The Two-Component Monooxygenase MeaXY Initiates the Downstream Pathway of Chloroacetanilide Herbicide Catabolism in Sphingomonads

ABSTRACT

Due to the extensive use of chloroacetanilide herbicides over the past 60 years, bacteria have evolved catabolic pathways to mineralize these compounds. In the upstream catabolic pathway, chloroacetanilide herbicides are transformed into the two common metabolites 2-methyl-6-ethylaniline (MEA) and 2,6-diethylaniline (DEA) through N-dealkylation and amide hydrolysis. The pathway downstream of MEA is initiated by the hydroxylation of aromatic rings, followed by its conversion to a substrate for ring cleavage after several steps. Most of the key genes in the pathway have been identified. However, the genes involved in the initial hydroxylation step of MEA are still unknown. As a special aniline derivative, MEA cannot be transformed by the aniline dioxygenases that have been characterized. Sphingobium baderi DE-13 can completely degrade MEA and use it as a sole carbon source for growth. In this work, an MEA degradation-deficient mutant of S. baderi DE-13 was isolated. MEA catabolism genes were predicted through comparative genomic analysis. The results of genetic complementation and heterologous expression demonstrated that the products of meaX and meaY are responsible for the initial step of MEA degradation in S. baderi DE-13. MeaXY is a two-component flavoprotein monooxygenase system that catalyzes the hydroxylation of MEA and DEA using NADH and flavin mononucleotide (FMN) as cofactors. Nuclear magnetic resonance (NMR) analysis confirmed that MeaXY hydroxylates MEA and DEA at the para-position. Transcription of meaX was enhanced remarkably upon induction of MEA or DEA in S. baderi DE-13. Additionally, meaX and meaY were highly conserved among other MEA-degrading sphingomonads. This study fills a gap in our knowledge of the biochemical pathway that carries out mineralization of chloroacetanilide herbicides in sphingomonads.

IMPORTANCE Much attention has been paid to the environmental fate of chloroacetanilide herbicides used for the past 60 years. Microbial degradation is considered an important mechanism in the degradation of these compounds. Bacterial degradation of chloroacetanilide herbicides has been investigated in many recent studies. Pure cultures or consortia able to mineralize these herbicides have been obtained. The catabolic pathway has been proposed, and most key genes involved have been identified. However, the genes responsible for the initiation step (from MEA to hydroxylated MEA or from DEA to hydroxylated DEA) of the downstream pathway have not been reported. The present study demonstrates that a two-component flavin-dependent monooxygenase system, MeaXY, catalyzes the para-hydroxylation of MEA or DEA in sphingomonads. Therefore, this work finds a missing link in the biochemical pathway that carries out the mineralization of chloroacetanilide herbicides in sphingomonads. Additionally, the results expand our understanding of the degradation of a special kind of aniline derivative.

INTRODUCTION

Since their invention in the 1960s, chloroacetanilide herbicides have become one of the most important and widely used classes of herbicides, which are applied to corn, cotton, soybean, and other crops for the control of annual grass and broadleaf weeds (1). Commonly used chloroacetanilide herbicides, such as alachlor, acetochlor, butachlor, and metolachlor, are N-alkoxyalkyl-N-chloroacetyl-substituted aniline derivatives (Fig. 1A). Due to their widespread use and relatively high water solubility, some chloroacetanilide herbicides and their metabolites have been frequently detected in soil and groundwater (2). Ecotoxicological data have suggested that some chloroacetanilide herbicides can cause DNA damage (3, 4) and tumor induction in fish, rat, and human cells (5–7). Alachlor, acetochlor, butachlor, and metolachlor are listed as B2, B2, L2, and C classes of carcinogens by the U.S. Environmental Protection Agency, respectively (8–10). Thus, much attention has been paid to the environmental fate of chloroacetanilide herbicides.

FIG 1.

The proposed bacterial catabolic pathway of chloroacetanilide herbicides. (A) Molecular structures of chloroacetanilide herbicides and their metabolites. (B) Proposed acetochlor-catabolic pathway and the enzymes involved. The upstream pathway of acetochlor degradation is shown in blue, and the downstream pathway is shown in green. The reaction investigated in this study is shown in red and indicated by a question mark.

Microbial degradation plays an important role in the elimination of chloroacetanilide herbicides in the environment (2). Enrichment culture techniques have been used with varied success in attempts to enrich and isolate chloroacetanilide herbicide-degrading microorganisms. A variety of bacterial strains able to degrade butachlor, alachlor, acetochlor, and metolachlor have been reported (11–18). Furthermore, some bacterial pure cultures (2) or consortia (15, 16) are able to mineralize these herbicides. Sphingomonas wittichii DC-6 was reported to mineralize chloroacetanilide herbicides, such as alachlor, acetochlor, and butachlor (2). In the consortium described by Li et al. (15), Sphingobium quisquiliarum DC-2 could only transform acetochlor to 2-methyl-6-ethylaniline (MEA), which was mineralized by Sphingobium baderi DE-13. In a consortium named T3, three bacteria could completely mineralize acetochlor via cooperative metabolism: acetochlor to 2-methyl-6′-ethyl-2-chloroacetanilide (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 (16). In addition, strains DC-6, DC-2, and DE-13 were isolated from Kunshan City, China, while consortium T3 was enriched from the sample collected in Zhenjiang City, China.

Based on the similar metabolites detected, a common bacterial catabolic pathway for chloroacetanilide herbicides has been proposed (Fig. 1B). Using acetochlor as an example, in the upstream pathway, initial N-dealkylation is followed by amide hydrolysis to generate MEA. In the downstream pathway, MEA was hydroxylated and hydrolytically deaminated to 2-methyl-6-ethyl-hydroquinone (MEHQ); MEHQ was further hydroxylated to 3-hydroxy-2-methyl-6-ethyl-hydroquinone (3-OH-MEHQ), which is the putative substrate of ring cleavage (2, 15, 18–20). Regarding the genetic basis of this pathway, a three-component Rieske nonheme iron oxygenase system, CndABC, or a cytochrome P450 system EthBAD responsible for initial N-dealkylation was identified from S. quisquiliarum DC-2 and Rhodococcus sp. strain T3-1, respectively (2, 20). The amidases CmeH and DamH in charge of amide hydrolysis were characterized in the S. quisquiliarum DC-2 and Delftia sp. strain T3-6, respectively (15, 19). The flavin-dependent monooxygenase system MeaBA able to transform MEHQ to 3-OH-MEHQ was identified in Sphingobium sp. strain MEA3-1 (18). However, the genes responsible for the hydroxylation of MEA are still unknown. In addition, the alkyl substituents of MEA are located on both sides of the amine group. This structure not only makes MEA very stable in natural environments (21) but also distinguishes its degradation pattern from the typical aniline catabolic pathway in bacteria. In a common degradation pathway, aniline is initially converted to catechol by aniline dioxygenases (22–24). However, MEA degradation is initiated with hydroxylation at the para-position (2, 15, 18), which may be catalyzed by a monooxygenase.

This study aims to identify the genes responsible for the initial hydroxylation step of MEA degradation in S. baderi DE-13. Given the unstable MEA-degrading phenotype of S. baderi DE-13, the following strategy was used in this work: a mutant deficient in MEA degradation was first isolated, whose genome sequence was compared with the wild type to predict candidate genes involved in MEA degradation; then, the candidate genes were subjected to experimental validation to confirm their specific function. Furthermore, the transcriptional response of identified genes to MEA and 2,6-diethylaniline (DEA) were also investigated. This study fills a gap in our knowledge of the biochemical pathway that mineralizes chloroacetanilide herbicides in sphingomonads and expands our understanding of the degradation of certain aniline derivatives.

RESULTS AND DISCUSSION

Isolation of the MEA degradation-deficient mutant strain DE-13-E9.

S. baderi DE-13 could completely degrade MEA and use it as a sole carbon source for growth (Fig. 2A). However, this MEA-degrading phenotype was unstable and prone to loss without selective pressure from MEA. Through continuous transfer on LB medium, an MEA degradation-deficient mutant strain, DE-13-E9, was isolated. The DE-13-E9 mutant showed no degradation activity toward MEA (Fig. 2B). Furthermore, the cell extract of strain DE-13 could catalyze MEA to hydroxylated MEA with exogenous addition of 100 μM NADH, while the cell extracts of the DE-13-9 mutant showed no detectable activity toward MEA under the same condition. These results indicated that the DE-13-E9 mutant lost at least the genes responsible for the initial degradation step of MEA.

FIG 2.

Degradation of MEA by wild-type strain DE-13 (A), mutant DE-13-E9 (B), and genetic complementation strain DE-13-E9 (pBmeaX) (C). Degradation experiments were carried out in MSM medium containing 0.5 mM MEA. The initial cell densities were set to an OD₆₀₀ of ∼0.05. The concentration of MEA was detected by HPLC, as described in Materials and Methods. The degradation curve is shown in green, and the growth curve is shown in blue. Results are the averages from three independent experiments, and error bars show standard deviations.

Prediction of MEA-catabolic genes based on comparative genomic analysis.

To find the genes lost in this MEA degradation-deficient mutant, the complete genome of strain DE-13 and the draft genome of the mutant DE-13-E9 were sequenced. The complete genome (4.58 Mb) of strain DE-13 revealed nine replicons, consisting of one circular chromosome and eight circular plasmids. The draft genome sequence of the DE-13-E9 mutant contained 159 contigs, constituting a total size of 4.48 Mb. As shown in Fig. S1, pairwise comparison results showed that compared to the wild type, the DE-13-E9 mutant lost a 14-kb fragment, which was named F-E9. The fragment is located on the chromosome of strain DE-13 from bp 157030 to 171030.

The candidate genes involved in the degradation of MEA were then predicted in fragment F-E9. The complete physical map of F-E9 is presented in Fig. 3. Computational analysis of this fragment identified 13 open reading frames (ORFs), and the proposed function for each ORF is listed in Table 1. Among these ORFs, one designated meaX was homologous to the genes encoding the oxygenase component of the two-component flavoprotein monooxygenase system (Fig. S2 and Table 1) and is likely responsible for the hydroxylation of MEA. MeaX consists of 406 amino acid residues and shares 30% and 24.8% identities with HsaA from Rhodococcus sp. strain RHA1 (25) and MeaA from Sphingobium sp. strain MEA3-1 (18), respectively. MeaX belongs to the group D flavin monooxygenases, which are enzymes with an acyl-coenzyme A (acyl-CoA) dehydrogenase fold (26). Large flavin mononucleotide (FMN)-specific reductase and small flavin reductase have been reported as the reductase components of group D flavin monooxygenases (27, 28).

FIG 3.

Physical map of the lost fragment in mutant DE-13-E9 and genetic complementation results. ORFs in orange are transposases or homologous proteins. Both meaX and meaY are drawn in red, and other ORFs are blue. Detailed information about these ORFs is listed in Table 1. The results of complementation of the mutant strain DE-13-E9 with various regions in different hosts (Sphingobium baderi DE-13-E9, Pseudomonas putida KT2440, Sphingomonas wittichii RW1, and Escherichia coli DH5α) are shown below the physical map. +, can degrade; −, cannot degrade.

TABLE 1.

Deduced function of each ORF identified within the lost fragment F-E9

Gene	Product sizea	Databaseb	Homologous proteinc	GenBank accession no.	Identity (%)	Proposed function
orf1	266	NR	Gammaproteobacteria transposase	WP_001375121	99	Transposase
		Swiss-Prot	Transposase for insertion sequence-like element IS431mec	P0A043	42
orf2	264	NR	Blastomonas sp. CACIA14H2 carnitine dehydratase	ESZ88780	69	Carnitine dehydratase
		Swiss-Prot	2-Methylacyl-CoA racemase	O09174	49
orf3	158	NR	Pseudomonas fluorescens NCIMB 11764 transcriptional regulator	AKV10929	63	Transcriptional regulator
		Swiss-Prot	Putative l-lactate dehydrogenase operon regulatory protein	P0ACL7	30
meaX	407	NR	Actinokineospora enzanensis hypothetical protein	WP_018684803	50	Oxygenase component of MEA monooxygenase
		Swiss-Prot	Flavin-dependent monooxygenase, oxygenase subunit HsaA	Q0S811	30
orf4	433	NR	Sphingomonas hypothetical protein	WP_048575098	67	Putative maleylacetate reductase I
		Swiss-Prot	Maleylacetate reductase I	P27137	29
orf5	464	NR	Novosphingobium barchaimii LL02 transposase IS5	KMS51534	100	Transposase
		Swiss-Prot	Probable beta-galactosidase B	B8NKI4	39
orf6	157	NR	Sphingomonas hypothetical protein	WP_048574811	100	Hypothetical protein
		Swiss-Prot	Ribosome-associated factor Y	Q5XAQ7	28
orf7	107	NR	Sphingomonas hypothetical protein	WP_048578651	100	Hypothetical protein
		Swiss-Prot	Leucyl-tRNA synthetase	Q7P0R1	27
meaY	173	NR	Sphingobium chungbukense hypothetical protein	WP_046764014	49	Reductase component of MEA monooxygenase
		Swiss-Prot	FMN reductase	A9CHR3	34
orf8	271	NR	Caulobacter henricii transposase	WP_035091777	84	Transposase
		Swiss-Prot	Putative transposase y4pF/y4sB	P55615	49
orf9	502	NR	Sphingomonas hypothetical protein	WP_030092541	99	Mobile element protein
		Swiss-Prot	Uncharacterized protein y4jA/y4nE/y4sE	P55501	67
orf10	329	NR	Novosphingobium sp. PP1Y ATPase AAA	WP_013832221	99	Mobile element protein
		Swiss-Prot	Putative insertion sequence ATP-binding protein y4iQ/y4nD/y4sD	P55500	76
orf11	541	NR	Sphingomonadaceae integrase	WP_030539573	100	Integrase
		Swiss-Prot	Uncharacterized protein y4rM	P55646	57

^aNumber of amino acids.

^bNR, NCBI Nonredundant Protein Sequences Database.

^cThe top BLASTP hit was selected.

The gene orf4, adjacent to meaX, was predicted to encode a 432-amino-acid protein with 29% identity to maleylacetate reductase I TfdFI from Ralstonia eutropha JMP134 (29). The gene located four ORFs from meaX, called meaY, was predicted to encode a protein of 173 amino acids that exhibits 34% identity with the FMN reductase RutF from Agrobacterium tumefaciens C58 (30). It was speculated that ORF4 or MeaY may be the reductase component of meaX.

As shown in Table 1, fragment F-E9 was flanked by the ORFs encoding transposases or other mobile element proteins (orf1, orf5, orf8, orf9, and orf10), which likely contributed to the instability of fragments F-E9 in strain DE-13. In addition, pairwise comparison results showed that the majority of fragment F-E9, including meaX, orf4, and meaY, was highly conserved (>95% similarities) in the draft genome sequences of MEA-degrading strains S. wittichii DC-6 (accession no. JMUB01000000) and Sphingobium sp. strain MEA3-1 (accession no. LECE01000000), indicating that the genes involved in MEA degradation could be horizontally transferred among these sphingomonads.

Genetic complementation and heterologous expression of MeaX in different hosts.

Genetic complementation was first used to identify the functions of meaX. As illustrated in Fig. 3, three fragments containing meaX-orf4, meaX, or orf4 were cloned into the plasmid pBBR1MCS-2. The resulting plasmids, pBmeaXorf4, pBmeaX, and pBorf4, were transferred into the DE-13-E9 mutant. Both plasmids pBmeaXorf4 and pBmeaX could restore the ability of the DE-13-E9 mutant to degrade and grow on MEA (Fig. 2C).

To further confirm the function of MeaX, plasmid pBmeaX, which contained meaX, was introduced into different bacterial strains, including S. wittichii RW1, P. putida KT2440, and Escherichia coli DH5α (Fig. 3). Except for E. coli DH5α, plasmid pBmeaX was able to confer the ability to transform MEA to all of the strains, producing a mixture of 4-OH-MEA, MEHQ, 2-methyl-6-ethyl-benzoquinone imine (MEBQI), and 2-methyl-6-ethyl-benzoquinone (MEBQ) (data not shown).

These results suggested that meaX was involved in the hydroxylation of MEA in strain DE-13. Moreover, MeaX did not have a high degree of specificity in the recognition of its partner flavin reductase, and the background flavin reductases in the tested hosts (strains DE-13, KT2440, and RW1), except E. coli DH5α, could support the activity of MeaX.

MeaXY catalyzes the monooxygenation of both MEA and DEA. (i) Expression and purification of MeaX and MeaY.

Recombinant MeaX and MeaY were individually overexpressed in E. coli BL21(DE3)/pG-KJE8 as N- and C-terminal 6His-tagged fusion proteins. SDS-PAGE analysis showed that the molecular masses of 6His-MeaX and MeaY-6His are approximately 46 and 21 kDa (Fig. S3), respectively, in line with the molecular weights deduced from their amino acid sequences.

(ii) Characterization of the reductase component of MEA/DEA monooxygenase.

ORF4 was also heterologously expressed and purified in E. coli BL21(DE3)/pG-KJE8, but the purified ORF4 (6His-ORF4 or ORF4-6His) did not have NAD(P)H oxidation activity in the presence of flavin adenine dinucleotide (FAD) or flavin mononucleotide (FMN), indicating that it is not the reductase component of MeaX (data not shown). On the other hand, in agreement with its predicted function, the enzyme assay indicated that MeaY could catalyze the oxidation of NADH, with an enzyme activity of 13.64 U · mg⁻¹, when FMN was used as the cofactor (defined as 100%). When FMN was replaced by FAD, the enzyme activity decreased to 4.45%, and when NADH was replaced by NADPH, the activity decreased to only 2.45%. These results showed that NADH was the favored electron donor and FMN was the favored electron acceptor for MeaY. The kinetic parameter results revealed K_m values for MeaY-6His toward NADH and FMN of 38.62 ± 5.05 and 21.22 ± 3.16 μM, respectively. The corresponding k_cat/K_m values were 0.24 ± 0.015 and 0.66 ± 0.046 s⁻¹ · μM⁻¹, respectively (values are averages of three measurements ± standard deviations) (Table 2).

TABLE 2.

Kinetic parameters^a of recombinant 6His-MeaX and MeaY-6His

Enzymeb	Substrate	Cofactor(s)	Km (μM)	kcat (s−1)	kcat/Km (s−1 · μM−1)
MeaY	NADH	FMN	38.62 ± 5.05	9.09 ± 0.69	0.24 ± 0.015
	FMN	NADH	21.22 ± 3.16	13.94 ± 1.73	0.66 ± 0.046
MeaXY	MEA	NADH, FMN	0.43 ± 0.053	0.74 ± 0.061	1.72 ± 0.084
	DEA	NADH, FMN	0.53 ± 0.024	0.79 ± 0.066	1.49 ± 0.12

^aValues are means ± standard deviations calculated from triplicate assays.

^bMolar ratio of MeaX/MeaY is 20:1.

(iii) Characterization of MEA/DEA monooxygenase MeaXY.

As shown in Fig. S4, purified 6His-MeaX and MeaY-6His could transform MEA to 4-OH-MEA (and DEA to 4-OH-DEA). The optimal molar ratio of 6His-MeaX and MeaY-6His was determined. The amount of 6His-MeaX was constant at 0.25 μM, while MeaY-6His ranged from 0.005 to 1.25 μM. The MEA hydroxylation activity reached a maximum when MeaY-6His was added at 0.0125 μM, which meant that MeaX-to-MeaY ratio of 1:20 was the optimal molar ratio (Fig. 4). The steady-state kinetics of the oxidase component against MEA and DEA were also evaluated. As shown in Table 2, the K_m values for MEA and DEA were 0.43 ± 0.053 and 0.53 ± 0.024 μM, respectively, and the corresponding values of k_cat/K_m were 1.72 ± 0.084 and 1.49 ± 0.12 s⁻¹ · μM⁻¹, respectively. These data indicated that MeaXY has similar affinity and catalytic activity for the two aniline derivatives (MEA and DEA).

FIG 4.

Relationship between the MEA monooxygenase activity and various concentrations of MeaY-6His while the concentration of 6His-MeaX was kept constant. The concentration of MeaY-6His was increased from 0.005 to 1.25 μM when the concentration of 6His-MeaX was kept constant at 0.25 μM. The 6His-MeaX specific activity against MEA was defined as 100% when 0.0125 μM MeaY-6His was added. Data were calculated from three independent replicates, and error bars indicate standard deviations.

MeaXY hydroxylates the aromatic rings of MEA and DEA at the para-position.

Previous studies have proposed that the aromatic ring of MEA is initially hydroxylated at the para-position in strain DE-13 (15) and Sphingobium sp. MEA3-1 (18). To verify this prediction, MeaXY-hydroxylated DEA and MEA were purified using silica gel column chromatography and subjected to nuclear magnetic resonance (NMR) analysis. As shown in Table 3, the spectrum of hydroxylated DEA was as follows: ¹H NMR (500 MHz, dimethyl sulfoxide-d6 [DMSO-d₆]) δ 8.21 (s, ¹H), 6.29 (s, 2H), 3.88 (s, 2H), 2.39 (dd, J = 7.5 Hz, 4H), 1.09 (t, J = 7.5 Hz, 6H). δ 6.29 ppm, 2.39 ppm, and 1.09 ppm have a peak area ratio of 2:4:6, which means that two benzene hydrogens share the same chemical environment as hydrogens of the two ethyl groups. Therefore, the only possible position for the hydroxyl was the para-position of the amido group. The ¹H NMR results for the hydroxylated MEA were: ¹H NMR (500 MHz, DMSO-d₆) δ 8.19 (s, 1H), 6.28 (s, 2H), 3.91 to 3.87 (m, 2H), 2.39 (dd, J = 7.5 Hz, 2H), 2.01 (s, 3H), 1.09 (t, J = 7.5 Hz, 3H). The δ 6.2821 ppm signal represents the two hydrogens of benzene, i.e., these two hydrogens share the same chemical environment in the NMR spectrum. Although anisomerous methyl and ethyl could theoretically place the two benzene hydrogens in different chemical environments, the hydroxyl plays the main role in the alteration of chemical shift. The hydroxyl alters the chemical shift of the ortho-position benzene hydrogen much more than the meta-position benzene hydrogen. Therefore, if hydroxyl was at the 3′ or 5′ position, the chemical shifts of the two benzene hydrogens would differ by approximately 0.5 ppm. Only when the hydroxyl group was at the para-position relative to the amido group could two benzene hydrogens share the same chemical shift values. These data showed that MeaXY hydroxylates the aromatic rings of MEA and DEA at the para-position, which is consistent with the reported tandem mass spectrometry (MS-MS) results in S. baderi DE-13 (15) and Sphingobium sp. strain MEA3-1 (18). Considering that meaX and meaY are highly conserved among strains DE-13, DC-6 (2), and MEA3-1 (18), it is reasonable to conclude that para-hydroxylation is the common catabolic step of MEA and DEA in these sphingomonads.

TABLE 3.

¹H NMR data analysis of hydroxylated MEA and DEA

Transcription levels of meaX and meaY in response to MEA or DEA.

The relative changes in the transcription of meaX and meaY under MEA- or DEA-induced and noninduced conditions were investigated using reverse transcription-quantitative PCR (RT-qPCR). As shown in Fig. 5, both MEA and DEA could upregulate the transcription of meaX and its adjacent gene, orf4. The increased range was similar between meaX and orf4: MEA upregulated 94.5-fold relative to meaX and 89.3-fold relative to orf4; DEA upregulated 60.8-fold relative to meaX and 52.6-fold relative to orf4. However, MEA and DEA had no effects on the transcription of meaY (Fig. 5). A FadR-type transcription regulator encoded by orf3 was predicted just upstream of meaX (Fig. 3 and Table 1) and is probably involved in the transcriptional regulation of meaX.

FIG 5.

Relative transcription levels of meaX, orf4, and meaY in strain DE-13 under MEA and DEA induction. The culture in glucose was set as the control, and relative expression levels were calculated using the 2^−ΔΔCT threshold cycle (C_T) method (49). Data were collected from three independent determinations, and the error bars indicate standard deviations.

Conclusion.

Taken together, this work demonstrates that the products of meaX and meaY are responsible for the initial step of MEA degradation in strain DE-13. meaX and meaY encode a two-component flavoprotein monooxygenase system that catalyzes the para-hydroxylation of MEA and DEA using NADH and FMN as cofactors. Additionally, transcription of meaX was significantly enhanced in response to MEA or DEA in strain DE-13. meaX and meaY were highly conserved in the two other MEA-degrading sphingomonads S. wittichii DC-6 and Sphingobium sp. strain MEA3-1. Previous studies have indicated that a chloroacetanilide herbicide-mineralizing pathway has evolved in some sphingomonads (2, 15, 18). This work describes a missing link in the biochemical pathway that carries out mineralization of chloroacetanilide herbicides in sphingomonads. Identification of other genes in this pathway and determining how they are regulated will be carried out in the future.

MATERIALS AND METHODS

Chemicals.

MEA (98% purity) and DEA (98% purity), purchased from J&K Chemical (Shanghai, China), were prepared as 0.2 M stock solutions in methanol and sterilized by membrane filtration (0.22 μm). Dimethyl sulfoxide-d6 (DMSO-d6) was obtained from Merck (Darmstadt, Germany).

Bacterial strains, culture conditions, plasmids, and oligonucleotides.

The strains and plasmids used in this study are listed in Table 4. The oligonucleotides used in this study are listed in Table 5. All bacterial strains were grown in Luria-Bertani (LB) medium. E. coli strains were grown at 37°C, and other bacterial strains were grown aerobically at 30°C. The growth of cells was evaluated using a colony counting method. Degradation of MEA or DEA was evaluated in mineral salt medium (MSM), which contained (in grams per liter) NaCl, 1.0; NH₄NO₃, 1.0; K₂HPO₄, 1.5; KH₂PO₄, 0.5; and MgSO₄·7H₂O, 0.2 (pH 7.0). When required, the final concentrations of antibiotics were as follows: streptomycin (Str), 100 mg · liter⁻¹; ampicillin (Ap), 100 mg · liter⁻¹; kanamycin (Km), 50 mg · liter⁻¹; and chloramphenicol (Cm), 30 mg · liter⁻¹.

TABLE 4.

Strains and plasmids used in this study

Strain or plasmid	Characteristicsa	Source or reference
Strains
Sphingobium baderi
DE-13	Wild-type, able to degrade MEA and grow on it as sole carbon source, Smr	15
DE-13-E9	Mutant of strain DE-13, unable to degrade MEA and DEA, Smr	This study
Pseudomonas putida KT2440	Unable to degrade MEA and DEA, Cmr	43
Sphingomonas wittichii RW1	Dibenzo-p-dioxin-degrading strain; unable to degrade MEA and DEA, Smr	44
Escherichia coli
DH5α	F− φ80dlacZΔM15 Δ(lacZYA-argF)U169 recA1 endA1 thi-1 supE44 relA1 deoR	TaKaRa
HB101 (pRK600)	Conjugation helper strain, Cmr	45
BL21(DE3)/pG-KJE8	F− ompT hsdS(rB− mB−) gal dcm lacY1 (DE3)/pG-KJE8 (Cmr)	TaKaRa
Plasmids
pMD19-T	TA clone vector, Apr	TaKaRa
pET-28a(+)	Expression vector, Kmr	TaKaRa
pET-29a(+)	Expression vector, Kmr	TaKaRa
pG-KJE8	Chaperone plasmid, Cmr	TaKaRa
pBBR1MCS-2	Broad-host-range cloning vector, Kmr	42
pBmeaXorf4	pBBR1MCS-2 harboring meaX and orf4, Kmr	This study
pBmeaX	pBBR1MCS-2 harboring meaX, Kmr	This study
pBorf4	pBBR1MCS-2 harboring orf4, Kmr	This study
pET28meaX	pET-28a(+) harboring meaX, Kmr	This study
pET29meaY	pET-29a(+) harboring meaY, Kmr	This study
pET28orf4	pET-28a(+) harboring orf4, Kmr	This study
pET29orf4	pET-29a(+) harboring orf4, Kmr	This study

^aSm^r, streptomycin resistance; Cm^r, chloramphenicol resistance; Ap^r, ampicillin resistance; Km^r, kanamycin resistance.

TABLE 5.

Oligonucleotides used in this study

Primer	Sequence (5′ to 3′)a	Purpose
F-E9-F	TTTGGGAGCAAACCAGATCAGGACA	Confirmation of the lost fragment in DE-13-E9
F-E9-R	CTCCTCCTCGAACCACGCCTTCAAC
RTX-F	CACTCTACGAAGACGCGGCCTT	RT-qPCR of meaX
RTX-R	ATCACCAGTTCCGAGTATGGCATC
RT4-F	CGCCGTCTCTTGAGCCTACTTTCG	RT-qPCR of orf4
RT4-R	GGCCGCACCTCGCTGAACAC
RTY-F	TGTCAGACGTTCTCGAGCCTT	RT-qPCR of meaY
RTY-R	TTCTTCGGTCGCCATGTAGC
RT16S-F	CCCTGGTAGTCCACGCCGTA	RT-qPCR of 16S rRNA gene
RT16S-R	CTGTCACCTAGCCAGCCGAAC
meaX-CF	AAGCTTCTCCAGGGCGTACATGGCTTCCCAGA (HindIII)	Complementation of meaX
meaX-CR	TCTAGACCGGCATATCGCAAGCATCCAGGAAC (XbaI)
orf4-CF	AAGCTTCACCTAATACGCCGTTCCTTTCATCT (HindIII)	Complementation of orf4
orf4-CR	TCTAGATCCTCAAGGGTCTCGACGGCGACATC (XbaI)
meaX-CF	AAGCTTCTCCAGGGCGTACATGGCTTCCCAGA (HindIII)	Complementation of meaX and orf4
orf4-CR	TCTAGATCCTCAAGGGTCTCGACGGCGACATC (XbaI)
meaX-EF	ATTACATATGACCCAAGCGATAGATGCGGGCGA (NdeI)	Construction of 6His-MeaX expression plasmid pET28meaX
meaX-ER	AGGTAAGCTTAAGATGAAAGGAACGGCGTATTAG (HindIII)
meaY-EF	GTAACATATGCAAAGGCCTGATGAGAATTCTA (NdeI)	Construction of MeaY-6His expression plasmid pET29meaX
meaY-ER	CTGTAAGCTTACGCTCAGTGCGGTCAGAGC (HindIII)
orf4-EF1	ATTACATATGGTGCGGGCCAACAGGACGGTT (NdeI)	Construction of 6His-ORF4 expression plasmid pET28orf4
orf4-ER1	AGGTAAGCTTAGTAGGCCATATTGAGAATCTCCA (HindIII)
orf4-EF2	CATATGGTGCGGGCCAACAGGACGGTT (NdeI)	Construction of ORF4-6His expression plasmid pET29orf4
orf4-ER2	CTCGAGGTAGGCCATATTGAGAATCTC (XhoI)

^aRestriction enzyme cutting sites are underlined (with the restriction enzyme in parentheses).

Isolation of spontaneous mutant strains.

After several continuous transfers in LB broth, the cells of strain DE-13 were spread on LB plates. Each colony was transferred to LB plates and MSM plates containing 0.5 mM MEA. Mutants that failed to grow on MEA were further evaluated as described below. An MEA degradation-deficient mutant strain was designated DE-13-E9.

Degradation test.

The tested cells growing in LB broth were washed twice with MSM medium and suspended in MSM medium containing 0.5 mM MEA. For strain DE-13 and its derivative, the final optical density at 600 nm (OD₆₀₀) was adjusted to approximately 0.05. For other strains, the final OD₆₀₀ was set at approximately 2.0. The substrate and its metabolites were extracted by dichloromethane. Anhydrous sodium sulfate was used to dehydrate the organic phase. For qualitative analysis, UV absorption of the solution was detected by a UV scanner UV-2450 (Shimadzu, Japan) at wavelengths from 200 to 350 nm. For quantitative analysis, the solvent dichloromethane was removed by stream of nitrogen gas, and the extracts were redissolved in 1 ml of methanol. Prepared samples were filtered through a 0.22-μm-pore Millipore membrane and analyzed using high-performance liquid chromatography (HPLC). Analysis was performed as follows: the separation column was a Shim-pack VP-ODS C₁₈ (250 mm by 4.6 mm), the mobile phase was methanol/water at a 80:20 (vol/vol) ratio with a flow rate of 0.8 ml · min⁻¹, and an SPD-20A detector was used to measuring UV absorption at 230 nm and 280 nm. The mass spectrum was collected using a Finnigan TSQ Quantum Ultra AM thermal triple-quadrupole mass spectrometer. The metabolites were ionized by electrospray with positive polarity, and characteristic fragment ions were detected using tandem mass spectrometry (MS-MS).

Genomic DNA sequencing analysis and PCR conditions.

The total genomic DNA was extracted according to the method of Pitcher et al. (31). Genome sequencing was performed by Majorbio Bio-Pharm Technology Co., Ltd. (Shanghai, China). The complete genome sequence of strain DE-13 was assembled using a method combining Illumina MiSeq sequencing technology, the Pacific Biosciences platform, and PCR validation (32–34). The draft genome sequence of strain DE-13-E9 was generated using the Illumina MiSeq platform. Annotation was performed using Glimmer 3.02 (35), tRNAscan-SE version 1.3.1 (36), and Barrnap 0.4.2 (37). BLASTN and BLASTP were used for the nucleotide sequence and deduced amino acid identity searches, respectively.

ORF comparison between the genomes of strain DE-13 and its mutant was performed using the RAST online analysis tool with the sequence-based model (38, 39). Lost fragments in the mutant strains were confirmed by PCR amplification and sequenced using primer sets F-E9-F/F-E9-R and F-D2-F/F-D2-R (Table 5). The lost fragments were compared to the genomes of strains DC-6 (2) and MEA3-1 (18) using dot plot analysis in OMIGA (40). All comparison results were visualized using the Circos software (41). In an ORF comparison between strains DE-13 and DE-13-E9, homologous genes whose amino acids shared >98% similarity were linked.

PCR amplification in this study was generally carried out as follows: the reaction solution contained 1× PCR buffer (50 mM KCl, 10 mM Tris-HCl, 0.1% Triton X-100, 2 mM MgSO₄ [pH 9.0]), approximately 0.1 ng of template DNA, 0.5 μM forward primer, 0.5 μM reverse primer, 0.2 mM dinucleoside triphosphate (dNTP), and 5 U of DNA polymerase (PrimeSTAR GXL DNA polymerase; TaKaRa, Japan) in 50 μl; amplification was performed in a Thermo Scientific Arktik cycler at 95°C 5 min for initial denaturation, 30 cycles of amplification (95°C for 10 s, 55 to 60°C for 5 s, 68°C for 1 min per 1 kb), 68°C 10 min for final extension, and storage at 4°C.

Genetic complementation and heterologous expression of MeaXY in different bacterial hosts.

Three fragments containing meaX-orf4, meaX, and orf4 were amplified using primer sets meaX-CF/orf4-CR, meaX-CF/meaX-CR, and orf4-CF/orf4-CR (Table 5) and cloned into the broad-host-range plasmid pBBR1MCS-2 (42) at the HindIII and XbaI sites, respectively. The sequences of inserted fragments in all constructed plasmids were verified through Sanger sequencing (3730xl DNA analyzer; Applied Biosystems). The plasmids pBBR1MCS-2, pBmeaXorf4, pBmeaX, and pBorf4 were introduced into strain DE-13-E9, Pseudomonas putida KT2440 (43), and Sphingomonas wittichii RW1 (44) by triparental conjugative transfer with the help of E. coli HB101 (pRK600) (45). Strains harboring genetic complementation plasmids were also confirmed by PCR and DNA sequencing. The MEA-degrading abilities of these strains were tested as described above.

Expression and purification of recombinant MeaX, MeaY, and ORF4.

The sequence of meaX was PCR amplified using the meaX-EF/meaX-ER primer set and cloned into the expression vector pET-28a(+) to give the expression plasmid pET28meaX. meaY was amplified using primer pair meaY-EF/meaY-ER and ligated into vector pET-29a(+), yielding plasmid pET29meaY. To express recombinant 6His-ORF4 and ORF4-6His, orf4 was amplified using the PCR primer sets orf4-EF1/orf4-ER1 and orf4-EF2/orf4-ER2, respectively. The amplified DNA fragments were cloned into the pET-28a(+) and pET-29a(+) vectors to obtain pET28orf4 and pET29orf4, respectively. All expression plasmids were verified by DNA sequencing.

The E. coli BL21(DE3)/pG-KJE8 strain was selected as the host; pG-KJE8 is a chaperone plasmid that can enhance the solubility of expressed protein in E. coli cells (46, 47). All expression plasmids described above were transferred into this host. Host cells harboring different plasmids were cultured in 100 ml of LB broth with 50 mg · liter⁻¹ Km and 30 mg · liter⁻¹ Cm at 37°C to an OD₆₀₀ of ∼0.2; 0.5 mg · ml⁻¹ l-arabinose and 5 ng · ml⁻¹ tetracycline were added to the culture. After an approximately 40-min incubation at 37°C, when the OD₆₀₀ reached about 0.6, 0.1 mM isopropyl β-d-1-thiogalactopyranoside (IPTG) was added to induce the expression of recombinant protein. After 10 h of incubation at 16°C with shaking 160 rpm, cells were harvested by centrifugation at 12,000 × g for 3 min, suspended in 15 ml of binding/washing buffer (50 mM NaH₂PO₄, 300 mM NaCl, 10 mM imidazole [pH 7.4]), and disrupted by sonication. After centrifugation at 12,000 × g for 30 min at 4°C, the supernatant was charged onto a 1-cm³ Co²⁺-charged resin (HiTrap Talon crude; GE Healthcare Life Sciences). Following washing with binding/washing buffer containing 50 mM imidazole, the target protein was eluted with 5 ml of elution buffer (50 mM NaH₂PO₄, 300 mM NaCl, 100 mM imidazole [pH 8.0]). Proteins nonspecifically bound to 6His-ORF4 and ORF4-6His were washed away using 30 mM imidazole containing binding/washing buffer and eluted by 5 ml of 50 mM imidazole-containing elution buffer.

All purified proteins were dialyzed in 1 liter of dialytic buffer (50 mM NaH₂PO₄, 100 mM NaCl, 20% glycerol, and 0.1 mM dithiothreitol [DTT] [pH 7.4]) and concentrated by ultrafiltration to 1 ml using a 10,000 molecular weight cutoff (MWCO) centrifugal filter (Merck Millipore, Germany). The purities of the purified enzymes were detected using 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels. Protein concentrations were quantified using the Bradford method with bovine serum albumin (BSA) as a standard (48).

Preparation of the cell extract.

Strains DE-13 and DE-13-E9 were individually grown to OD₆₀₀ of 1.0 in 100 ml of LB broth. The cells were harvested, suspended in 15 ml PBA buffer (137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, and 2 mM KH₂PO₄ [pH 7.4]), and disrupted by sonication; this was followed by centrifugation at 15,000 × g for 30 min at 4°C, and the supernatant was transferred to a new tube as the cell extract.

Enzyme assays.

All enzyme reaction mixtures were measured in PBA buffer with a final volume of 1 ml. NAD(P)H consumption at 340 nm was used for the determination of oxidoreductase activity, and at this wavelength, the molar extinction coefficient for NAD(P)H was 6,220 M⁻¹ · cm⁻¹. In the enzyme kinetics measurement, seven concentrations of NADH or NADPH (10, 20, 50, 80, 120, 160, and 200 μM) were measured in the presence of 10 μM FAD or FMN; six concentrations of FAD or FMN (1, 3, 6, 10, 15, and 20 μM) were measured in the presence of 100 μM NADH or NADPH. A 0.025 μM recombinant MeaY-6His solution was used in each of these assays; the reactions were started by addition of NAD(P)H and incubated at 30°C for 5 min.

To evaluate the relationship between the molar ratio of MeaX:MeaY and MEA hydroxylation activity, purified 6His-MeaX was kept constant at 0.25 μM, while the concentration of MeaY-6His varied from 0.005 to 1.25 μM (0.005, 0.0125, 0.025, 0.125, 0.25, and 1.25 μM). The concentration of MEA was 150 μM. The reaction mixture was incubated at 30°C for 10 min and stopped by adding an equal volume of methanol. Twenty microliters of the mixture was analyzed by HPLC, as described above. The concentration of MEA was calculated according to a standard curve, and relative degradation rates were also calculated when the highest degradation rate was selected as 100%.

In the case of oxygenase activity determination (recombinant MeaXY against MEA and DEA), seven concentrations of substrates (75, 112.5, 150, 187.5, 225, 262.5, and 300 μM) were used in the presence of 100 μM NADH, 10 μM FMN, 0.25 μM 6His-MeaX, and 0.0125 μM MeaY-6His; 500 U · ml⁻¹ catalase (bovine) was also added to the reaction mixture to regenerate trapped oxygen from hydrogen peroxide. The reaction mixture was incubated at 30°C for 10 min and the reaction stopped by adding an equal volume of methanol. Finally, 20 μl of the mixture was analyzed by HPLC, as described above. The concentration of MEA or DEA was calculated according to standard curves.

Kinetic data were evaluated by nonlinear regression analysis with the Michaelis-Menten equation v = V_max × [S]/(K_m + [S]), where v is the reaction rate and [S] is the concentration of substrate; the k_cat was calculated using the equation V_max = k_cat × [total enzyme concentration]. All data were collected from three independent determinations. One unit of enzyme activity was defined as the amount of enzyme required to catalyze 1 μmol substrate per min at 30°C.

Identification of MeaXY-hydroxylated MEA and DEA by NMR.

MeaXY-hydroxylated MEA and DEA were purified using silica gel column chromatography (3 by 40 cm), and the mobile phase was ligroin-chloroform-methyl cyanides at a 20:6:5 ratio. The eluate was collected per 20 ml, and the target collected eluate for NMR analysis was confirmed by HPLC and MS-MS analysis. In NMR analysis, 0.7 ml of dimethyl sulfoxide-d6 was used for the dissolution of 3 mg of purified product in a 5-mm NMR tube, and ¹H spectra were recorded at 500 MHz using a spectrometer with standard H1 parameters (Bruker Avance III, 500 MHz). The results were analyzed using Bruker TopSpin 3.1 software. Chemical shifts are reported in delta (δ) units, parts per million (ppm) downfield from tetramethylsilane, or ppm relative to the center of the singlet at 2.50 ppm for DMSO-d6. Coupling constant J values are reported in hertz (Hz), and the splitting patterns were designated s, singlet; d, doublet; t, triplet; dd, double of doublets; m, multiplet.

RT-qPCR.

Cells of strain DE-13 were cultured to the logarithmic phase in LB broth, suspended in MSM medium containing 0.5 mM MEA, DEA, or glucose, and cultured at 30°C for 8 h. Total RNA was extracted using MiniBEST universal RNA extraction kit (TaKaRa, Dalian, China), according to Wang et al. (49). Genomic DNA was eliminated with the genomic DNA (gDNA) eraser (TaKaRa) at 42°C for 2 min. Reverse transcription was conducted in the following reaction system at 37°C for 15 min: 1 μg of total RNA, 50 pmol random primer, and 5 U of reverse transcription enzyme (PrimeScript RTase; TaKaRa). Transcription levels of meaX, orf4, and meaY were performed in the 7300 real-time PCR system (Applied Biosystems, USA) using SYBR Premix Ex Taq RT-PCR kit (Tli RNaseH Plus; TaKaRa), according to the manufacturer's manual. The 16S rRNA gene was set as an internal standard, and the 2^−ΔΔCT threshold cycle (C_T) method was carried out to quantify relative expression (50). All primers used in the RT-qPCR are listed in Table 5.

Accession number(s).

The sequence of the lost DNA fragment in strains DE-13-E9 was submitted to GenBank under the accession number KT962120 (nucleotides 1030 to 15030 were lost in the mutant strain). The genome sequence of strain DE-13, including one circular chromosome and eight circular plasmids, was submitted to GenBank under the accession numbers CP013264 to CP013272.