luria-Bertani (LB) medium contained 10.0 g/liter tryptone, 5.0 g/liter yeast extract, and 10.0 g/liter NaCl, pH 7.0.
Isolation of spontaneous mutants unable to degrade acetochlor.
Rhodococcus sp. T3-1 (China Center for Type Culture Collection [CCTCC] no. M 2012525) was streaked on an LB plate, and an independent colony was transferred to a fresh LB plate. After 23 transfers, a colony with different morphology was observed and purified, and its ability to degrade acetochlor was determined. The enterobacterial repetitive intergenic consensus (ERIC)-PCR pattern (6) and 16S rRNA gene sequence of the mutant strain were determined and compared with those of the wild type. Strains that lost the ability to degrade acetochlor were considered mutants.
Bacterial cultivation and crude cell extract preparation.
Mutant and wild-type strains of Rhodococcus sp. T3-1 were cultivated in appropriate volumes of LB broth to exponential phase at 30°C in a 180-rpm shaking incubator. Cells were harvested for 15 min at 5,000 rpm (Beckman [USA] Allegra X-22R) at 4°C. The pellets were resuspended and washed with buffer A (20 mM Tris-HCl [pH 7.0]) and collected by centrifugation as described above. The collected cells (1 g) were ground with quartz sand in liquid nitrogen. The homogenate was resuspended in solution with buffer B (20 mM Tris-HCl, pH 7.0, 40% sucrose, 0.1 M NaCl, 10 mM MgCl2, 2 mM β-mercaptoethanol) and centrifuged at 12,000 rpm for 20 min at 4°C to remove the quartz sand and cell debris. The supernatants obtained from this step were referred to as crude cell extracts. Protein concentrations were determined using the Bradford method (7) with bovine serum albumin as a standard. Total proteins of the supernatants of the mutant and wild type were analyzed by denaturing SDS-10% (wt/vol) polyacrylamide gradient gel electrophoresis.
Standard assay for enzyme activities with cell extracts.
Acetochlor N-deethoxymethylase activity was detected indirectly by a two-step catalytic chromogenic method. Acetochlor was first transformed to CMEPA by N-deethoxymethylase, and then CMEPA was hydrolyzed to MEA by DamH from Delftia sp. T3-6 (8). MEA could be measured conveniently by a spectrophotometric method.
To set up the reaction mixture, 100 μl crude cell extract was added to 400 μl buffer B containing 1 mM acetochlor and 1 mM NADH. Purified DamH (10 μl) was added to the mixture to start the reaction, and the reaction mixture was incubated at 30°C for 30 min. After the incubation, 5 μl 0.2 M potassium hexacyanoferrate(III) and 5 μl 0.1 M 4-aminoantipyrene were added to the reaction mixture. MEA reacted with 4-aminoantipyrene to form a purple compound with maximum absorption at 545 nm (9).
Enzyme purification.
All purification steps for acetochlor N-deethoxymethylase were performed at a temperature of 4°C to avoid any possible enzyme denaturation. The cell extracts were fractionated with ammonium sulfate, and the precipitate (70 to 80% saturation) was harvested by centrifugation at 12,000 rpm for 20 min. The precipitates were dissolved in ∼100 ml of buffer A (20 mM Tris-HCl [pH 7.0] containing 70% ammonium sulfate) (10). The enzyme was then applied to a Butyl-650M hydrophobic interaction chromatograph (Toyopearl, Japan). The eluted fractions from the chromatographic separations were monitored at 280 nm and collected at 1-ml intervals. The fractions containing acetochlor N-deethoxymethylase activity were collected and concentrated with a Millipore Centriprep Centifugal Filter Concentrator (11).
Amide hydrolase (DamH) was purified from recombinant E. coli BL21(DE3) by Ni-nitrilotriacetic acid (NTA) affinity chromatography as described previously (8).
Protein sequencing and mass spectroscopy analysis.
The molecular weights of purified proteins were estimated using electrophoresis on SDS-12% PAGE by the method of Laemmli (12). The stained gel band was excised and used for peptide mass fingerprint analysis (Bo-Yuan Biological Technology Co. Ltd.). The resulting peptide fragments were analyzed by searches in Mascot website databases (Matrix Science) to determine the sequences of the peptide fragments.
Gene cloning and expression of EthABCD.
Genomic DNA was extracted from Rhodococcus sp. T3-1 as described previously to serve as the PCR template (13). Oligonucleotide primers were designed to amplify the eth genes (Table 1). The full-length eth gene cluster was PCR amplified using the following primers: EthA-F (forward), containing an NdeI site (underlined) at the initiation site of the ethA gene, and EthD-R (reverse), containing a NotI site (underlined) after the ethD gene. The ethABC gene cluster was PCR amplified using the following primers: EthA-F and EthC-R(N) (reverse, containing a NotI site [underlined] after the ethC gene). In order to construct the ethABD gene cluster, overlapping PCR was carried out to delete the ethC gene of ethABCD. The forward primer EthB-R(2) and the reverse primer EthD-F(2) were a pair of overlapping complementary primers that were synthesized to obtain the structural genes without ethC. The sequence of ethR was obtained by self-formed adaptor (SEFA) PCR (14), and the primers used are presented in Table 1. The primers EthR-sp1, EthR-sp2, and EthR-sp3 were used to amplify the sequence of ethR.
TABLE 1.
Oligonucleotide primers used in PCR
| Primer | Sequencea (5′ to 3′) | Target gene(s) |
|---|---|---|
| EthA-F | CATATGATCATCATCGGCGCCGGGCAG | ethA |
| EthA-R | CTCGAGGCGAGCCGTGGCGACCTCCCGGGCGATGT | |
| EthB-F | CATATGACACTGTCACTGGCCAC | ethB |
| EthB-R | GAGCTCCTTCGGGTAGATCCGCAC | |
| EthC-F | CATATGCCCAAGATCACCTTCTCCCAAT | ethC |
| EthC-R | CTCGAGGAACGCGTCGGGGACCTCCAG | |
| EthD-F | CATATGTATCAGATCGTGGCCTGCT | ethD |
| EthD-R | TGCGGCCGCGGTCCGGTCGACCTCATCCCCGAT | |
| EthA-F | CATATGATCATCATCGGCGCCGGGCAG | ethABC |
| EthC-R(N) | GCGGCCGCTCAGAACGCGTCGGGGACCTCCAGC | |
| EthA-F | CATATGATCATCATCGGCGCCGGGCAG | ethABD |
| EthB-R(2) | CCGGACGCGTTCACTTCGGGTAGATCCGCAC | |
| EthD-F(2) | GATCTACCCGAAGTGAACGCGTCCGGCACCAG | |
| EthD-R | TGCGGCCGCGGTCCGGTCGACCTCATCCCCGAT | |
| EthR-sp3 | GACATCGCTGCCTGCNNNNNNNNNTGCCCG | ethR |
| EthR-sp2 | GTTGATGAGATACGCCTTCG | |
| EthR-sp1 | CGACCCGAACTCAGCGACAC |
Restriction sites are underlined.
The PCR products were digested with NdeI and NotI, inserted into the NdeI-NotI sites of pET29a(+) to obtain the plasmid pET29a harboring eth genes (Table 2), and then transformed into E. coli BL21(DE3). The transformants were grown in LB medium containing 50 μg/ml kanamycin at 37°C until they reached mid-log phase; they were then induced with 0.2 mM IPTG (isopropyl-β-d-thiogalactopyranoside) at 18°C for 24 h.
TABLE 2.
Plasmids used in this study
| Plasmid | Characteristic(s) | Source or reference |
|---|---|---|
| pMD19-T | T-A clone vectors; Ampr | TaKaRa |
| pET-29a(+) | Expression vector; Kmr | Laboratory stock |
| pET-ethA | pET-29a(+) derivative carrying ethA | This study |
| pET-ethB | pET-29a(+) derivative carrying ethB | This study |
| pET-ethC | pET-29a(+) derivative carrying ethC | This study |
| pET-ethD | pET-29a(+) derivative carrying ethD | This study |
| pET-ethABCD | pET-29a(+) derivative carrying ethABCD | This study |
| pET-ethABC | pET-29a(+) derivative carrying ethABC | This study |
| pET-ethABD | pET-29a(+) derivative carrying ethABD | This study |
Crude cell extracts of various recombinant eth genes were prepared from the transformed E. coli strains; cells were suspended in buffer A and were disrupted by sonication (Sonicator 201M; Kubota, Japan). After centrifugation at 12,000 rpm for 20 min at 4°C, the resulting supernatant was used as the crude cell extract. The expression levels of EthA, EthB, EthC, and EthD in the crude extracts were examined using SDS-PAGE analysis.
Product analysis of reactions using resting cells of the transformed E. coli strains.
The transformants were grown in LB broth containing 50 mg/liter acetochlor and 50 μg/ml kanamycin at 37°C until they reached mid-log phase; they were then induced with 0.2 mM IPTG at 18°C for 48 h.
Acetochlor and CMEPA in the cultures were extracted with an equal volume of dichloromethane, and the extracts were dried over anhydrous Na2SO4. The extracts were evaporated at room temperature. Residual material was redissolved in an equal volume of methanol and examined by high-performance liquid chromatography (HPLC) with a UV detector (LC-20AD; Shimadzu, Japan). The mobile phase was 80% methanol, and the flow rate was 1.0 ml · min−1. Compounds were detected spectrophotometrically at a wavelength of 243 nm. The separation column (Inertsil ODS-SP; 4.6 by 250 mm) was filled with Kromasil 100-5C18, and the injection volume was 20 μl (1).
The metabolites formed during degradation were identified by gas chromatography-mass spectrometry (GC-MS) according to the methods described by Zhang et al. (15).
Extraction of nucleic acids, RNA quantification, and cDNA synthesis.
Rhodococcus sp. T3-1 and the mutant were cultivated using acetochlor as a carbon and energy source, as previously described, and cells cultivated in minimal salts medium containing 2 mM glucose were used as controls. A total of 10 ml of bacterial culture from the growth phase, with an optical density at 600 nm (OD600) of approximately 0.2, was filtered using polyethersulfone filters with a 0.2-μm pore size (Pall Corporation). Genomic DNA was extracted from Rhodococcus sp. T3-1 and the mutant cells and purified as described previously. The RNA extraction and the cDNA synthesis were carried out according to the method of Jechalke et al. (16). Oligonucleotide primers were used to amplify the eth gene as described above.
Nucleotide sequence accession numbers.
The sequences of the eth genes of Rhodococcus sp. T3-1 have been deposited in the GenBank database under accession numbers KM225842 (ethR), KM225843 (ethA), KM225844 (ethB), KM225845 (ethC), and KM225846 (ethD).
RESULTS
Isolation of acetochlor degradation-deficient mutants of Rhodococcus sp. T3-1.
In an attempt to test the stability of the acetochlor degradation phenotype, Rhodococcus sp. T3-1 bacteria were cultivated on LB plates and transferred continuously 23 times. A colony with different morphology was observed after 23 transfers. The colony was much rougher than that of strain T3-1 (Fig. 1). Strain T3-1 degraded acetochlor to CMEPA, which could be further degraded to MEA by the amide hydrolase DamH. This process could be simply traced by following the color development of MEA and 4-aminoantipyrene. Among the tested cultures from the continuous passages, a colony with different morphology was observed after 23 transfers, which had lost the ability to degrade aectochlor. The colony of the mutant strain was much rougher than that of strain T3-1. The 16S rRNA gene sequence of the mutant strain shared 100% identity with that of strain T3-1. The ERIC-PCR patterns of the two strains were compared and showed identical band distributions. These results indicated that the rough colony was indeed an acetochlor degradation-deficient mutant strain of strain T3-1. The strain was designated Rhodococcus sp. T3-1 MT. The results of continuous passages showed the acetochlor degradation phenotype was very stable in Rhodococcus sp. T3-1.
Comparison and analysis of total proteins of Rhodococcus sp. T3-1 and T3-1 MT.
Since the morphological and degradative characters of the mutant strain T3-1 MT were greatly changed, differences in the protein profiles of the strains might give clues to the enzymes related to acetochlor degradation. Crude cell extracts of both strains were analyzed by SDS-PAGE (Fig. 1). The two strains showed similar protein patterns, except that a protein band of about 43 kDa was missing in the mutant strain T3-1 MT. We deduced that this protein was related to the degradation of acetochlor. The protein band was cut, and the gel slice was used for peptide mass fingerprint analysis by liquid chromatography-tandem MS (LC–MS-MS). Peptide microsequencing yielded several peptide fragments with the following sequences: TIPGAILEGIR, AAAYKEKIQQAAVTLVEELLDRREFDAVLDFAQMMPMRVFMEVLGVEPDIEQRR, and YATDADVFAHDTLVDPYDTYR. These peptide sequences matched those of a cytochrome P450 monooxygenase, EthB, from Rhodococcus ruber IFP 2001 (gi|16551194) (17).
FIG 1.
Profiles of the total proteins of Rhodococcus sp. T3-1 and T3-1 MT as revealed by SDS-PAGE. Lane WT, Rhodococcus sp. T3-1 (wild type) total protein; lane MT. Rhodococcus sp. T3-1 MT total protein; lane M, protein molecular mass marker. The arrow indicates the protein band that is missing in the mutant strain T3-1 MT.
Purification of acetochlor N-deethoxymethylase from Rhodococcus sp. T3-1.
Because it was difficult to genetically manipulate Rhodococcus strains, we tried to purify the N-deethoxymethylase from Rhodococcus sp. T3-1. Acetochlor N-deethoxymethylase from Rhodococcus sp. T3-1 was purified using a two-step procedure described previously. In the first step, the culture supernatant was fractionated with ammonium sulfate. The fraction F6 showed activity higher than that of F1, with 55.7% protein recovery. No activity was detected in the other precipitated fractions.
The fraction F6, which gave the highest activity, was loaded onto to a Butyl-650M hydrophobic interaction chromatography column. The eluted fractions from the chromatographic separations were monitored using electrophoresis on SDS-12% PAGE, and the acetochlor N-deethoxymethylase activity of each fraction was detected. As shown in Fig. 2, the fractions (181 to 202) with acetochlor N-deethoxymethylase activity exhibited three bands on SDS-PAGE with molecular masses of approximately 45 kDa, 43 kDa, and 11 kDa.
FIG 2.
Analysis of the sample purified by a Butyl-650M column from Rhodococcus sp. T3-1 on SDS-PAGE. Lane 1, sample of fraction 181; lane 2, sample of fraction 184; lane 3, sample of fraction 187; lane 4, sample of fraction 190; lane 5, sample of fraction 193; lane 6, sample of fraction 196; lane 7, sample of fraction 199; lane 8, sample of fraction 202.
Peptide fingerprint analysis identified the three components, and the results are shown in Table 3. The three components were determined to be similar to a ferredoxin reductase, EthA (45 kDa; 98% identity);, a ferredoxin, EthD (11 kDa; 96% identity); and a cytochrome P450 monooxygenase, EthB (43 kDa; 93% identity). EthABD was a multicomponent cytochrome P450 monooxygenase system involved in the hydroxylation of ethoxy residues in the fuel oxygenate ethyl tert-butyl ether (ETBE) in R. ruber (17). The result of purification was consistent with that observed in the mutant strain, T3-1 MT.
TABLE 3.
Results of peptide mass of fingerprint analysis
| Identifier | Mass (Da) | Protein | Species or strain | Peptide |
|---|---|---|---|---|
| gi|16551194 | 43,746 | EthA | R. ruber | RLESVQNAVDHARH |
| RLDIDLITGDGVTRI | ||||
| RELSVPGADLAGVEALRT | ||||
| RGGPRPVAPVDGSPAAFDLKH | ||||
| RLTIVGDEPGLPYQRPPLSKA | ||||
| KIQIAGIGAQGAESVVIGDEAAERC | ||||
| gi|16551195 | 45,139 | EthB | R. ruber | KQHINNLIRS |
| MTLSLATAQERY | ||||
| KTIPGAILEGIRF | ||||
| RDVEYDDIVIPAGSRT | ||||
| KIQQAAVTLVEELLDRR | ||||
| RVFMEVLGVEPDIEQRR | ||||
| REFDAVLDFAQMMPMRV | ||||
| RYATDADVFAHDTLVDPYDTYRS | ||||
| gi|490036008 | 11,602 | EthD | R. ruber BKS 20-38 | KLALKQQGIRN |
| RLVAAYNHPEDPEKF | ||||
| RLVAAYNHPEDPEKFLDHYRN |
Cloning and expression of the ethABCD genes and ethR.
The amino acid sequences from peptide fingerprint analysis were used to design primers to amplify the eth genes from the genome of strain T3-1. The ethA, ethB, ethC, ethD, and ethABCD genes were amplified by each primer as described previously. The cloned ethABCD gene cluster was 3,219 bp in length, encoding a ferredoxin reductase, EthA; a cytochrome P-450 monooxygenase, EthB; a ferredoxin, EthC; and EthD, with unknown function (17–19). The corresponding proteins were searched against the GenBank database using the BLASTP program (http://blast.ncbi.nlm.nih.gov). The searches revealed that the most closely related proteins were a ferredoxin reductase, EthA (99% identity); a cytochrome P-450, EthB (98% identity); a ferredoxin, EthC (97% identity); and a ferredoxin, EthD (96% identity), from R. ruber (Fig. 3). ethR was cloned by SEFA-PCR, and its amino acid sequence shared 99% identity with that of EthR from R. ruber with an A12V site mutation.
FIG 3.
Phylogenetic tree of EthABCD and related proteins constructed by the neighbor-joining method. The branches corresponding to partitions reproduced in less than 50% of bootstrap replicates are collapsed. The strain names are shown. The scale bar indicates amino acid residue substitutions per amino acid position.
ethA, ethB, ethC, ethD, and ethABCD were cloned into the expression vector pET29a to verify their biochemical functions. No activities could be detected for the lysate of E. coli BL21(DE3) expressing ethABCD or the mixture of lysates of ethA, ethB, ethC, and ethD. Their expression in E. coli BL21(DE3) was analyzed by SDS-PAGE (data not shown). No evident expression of the cloned genes was observed in these E. coli strains.
Acetochlor transformation product analysis with resting E. coli cells expressing ethABCD and ethABD.
Although acetochlor degradation activity could not be detected with E. coli lysates, resting cells expressing ethABCD showed weak activities to acetochlor. A new product peak appeared at 4.959 min. The metabolite was identified as CMEPA by comparing its retention time (Rt) (4.959 min) from HPLC analysis and its charge-to-mass ratio (m/z) (212) from LC-MS analysis with those of an authentic CMEPA standard (Fig. 4b). Considering that EthC was not detected in the peptide mass fingerprint analysis of the purified enzyme, subclones of ethABC and ethABD were constructed to identify the function of EthC with EthD in acetochlor N-deethoxymethylation. The results revealed that the activity of N-deethoxymethylase, which converts acetochlor to CMEPA, was detected in the case of ethABD expressed in E. coli BL21(DE3) (Fig. 4a), indicating that EthD played an important role in acetochlor N-deethoxymethylation.
FIG 4.
Product analysis of reactions using whole cells of the transformed E. coli strain. (a) HPLC analysis of the acetochlor conversion product for E. coli BL21(DE3) [pET-29a(+)-ethABD]. (b) GC-MS analysis of the acetochlor conversion product for E. coli BL21(DE3) [pET-29a(+)-ethABD].
PCR and RT-PCR analyses of Rhodococcus sp. T3-1 and T3-1 MT.
In order to clarify the reason why T3-1 MT lost the ability to transform acetochlor, the genes in the eth cluster in the mutant were amplified by PCR. The result showed that none of the eth genes could be amplified from the mutant (data not shown). This indicated that the eth genes were lost in the mutant strain. Expression of the eth gene cluster in T3-1 was analyzed by reverse transcription (RT)-PCR with cells grown under different conditions. The results showed that eth genes were expressed under both induced (with acetochlor as the sole carbon source) and uninduced (with glucose as the sole carbon source) conditions (Fig. 5). This showed that the enzyme could be purified from cultures in LB broth without addition of acetochlor.
FIG 5.
Agarose gel electrophoresis of fragments of ethA, ethB, ethC, and ethD of Rhodococcus sp. T3-1 by RNA reverse transcription-PCR. Lane 1, ethA, glucose; lane 2, ethA, acetochlor; lane 3, ethB, glucose; lane 4, ethB, acetochlor; lane M, DL 5000 marker; lane 5, ethC, glucose; lane 6, ethC, acetochlor; lane 7, ethD, glucose; lane 8, ethD, acetochlor.
DISCUSSION
Rhodococcus sp. T3-1 could grow in MSM with acetochlor as the sole carbon source. Strain T3-1 showed high catalysis activity to convert acetochlor to CMEPA using whole cells (1), but no N-deethoxymethylase activity could be detected with the crude enzyme prepared by ultrasonic fragmentation. We tested cell disruption methods by protoplast lysis and grinding in liquid nitrogen. Crude cell extraction by grinding in liquid nitrogen showed weak activity to acetochlor with a special reaction buffer B; 40% sucrose and 0.1 M NaCl in the buffer played key roles in condensing the concentration and maintaining the stability of the enzyme, and β-mercaptoethanol prevented the free cysteine thiol group in the protein from forming a disulfide bond. Under optimized conditions, a multicomponent acetochlor N-deethoxymethylase was purified from Rhodococcus sp. T3-1. This N-deethoxymethylase was encoded by the ethABCD gene cluster involved in methyl tert-butyl ether (MTBE) and ETBE degradation (16–24). The functions of EthABCD in bacteria were genetically verified by gene complementation in mutant strains.
It was reported that a three-component Rieske nonheme iron oxygenase (RO) system catalyzed the N-dealkylation of acetochlor in Sphingomonas sp. strains DC-6 and DC-2 (25). Such an RO involved in the dicamba (3,6-dichloro-2-methoxybenzoic acid) O-demethylation was composed of a reductase, a ferredoxin, and an oxygenase (10, 26, 27). The structures of the oxygenases of the RO systems were highly conserved (28). However, in Rhodococcus sp. T3-1, the N-ethoxymethylation was accomplished by a cytochrome P450 system. Cytochrome P450 monooxygenase systems are versatile biocatalysts that introduce oxygen into a vast range of molecules (29). Research during the last few decades revealed that cytochrome P450 monooxygenases could metabolize herbicides and other xenobiotics (30). Cytochrome P450 was always associated with the electron transport component, which consisted of a cytochrome P450 reductase, a ferredoxin, and a cytochrome P450 monooxygenase. Cytochrome P450 could perform the function of oxidation only when it constituted a multienzyme complex together with oxidoreductase and ferredoxin (17). We deduced that the interaction among the subunits of the P450 complex is weak. Violent cell disruption could dissociate the complex and cause it to lose enzyme activity.
Many strains harboring the ethRABCD gene cluster have been reported, and its function was also studied by genetic methods (16–24). The ethABCD cluster encoded a ferredoxin reductase (EthA), a cytochrome P450 (EthB), a ferredoxin (EthC), and a protein with an unknown function (EthD) (17). ethR encoded a regulatory protein of the AraC/XylS family, and in strains containing ethR, the expression of ethB and ethD was induced by ETBE (17). Constitutive expression of the eth cluster in Rhodococcus sp. T3-1 was verified by examination of the transcription of eth genes with RT-PCR (Fig. 5). This is contrary to what was observed in R. ruber (17), although their regulatory proteins, EthR, shared very high identity (>99%). In the Aquincola tertiaricarbonis L108 strain with ethR lost, EthA, EthB, EthC, and EthD were expressed without induction (18, 19). We deduced that other regulation factors controlling the expression of the eth operon exist or that the A12V mutation led to the loss of function. The eth gene cluster could be spontaneously deleted in MTBE- or ETBE-degrading bacteria (17, 19). Strain T3-1 lost its ability to degrade acetochlor after repeated transfers, indicating a similar transposon structure around the eth gene cluster in this strain.
In the process of MTBE degradation, the reductase (EthA), the ferredoxin (EthC), and the cytochrome P450 (EthB) constituted an electron transport chain. EthD was considered not to be involved in the reaction (18), though gene complementation showed EthD was essential for MTBE degradation by R. ruber IFP 2001 (17). However, the function of EthD is unknown. The amino acid sequence of EthD is similar to that of a protein with an unknown function (Genpept accession no. U17130_4; 40% identity) that is encoded by the orf4 gene from Rhodococcus erythropolis. The orf4 gene is located in a gene cluster, thcRBCD, that also encodes a cytochrome P450 system, ThcD-ThcC-ThcB, catalyzing the N-dealkylation reaction of thiocarbamate compounds (17, 31). Our results of heterologous expression in E. coli showed that EthC was not required for acetochlor degradation while EthD was essential. Considering the catalytic mechanism of the P450 system, EthD might be a novel type of ferredoxin in the electron transport chain.
The cytochrome P450 monooxygenase EthABCD catalyzes hydroxylation of methoxy and ethoxy residues in the fuel oxygenates MTBE, ETBE, and tert-amyl methyl ether (TAME). In the reaction process of ETBE hydroxylation, the electron transport chain, consisting of EthA-EthC-EthB, converted the methoxy to the unstable intermediate TBF, and then the TBF was hydrolyzed to TBA and formaldehyde (32). Our results showed that EthABD could also perform N-dealkylation. This expanded the substrate of the Eth P-450 system.
The low-level expression of ethABCD in E. coli BL21(DE3) posed a difficulty in obtaining a sufficient amount of EthABD for biochemical analysis. Further research to improve the expression of ethABCD in E. coli is needed to clarify the biochemical character of the novel N-dealkylation function of the Eth P-450 system.
ACKNOWLEDGMENTS
Grants from the Natural Science Foundation of Jiangsu Province, China (no. BK2012029); the Natural Science Foundation of China (31270095); and the National Science and Technology Support Program (2012BAD14B02) supported this work.
REFERENCES
- 1.Hou Y, Dong W, Wang F, Li J, Shen W, Li Y, Cui Z. 2014. Degradation of acetochlor by a bacterial consortium of Rhodococcus sp. T3-1, Delftia sp. T3-6, and Sphingobium sp. MEA3-1. Lett Appl Microbiol 59:35–42. doi: 10.1111/lam.12242. [DOI] [PubMed] [Google Scholar]
- 2.Xiao N, Jing B, Ge F, Liu X. 2006. The fate of herbicide acetochlor and its toxicity to Eisenia fetida under laboratory conditions. Chemosphere 62:1366–1373. doi: 10.1016/j.chemosphere.2005.07.043. [DOI] [PubMed] [Google Scholar]
- 3.Li Y, Chen Q, Wang C-H, Cai S, He J, Huang X, Li S-P. 2013. Degradation of acetochlor by consortium of two bacterial strains and cloning of a novel amidase gene involved in acetochlor-degrading pathway. Bioresour Technol 148:628–631. doi: 10.1016/j.biortech.2013.09.038. [DOI] [PubMed] [Google Scholar]
- 4.Jablonkai I. 2000. Microbial and photolytic degradation of the herbicide acetochlor. Int J Environ Anal Chem 78:1–8. doi: 10.1080/03067310008032688. [DOI] [Google Scholar]
- 5.Dagnac T, Jeannot R, Mouvet C, Baran N. 2002. Determination of oxanilic and sulfonic acid metabolites of acetochlor in soils by liquid chromatography-electrospray ionisation mass spectrometry. J Chromatogr A 957:69–77. doi: 10.1016/S0021-9673(02)00310-2. [DOI] [PubMed] [Google Scholar]
- 6.de Bruijn FJ. 1992. Use of repetitive (repetitive extragenic palindromic and enterobacterial repetitive intergeneric consensus) sequences and the polymerase chain reaction to fingerprint the genomes of Rhizobium meliloti isolates and other soil bacteria. Appl Environ Microbiol 58:2180–2187. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7.Bradford MM. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72:248–254. doi: 10.1016/0003-2697(76)90527-3. [DOI] [PubMed] [Google Scholar]
- 8.Wang F, Hou Y, Zhou J, Li Z, Huang Y, Cui Z. 2014. Purification of an amide hydrolase DamH from Delftia sp. T3-6 and its gene cloning, expression, and biochemical characterization. Appl Microbiol Biotechnol 98:7491–7499. doi: 10.1007/s00253-014-5710-y. [DOI] [PubMed] [Google Scholar]
- 9.El-Dib MA, Abdel-Rahman MO, Aly OA. 1975. 4-Aminoantipyrine as a chromogenic agent for aromatic amine determination in natural water. Water Res 9:513–516. doi: 10.1016/0043-1354(75)90076-7. [DOI] [Google Scholar]
- 10.Fox B, Froland W, Dege J, Lipscomb J. 1989. Methane monooxygenase from Methylosinus trichosporium OB3b. Purification and properties of a three-component system with high specific activity from a type II methanotroph. J Biol Chem 264:10023–10033. [PubMed] [Google Scholar]
- 11.Boden R, Borodina E, Wood AP, Kelly DP, Murrell JC, Schäfer H. 2011. Purification and characterization of dimethylsulfide monooxygenase from Hyphomicrobium sulfonivorans. J Bacteriol 193:1250–1258. doi: 10.1128/JB.00977-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 12.Laemmli UK. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685. doi: 10.1038/227680a0. [DOI] [PubMed] [Google Scholar]
- 13.Sambrook J, Russell D. 2001. Molecular cloning: a laboratory manual, 3rd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. [Google Scholar]
- 14.Wang S, He J, Cui Z, Li S. 2007. Self-formed adaptor PCR: a simple and efficient method for chromosome walking. Appl Environ Microbiol 73:5048–5051. doi: 10.1128/AEM.02973-06. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15.Zhang J, Zheng J-W, Liang B, Wang C-H, Cai S, Ni Y-Y, He J, Li S-P. 2011. Biodegradation of chloroacetamide herbicides by Paracoccus sp. FLY-8 in vitro. J Agric Food Chem 59:4614–4621. doi: 10.1021/jf104695g. [DOI] [PubMed] [Google Scholar]
- 16.Jechalke S, Rosell M, Martínez-Lavanchy PM, Pérez-Leiva P, Rohwerder T, Vogt C, Richnow HH. 2011. Linking low-level stable isotope fractionation to expression of the cytochrome P450 monooxygenase-encoding ethB gene for elucidation of methyl tert-butyl ether biodegradation in aerated treatment pond systems. Appl Environ Microbiol 77:1086–1096. doi: 10.1128/AEM.01698-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17.Chauvaux S, Chevalier F, Le Dantec C, Fayolle F, Miras I, Kunst F, Beguin P. 2001. Cloning of a genetically unstable cytochrome P-450 gene cluster involved in degradation of the pollutant ethyltert-butyl ether by Rhodococcus ruber. J Bacteriol 183:6551–6557. doi: 10.1128/JB.183.22.6551-6557.2001. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Lopes Ferreira N, Malandain C, Fayolle-Guichard F. 2006. Enzymes and genes involved in the aerobic biodegradation of methyl tert-butyl ether (MTBE). Appl Microbiol Biotechnol 72:252–262. doi: 10.1007/s00253-006-0494-3. [DOI] [PubMed] [Google Scholar]
- 19.Schuster J, Purswani J, Breuer U, Pozo C, Harms H, Müller RH, Rohwerder T. 2013. Constitutive expression of the cytochrome P450 EthABCD monooxygenase system enables degradation of synthetic dialkyl ethers in Aquincola tertiaricarbonis L108. Appl Environ Microbiol 79:2321–2327. doi: 10.1128/AEM.03348-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20.François A, Mathis H, Godefroy D, Piveteau P, Fayolle F, Monot F. 2002. Biodegradation of methyl tert-butyl ether and other fuel oxygenates by a new strain, Mycobacterium austroafricanum IFP 2012. Appl Environ Microbiol 68:2754–2762. doi: 10.1128/AEM.68.6.2754-2762.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 21.Malandain C, Fayolle-Guichard F, Vogel TM. 2010. Cytochromes P450-mediated degradation of fuel oxygenates by environmental isolates. FEMS Microbiol Ecol 72:289–296. doi: 10.1111/j.1574-6941.2010.00847.x. [DOI] [PubMed] [Google Scholar]
- 22.Auffret M, Labbé D, Thouand G, Greer CW, Fayolle-Guichard F. 2009. Degradation of a mixture of hydrocarbons, gasoline, and diesel oil additives by Rhodococcus aetherivorans and Rhodococcus wratislaviensis. Appl Environ Microbiol 75:7774–7782. doi: 10.1128/AEM.01117-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 23.Le Digabel Y, Demanèche S, Benoit Y, Vogel TM, Fayolle-Guichard F. 2013. Ethyl tert-butyl ether (ETBE) biodegradation by a syntrophic association of Rhodococcus sp. IFP 2042 and Bradyrhizobium sp. IFP 2049 isolated from a polluted aquifer. Appl Microbiol Biotechnol 97:10531–10539. doi: 10.1007/s00253-013-4803-3. [DOI] [PubMed] [Google Scholar]
- 24.Liu H, Yan J, Wang Q, Karlson UG, Zou G, Yuan Z. 2009. Biodegradation of methyl tert-butyl ether by enriched bacterial culture. Curr Microbiol 59:30–34. doi: 10.1007/s00284-009-9391-1. [DOI] [PubMed] [Google Scholar]
- 25.Chen Q, Wang C-H, Deng S-K, Wu Y-D, Li Y, Yao L, Jiang J-D, Yan X, He J, Li S-P. 2014. Novel three-component Rieske non-heme iron oxygenase system catalyzing the N-dealkylation of chloroacetanilide herbicides in Sphingomonads DC-6 and DC-2. Appl Environ Microbiol 80:5078–5085. doi: 10.1128/AEM.00659-14. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 26.Wang X, Li B, Herman PL, Weeks DP. 1997. A three-component enzyme system catalyzes the O demethylation of the herbicide dicamba in Pseudomonas maltophilia DI-6. Appl Environ Microbiol 63:1623–1626. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 27.Herman PL, Behrens M, Chakraborty S, Chrastil BM, Barycki J, Weeks DP. 2005. A three-component dicamba O-demethylase from Pseudomonas maltophilia, strain DI-6: gene isolation, characterization, and heterologous expression. J Biol Chem 280:24759–24767. doi: 10.1074/jbc.M500597200. [DOI] [PubMed] [Google Scholar]
- 28.Dumitru R, Jiang WZ, Weeks DP, Wilson MA. 2009. Crystal structure of dicamba monooxygenase: a Rieske nonheme oxygenase that catalyzes oxidative demethylation. J Mol Biol 392:498–510. doi: 10.1016/j.jmb.2009.07.021. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 29.Urlacher VB, Eiben S. 2006. Cytochrome P450 monooxygenases: perspectives for synthetic application. Trends Biotechnol 24:324–330. doi: 10.1016/j.tibtech.2006.05.002. [DOI] [PubMed] [Google Scholar]
- 30.Kreuz K, Tommasini R, Martinoia E. 1996. Old enzymes for a new job (herbicide detoxification in plants). Plant Physiol 111:349. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 31.Nagy I, Schoofs G, Compernolle F, Proost P, Vanderleyden J, De Mot R. 1995. Degradation of the thiocarbamate herbicide EPTC (S-ethyl dipropylcarbamothioate) and biosafening by Rhodococcus sp. strain NI86/21 involve an inducible cytochrome P-450 system and aldehyde dehydrogenase. J Bacteriol 177:676–687. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 32.Zhang R-L, Huang G-Q, Lian J-Y, Li X-G. 2007. Degradation of MTBE and TEA by a new isolate from MTBE-contaminated soil. J Environ Sci 19:1120–1124. doi: 10.1016/S1001-0742(07)60182-X. [DOI] [PubMed] [Google Scholar]