Two dcm Gene Clusters Essential for the Degradation of Diclofop

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Agricultural and Environmental Chemistry

Two dcm Gene Clusters Essential for the Degradation of Diclofop-methyl in a Microbial Consortium of Rhodococcus sp. JT-3 and Brevundimonas sp. JT-9 Hui Zhang, Ting Yu, Jie Li, Yi-Ran Wang, Guang-Li Wang, Feng Li, Yuan Liu, Ming-Hua Xiong, and Yingqun Ma J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.8b05382 • Publication Date (Web): 30 Oct 2018 Downloaded from http://pubs.acs.org on October 30, 2018

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Journal of Agricultural and Food Chemistry

Two dcm Gene Clusters Essential for the Degradation of Diclofop-methyl in a Microbial Consortium of Rhodococcus sp. JT-3 and Brevundimonas sp. JT-9

Hui Zhanga, Ting Yua, Jie Lia, Yi-Ran Wanga, Guang-Li Wanga*, Feng Lia*, Yuan Liua, Ming-Hua Xionga, Ying-Qun Mab

a College of Life Sciences, Huaibei Normal University, Huaibei 235000, China b Advanced Environmental Biotechnology Centre, Nanyang Environment & Water Research Institute, Nanyang Technological University, 1 Cleantech Loop, Singapore 637141, Singapore

*Corresponding

author:

Guang-Li Wang, [email protected]

Address: College of Life Sciences, Huaibei Normal University, Huaibei 235000, China Tel/Fax: +86 561 3803024

ACS Paragon Plus Environment


Abstract: The metabolism of widely used aryloxyphenoxypropionate herbicide has been extensively studied in microbes. However, the information on the degradation of diclofop-methyl (DCM) is limited, with no genetic and biochemical investigation reported. The consortium L1 of Rhodococcus sp. JT-3 and Brevundimonas sp. JT-9 was able to degrade DCM through a synergistic metabolism. To elaborate the molecular mechanism of DCM degradation, the metabolic pathway for DCM was first investigated. DCM was initially transformed by strain JT-3 to diclofop acid and then by strain JT-9 to 2-(4-hydroxyphenoxy) propionic acid as well as 2,4-dichlorophenol. Subsequently, the two dcm gene clusters, dcmAE and dcmB1B2CD, involved in further degradation of 2,4-dichlorophenol, were successfully cloned from strain JT-3 and the functions of each gene product were identified. DcmA, a glutathione-dependent dehalogenase, was responsible for catalyzing the reductive dehalogenation of 2,4-dichlorophenol to 4-chlorophenol, which was then converted by the two-component monooxygenase DcmB1B2 to 4-chlorocatechol as the ring cleavage substrate of the dioxygenase DcmC. In this study, the overall DCM degradation pathway of the consortium L1 was proposed and, particularly, the lower part on the DCP degradation was characterized at the genetic and biochemical levels.

Keywords: Diclofop-methyl; Microbial consortium; Microbial degradation; Molecular mechanism


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1 Introduction As one of the representative aryloxyphenoxypropionate (AOPP) herbicides, diclofop-methyl (DCM, (R, S)-methyl-2-[4-(2,4-dichlorophenoxy) phenoxy] propionate), with potent inhibition of acetyl-CoA carboxylase as well as photosynthesis and meristem activity, is widely used for the control of annual and perennial grass weeds in broadleaf crop fields such as wheat, barley, and legumes.1, 2 The US Environment Protection Agency reported that the total annual domestic usage of DCM is approximately 750,000 pounds of as an active major ingredient.3 Due to its widespread application, DCM and its transformation products can be transported to surface waters and shallow groundwater, leading to a high environmental pollution risk.4-6 In addition, long-term exposure to high concentrations of DCM has been shown to cause lung injury, such as pneumoconiosis,7 and chromosomal aberration of mammalian lymphocytes both in vivo and in vitro.8 Thus, the environmental removal and degradation mechanisms of DCM have attracted considerable attention. Compared to the available physical and chemical removal methods, microbial degradation is thought to be a more effective and thorough strategy to eliminate DCM from a contaminated environment. To date, several strains capable of degrading AOPP herbicides, including the fluazifop-P-butyl-degrading strain Aquamicrobium sp. FPB-1,9 the fenoxaprop-P-ethyl (FE)-degrading strain Acinetobacter sp. DL-2,10 the cyhalofop-butyl-degrading strain Pseudomonas azotoformans QDZ-1,11 and the quizalofop-P-ethyl (QE)degrading strain Ochrobactrum sp. QE-9,12 have been isolated based on their ability to utilize AOPP herbicide as a sole carbon and energy source. However, these strains transform the AOPP herbicides into the corresponding acids by a specific carboxylesterase, but show no further degradation, resulting in incomplete degradation. Recently, an FE-mineralizing bacterial consortium W1 was isolated by soil-enrichment culture and the FE degradation mechanism was extensively investigated. Through this, fenoxaprop acid, 6-chloro-



2,3-dihydrobenzoxazol-2-one (CDHB), 2-(4-hydroxyphenoxy) propionic acid (HPP), and 2-amino-5-

chlorophenol were identified as intermediate metabolites by HPLC/MS.13, 14 Acinetobacter sp. DL-2, which is responsible for the transformation of FE to fenoxaprop acid, and Pigmentiphaga sp. DL-8, which is involved in the degradation of CDHB and HPP, were successfully isolated from the bacterial consortium W1. However, a single bacterial strain with the ability of converting fenoxaprop acid to CDHB and HPP, which is considered the critical step in the degradation process of FE, has not been isolated.13 Similarly, the degradation of [14C]DCM to [14C]CO2 by a biofilm consortium comprising nine bacterial strains and one alga, demonstrated that environmental metabolism of DCM required microbial synergetic degradation.15 In 1996, a DCM degrading bacteria consortium consisting of Chryseomonas luteola and Sphingomonas paucimobilis was isolated from herbicide-polluted soil in Canada.16 The Sphingomonas strain was able to degrade the initial hydrolyzing product of DCM, diclofop acid (DC), to 4-(2,4-dichlorophenoxy) phenol, 2,4dichlorophenol (DCP) and phenol, yet, accumulation of DCP in the growth medium was observed, suggesting that this strain alone was unable to completely degrade DC as well as DCM.17 The information on the microbial degradation of DCM is limited, with no genetic and biochemical investigations reported. This study aims to investigate the molecular mechanism of DCM degradation by the consortium L1, which was isolated from QE-contaminated soil using QE as a sole carbon source.18 Interestingly, the consortium L1 was proven to be able to degrade DCM by co-operation between strains JT-3 and JT-9. In this study, the overall DCM degradation pathway was elucidated through the degradation tests using the two consortium members JT-3 and JT-9 and the chemical analyses of the DCM intermediates. In addition, the two gene clusters involved in the lower pathway of DCM degradation were cloned from strain JT-3 and the functions of each gene product were characterized. These results suggest a novel DCM


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degradation pathway by the bacterial consortium.

2 Materials and methods 2.1 Chemicals, media, and strains DCM (purity, 99.5%), DC (purity, 99.2%), HPP (purity, 95%), DCP (purity, 99%), 4-chlorophenol (4CP) (purity, 99%), and 4-chlorocatechol (4-CC) (purity, 98%) were purchased from J&K Chemical Co., Ltd., China. All other chemicals and reagents were of analytical or high-performance liquid chromatography (HPLC) grade and are commercially available. The primers used in this study were synthetized from Takara Biomedical Technology (Dalian) Co., Ltd., China (Supplementary Table S1). Luria–Bertani (LB) medium and mineral salts medium (MM) were used for bacterial cultivation.12 When necessary, kanamycin (50 μg mL–1) or chloramphenicol (Cm, 34 μg mL–1) were added. 2.2 Synergistic degradation and growth of the consortium L1 Strains JT-3 and JT-9 were grown in LB medium for approximately 24 h at 30 °C and 180 rpm. The cells were harvested by centrifugation (5000 g, 10 min) and washed twice with fresh MM. Cell density was then adjusted to approximately 1.0 × 108 CFU mL–1, and an aliquot (2%, v/v) was inoculated into 100 mL of MM medium supplemented with 100 mg L-1 of DCM or its intermediates as the sole carbon and energy source. All cultures were incubated at 30 °C and 160 rpm on a rotary shaker. At regular intervals, 5 mL of samples were withdrawn from the cultures for analysis. In order to identify the intermediates of DCP prior to ring cleavage, 1 mM 2,2’-dipyridyl was used as an inhibitor of aromatic ring dioxygenases in the MM.19 The concentrations of DCM and its metabolic intermediates were determined by HPLC analysis based on a peak area from the calibration curve. The amount of cells of each strain was calculated by a dilution and plate counting method.18 Of note, the individual degradation of each strain was detected under the same conditions



as the synergistic degradation of both strains. 2.3 Chemical analysis For HPLC analysis, each sample was extracted twice with an equal volume of dichloromethane. The extracts were pooled and dried with anhydrous sodium sulfate. After rotary evaporation, the residues were dissolved in 1 mL of methanol. HPLC analysis was performed as described previously with minor modifications.12 To identify the metabolic intermediates of DCM transformed by the consortium L1 or by each strain, the samples were subjected to LC ion-trap mass spectrometry (LC-ITMS) analysis.18 Electrospray ionization (ESI) was performed in negative ion mode and full scan in the range 100–500 m/z. Due to the poor ionization efficiency for DCP metabolites in the LC-ITMS analysis, gas chromatography-mass spectrometry (GC-MS) analysis was further conducted to determine the identity of the metabolites. The supernatant from each sample was acidified prior to extraction with ether, as previously described.20 The resulting organic phases were concentrated and analyzed using a Thermo Fisher Trace GC Ultra gas chromatograph equipped with an HP-5MS capillary column (30 m × 0.25 mm × 0.25 μm) and coupled to a Thermo Fisher ITQ 900 ion trap mass spectrometer. Helium was used as the carrier gas at a flow rate of 1 mL min–1. The oven was initially held at 40 °C for 1 min, ramped to 240 °C at a rate of 5 °C min–1, and held for 3 min. Mass spectra were recorded in the range 40–200 m/z. The intermediates were identified through comparing the GC retention times and mass spectra of those authentic compounds available in NIST98 mass spectra data library. 2.4 Real-time quantitative PCR (RT-qPCR) The genomic DNA from strain JT-3 was isolated using the high salt precipitation method12 and submitted to BGI Technology Service Co., Ltd. (Shenzhen, China) for whole-genome sequencing and


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annotation. Target gene primers were designed and used to investigate the transcriptional levels of the gene clusters dcmAE and dcmB1B2CD in response to the various substrates. Strains JT-3 and JT-9 were grown in LB medium at 30 °C. Cells were harvested, washed twice, and diluted to approximately 1.0 × 108 CFU mL– 1

with fresh MM medium. An aliquot (2%, v/v) was inoculated into 100 mL of MM medium containing 100

mg L-1 DCM and incubated at 30 °C and 160 rpm for 54 h. For DCP degradation induction, only strain JT-3 was inoculated into MM medium containing 100 mg L-1 DCP and cultured under the same conditions. In control experiments, yeast extract (0.2%, w/v) was added instead of DCM and DCP. Total RNA was extracted with the RNAiso Plus (Takara) and reverse transcription PCR was performed with the PrimeScript RT reagent kit (Takara) in a 20-μL reaction mixture to obtain cDNA. RT-qPCR was performed with a 7300 Fast Real-Time PCR System (Applied Biosystems) in a 25 μL reaction mixture using SYBR Premix Ex Taq II (Takara). All samples were run in triplicate, in three independent experiments. The relative transcriptional levels of the target genes were evaluated according to the 2-∆∆CT method using the 16S rRNA gene as a reference.21 2.5 Protein expression and purification The genes dcmA, dcmB1, dcmB2 and dcmC were amplified from the genomic DNA of strain JT-3 by PCR. The resulting PCR products were respectively digested and cloned into the corresponding sites of pET29a(+) to generate the expression plasmids. The plasmids were subsequently transformed into E. coli (DE3) for protein expression. N-terminal 6 × histidine-tagged (H6) recombinant proteins

were purified using a Ni-

NTA affinity chromatography column (Sangon Biotech, Shanghai, China).12 The protein concentrations were determined by the Bradford assay using bovine serum albumin as the standard. 2.6 Enzyme assays



The dehalogenase activity of DcmA against DCP was measured under anaerobic conditions by adding a reduced form of glutathione (GSH) to remove residual oxygen from the solution. The reaction mixture contained 1 mM GSH, 1–100 μg of purified H6-DcmA, and 50 μM DCP in 10 mL of phosphate buffer (50 mM, pH7.5). The reaction was initiated with the addition of the substrate. The reaction mixture was extracted with ethyl acetate after acidification and subjected to GC-MS analysis to determine the substrate concentration. For kinetic assays, DCP was appropriately diluted into at least six different concentrations in the range 0–150 μM. Data from three independent experiments were fit into the Lineweaver–Burk transformation of the Michaelis–Menten equation with OriginPro 8 software (OriginLab, USA). One unit (U) of activity was defined as the amount of enzyme consuming 1 μmol of substrate per min at 30 °C. The nicotinamide adenine dinucleotide (NADH) oxidizing activity of DcmB2 was assayed by measuring the decrease in absorbance of NADH at 320 nm (ε = 6,220 M-1 cm-1) in 1 mL of phosphate buffer (50 mM, pH7.5) containing 200 μM NADH and 25 μM flavin adenine dinucleotide (FAD) at 25 °C. The reaction was initiated by addition of purified H6-DcmB2 (0.8 μg). The enzyme activity of 4-CP monooxygenase (DcmB1) was assayed in 50 mM phosphate buffer (pH 7.5), as previously described,20 with several modifications. The reaction mixture contained 200 μM NADH, 25 μM FAD, purified H6-DcmB2 (0.1–1 μg), and purified H6DcmB1 (10–100 μg) in a final volume of 1 mL. The assay was initiated by addition of substrate 4-CP. One unit (U) of activity was defined as the amount of enzyme required for oxidization of 1 μmol of NADH for reductase DcmB2 or 1 μmol of 4-CP for 4-CP monooxygenase DcmB1 per min. The products of the reaction catalyzed by purified H6-DcmB1 and H6-DcmB2 were detected by GC-MS analysis. The detection of activities of the enzymes involved in the sequential transformation of 4-CC was performed as follows. The ring cleavage activity of 4-CC dioxygenase (DcmC) was assessed by monitoring the change in absorption at


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260 nm due to the formation of 3-chloro-cis,cis-muconate (3-CM) from 4-CC.22 Further, the release of chloride ions was detected and quantified by the measurement of absorbance at 460 nm according to the standard curve for chloride ions from known various concentrations of NaCl solutions, as previously described.23 2.7 Gene knockout and complementation of dcmA The gene knockout plasmid pK18mobsacB-dcmA was constructed by fusing the upstream (553 bp) and downstream (499 bp) fragments of the dcmA gene and chloramphenicol resistance gene (1075 bp) from plasmid pXMJ1920 to EcoRI/HindIII-digested vector pK18mobsacB24 using in-fusion HD cloning kit (Takara). The resulting plasmid was then introducted into strain JT-3 by electrotransformation.20 The singlecrossover colonies were screened on LB plates containing Cm and then replica plated on LB plates supplemented with 10% (w/v) sucrose. The Cm-resistant but kanamycin-sensitive double-crossover recombinants were selected for PCR analysis.20 Plasmid pRESQ-dcmA for gene complementation was generated by fusion of the dcmA gene and its native promoter into BglII-digested vector pRESQ.25 Thereafter, the construct was introduced into the mutant strain JT-3∆dcmA by electrotransformation to obtain the dcmAcomplemented strain JT-3∆dcmA(dcmA), regaining the ability of utilizing DCP as the sole carbon source for growth. The gene disruption and gene complementation in these mutants were confirmed by PCR and sequencing. 2.8 Nucleotide sequence accession numbers The nucleotide sequences of approximately 4.9-kb and 8.4-kb DNA fragments harboring dcmAE dcmB1B2CD, respectively, from Rhodococcus sp. strain JT-3 described in this study were deposited in the GenBank database under accession numbers MH579712 and MH579713.



3 Results and discussion 3.1 Synergistic degradation and growth of the consortium L1 DCM degradation by the consortium L1 was assessed using it as the sole carbon source. When strains JT-3 and JT-9 were simultaneously inoculated into MM containing 100 mg L–1 of DCM, DCM was degraded within 72 h (Figure 1A). The amount of cells of strain JT-3 increased from 2.0 × 106 CFU mL–1 to 11.9 × 106 CFU mL–1 in 84 h. For strain JT-9, the cells rapidly increased after approximately 24-h lag time, reaching 8.4 × 106 CFU mL–1 from 2.0 × 106 CFU mL–1 at 72 h. In our previous study, an esterase, named EstS-JT, catalyzing the initial hydrolysis of DCM to DC was found from strain JT-3 and the function was confirmed by chemical analysis of the enzymatic reaction using LC-MS18. Thus, to investigate the degradation pathway lower than DC, the degradation dynamics of DC by both strains JT-3 and JT-9 was examined in MM using DC as the sole carbon source. The two strains exhibited similar degradation and growth curves when inoculated into DCM-containing MM except for the shorter lag phase (12 h) (Figure 1B). These observations indicate that strains JT-3 and JT-9, inoculated as a consortium, are capable of utilizing DCM or DC as the sole carbon source for growth. In order to estimate the roles of each strain during the DCM degradation, each strain was inoculated to MM supplemented with DC. When strain JT-9 was inoculated alone in DC-containing MM, it was able to degrade 100 mg L–1 of DC in 84 h, reaching a cell density of 7.8 × 106 CFU mL–1 at 84 h (Figure 1C), suggesting that strain JT-9 could utilize DC as its sole carbon source for growth. In contrast, a negligible change in DC concentration was observed in MM inoculated with strain JT-3 (Figure 1C), indicating that strain JT-3 was incapable of degrading DC. Interestingly, though strain JT-9 could utilize DC as the sole carbon source for growth, an accumulation of DCP was observed during the degradation of DC by strain


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JT-9 alone, suggesting that the degradation of DC requires the cooperative metabolism of strains JT-9 and JT-3. Thus, DCP degradation by each individual strain was performed in MM; as expected, 100 mg L–1 of DCP was degraded by strain JT-3 in 60 h (Figure 1D). The cell density quickly increased to 11.1 × 105 CFU mL–1 at 60 h and remained stable for 12 h. Conversely, inoculation with strain JT-9 led to no significant change in DCP concentration, with a gradual decrease in the growth curve. These results clearly demonstrate that strain JT-3 initiated the conversion of DCM to DC, which was used as the carbon source by strain JT-9 for further degradation. Subsequently, DCP, as a metabolite of DC, was degraded by strain JT-3. Thus a synergistic metabolism in the consortium L1 occurs during DCM degradation, with mutual assistance to achieve cell growth and degradation of DCM. Several bacterial consortia have been described to synergistically degrade a variety of environmental contaminants, such as FE13 and bromoxynil octanoate.26 Indeed, bromoxynil octanoate was shown to be degraded by the consortium consisting of two bacterial strains (Sphingopyxis sp. OB-3 and Comamonas sp. 7D-2) through a similar synergistic mechanism.26 The two members of the consortium facilitated the growth of each other through cross-feeding with metabolites from the degradation pathway. 3.2 Identification of metabolic intermediates In order to determine the metabolic intermediates of DCM degradation by the consortium L1, all samples were analyzed at 5 days after inoculation by HPLC and LC-ITMS. Three metabolites were detected, with retention times of 7.57, 5.36, and 10.71 min, respectively (Figure 2A). Metabolite I was identified as DC, with a protonated molecular parent ion peak at m/z 327.14 (Figure 2B) and characteristic second-order fragment ion peaks at m/z 281.08 (loss of –COOH) and at m/z 252.92 (loss of –CH3CH2) (Figure 2C). Metabolite II displayed a protonated parent ion peak at m/z 163.12 (Figure 2D) and characteristic second-



order fragment ion peak at m/z 147.36 (loss of –OH) (Figure 2E), and was identified as DCP. Metabolite III had a protonated parent ion peak at m/z 182.16 (Figure 2F) and characteristic second-order fragment ion peaks at m/z 139.02 (loss of –COOH) and at m/z 108.48 (loss of –CH3CH2) (Figure 2G). Thus, it was assigned to HPP, a major intermediate that has been shown to be present in the microbial metabolism of another AOPP herbicide, FE.10 On the basis of the identification of DCM metabolites, a plausible DCM degradation pathway by both strains JT-3 and JT-9 was proposed (Figure 3B). Nevertheless, the DCM degradation presented herein was significantly different from the previously reported metabolisms of clodinafop27 and DCM,17 in which the corresponding acid, DC, produced by the breakdown of the ester bond of DCM, was transformed to DCP and HPP by hydrolyzing the C–O bond between two benzene rings rather than directly by removal of propionic acid to form phenoxy-phenol. Unfortunately, the gene responsible for the conversion of DC to HPP and DCP is yet not found in strain JT-9. To further determine how DCP was degraded by strain JT-3 in the consortium L1, strain JT-3 was inoculated in MM containing 100 mg L–1 DCP and 1 mM 2,2’-dipyridyl. At 48 h after inoculation, two metabolites with GC retention times of 11.28 and 14.52 min, respectively, were accumulated with DCP consumption (Figure 4A). Compared to the mass spectra of the authentic compounds, these two metabolites, IV and V, were identified as 4-CP and 4-CC by GC-MS analysis, respectively (Figure 4B–E). Additionally, quantitative analysis of the intermediates indicated that the stoichiometric amount of DCP consumption (482 μM) was nearly equal to the total yield of both 4-CP (411 μM) and 4-CC (28 μM). Furthermore, with the increase in degradation time, a gradual accumulation of 4-CC, coupled with the continuous consumption of 4-CP, was observed (data not shown), demonstrating that strain JT-3 degraded DCP to 4-CP and then 4-CC.


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In the absence of 2,2’-dipyridyl, 4-CC was not detected, suggesting further degradation of 4-CC by the ringcleavage (the data was shown later) (Figure 3B). DCP has also been found to be the first intermediate during the degradation of herbicide 2,4-dichlorophenoxyacetic acid (2,4-D),28, 29 wherein 2,4-D was converted into 3,5-dichlorocatechol by hydroxylase and subsequently used as the ring cleavage substrate for further degradation. In the case of DCP degradation by strain JT-3, 4-CP, as the initial intermediate of DCP degradation by the dehalogenation reaction, was detected for the first time herein. Recently, Pandey et al.30 described a degradation pathway of 2-chloro-4-nitrophenol (2C4NP) in Burkholderia sp. strain SJ98 by a similar dehalogenation under aerobic condition. The first step of 2C4NP degradation by strain SJ98 was the reductive dehalogenation, resulting in the formation of 4-nitrophenol. 3.3 Sequence analyses of the DCM metabolic gene clusters According to the proposed DCM degradation pathway based on the identified intermediates during DCM and DCP degradation, two gene clusters likely involved in DCP degradation were found on contigs 3 and 7 of the draft genome sequence of strain JT-3 and designated dcmAE and dcmB1B2CD, respectively (Figure 3A). Among dcmAE, dcmA, encodes a protein (DcmA) of 351 amino acid residues, which exhibits 55% and 30% similarity with 2,5-dichlorohydroquinone reductive dechlorinase (AAN64243) from Sphingomonas paucimobilis B90 and glutathione S-transferase family protein (AGI68666) from Octadecabacter antarcticus 307, respectively, suggesting that it appeared to belong to the glutathione Stransferase family and was involved in the reductive dechlorination of DCP during DCM degradation. On both the 5’- and 3’-flankinng regions of dcmA, two complete IS6100 transposable element including inverted repeats were found, in which gene tnpA encoding proteins had 99% identity to the IS6100 transposase. Such gene organization flanked by insertion elements was reported to exist in the 2,4-D degradation gene cluster



of Delftia acidovorans strain P4a, isolated from heavily organochlorine-contaminated soil.31 The dcmE gene encodes a protein (DcmE) of 354 amino acid residues, which displays 58% homology with the maleylacetate reductase MacA (AAC38802) from Rhodococcus opacus 1CP. Similar phenomenon is found that the gene (macA) encoding MacA was relatively distant from the major counterparts responsible for phenol metabolism, suggesting that the dcmE and macA genes are not members of the specialized aromatic compound metabolic genes.32 For another gene cluster, dcmB1B2CD, DcmB1 and DcmB2 showed 79% and 56% similarity to the oxygenase and reductase components, respectively, of the 4-nitrophenol hydroxylase (Q8RQQ0) from Rhodococcus sp. PN1, suggesting that DcmB1B2 probably belongs to the two-component flavin-diffusible monooxygenase (TC-FDM) family. DcmC and DcmD shared moderate similarity to chlorocatechol 1,2dioxygenase (AAC38249, 41%) and chloromuconate cycloisomerase (AAC38251, 40%), respectively, from Rhodococcus opacus 1CP. 3.4 Transcriptional analyses of the gene cluster dcmAE and dcmB1B2CD To estimate whether the gene clusters dcmAE and dcmB1B2CD are involved in DCM degradation, the transcriptional levels of these two gene clusters in response to substrates DCM and DCP were investigated. At 54 h after inoculation by both JT-3 and JT-9 in MM with or without the substrate DCM, total RNA was extracted from strain JT-3 and utilized as the template for RT-qPCR. RT-qPCR results revealed that the transcriptional levels of dcmA, dcmB1B2, dcmC, dcmD, and dcmE were dramatically increased compared to those from the samples without the addition of DCM, with a 345-, 271-, 57-, 49-, and 82-fold increase, respectively (Figure 5). The significant diversity in transcription levels of dcmA, dcmE, dcmB1B2, and dcmC was probably the reason that each gene was controlled by the respective independent transcriptional operons. Conversely, the approximate equivalence of transcriptional levels between dcmC and dcmD was suggested


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to be located in a single transcriptional operon. Likewise, for the samples from MM containing DCP inoculated by strain JT-3, the transcriptional levels of these genes were also obviously up-regulated. Thus, the results indicated that these tested genes were likely involved in DCM degradation by the consortium L1. 3.5 DcmA catalyzes the reductive dechlorination of DCP to form 4-CP Recombinant H6-DcmA was expressed in E. coli BL21 (DE3) by induction of 0.1 mM isopropyl β-Dthiogalactopyranoside (IPTG) at 16 °C overnight. Following purification and concentration, SDS-PAGE analysis only displayed a slight single protein band of approximately 44 kDa (Supplementary Figure S1), matching the calculated molecular mass of the recombinant H6-DcmA. Conversion of DCP by the purified protein was performed in the reaction mixture containing GSH and the metabolites from DCP were analyzed by GC-MS. One metabolite was detected at 11.28 min in the chromatogram and it showed a similar mass spectrum to that of authentic 4-CP (Supplementary Figure S2A). The dechlorination activity assay revealed that the catalytic velocity of H6-DcmA against DCP was strictly dependent on the presence of GSH (Supplementary Figure S3). These results suggest that DcmA catalyzes DCP to 4-CP in the presence of GSH through a mechanism analogous to that for tetrachlorohydroquinone dehalogenation by the GSH-dependent dehalogenase of Sphingobium chlorophenolicum,33 in which the enzyme activated GSH as a nucleophilic group to form a complex between GSH and DCP, leading to the displacement of chlorine in DCP with GSH. The Km value for GSH of H6-DcmA was 98 μM and maximal activity (3.2 U mg–1) was achieved in the presence of 1mM GSH, nearly equivalent to the value of dechlorinase BphK from Burkholderia xenovorans strain LB400.34 When the concentration of GSH added in the reaction mixture was far in excess of the amount required, the concentration of the DCP substrate varied from 0 to 150 μM. The kinetic analysis indicated the Km value of H6-DcmA was 26.7  3.1 μM, the catalytic efficiency kcat/Km and Vmax values were 0.11  0.02



μM–1 S–1 and 3.86  0.43 μmol min-1 mg-1, respectively. 3.6 DcmB1B2 catalyzes the hydroxylation of 4-CP to form 4-CC Recombinant H6-DcmB1 and H6-DcmB2 were individually expressed in E. coli BL21 (DE3) and purified by Ni-NTA affinity chromatography. SDS-PAGE analysis showed that the molecular masses of the purified H6-DcmB1 and H6-DcmB2 were approximately 65 and 26 kDa, respectively (Supplementary Figure S1), corresponding to their molecular weights based on the deduced amino acid sequences. As the reductase component of 4-CP 4-monooxygenase, the maximal NADH-oxidizing activity of DcmB2 was 1.4 U mg–1 in presence of FAD. The reaction was performed in the excess amount of FAD (25 μM) to determine the kinetic parameters of DcmB2 for NADH, the Km values of H6-DcmB2 for NADH was 31.7  3.5 μM, the catalytic efficiency kcat/Km and Vmax values were 0.29  0.03 μM–1 S–1 and 21.4  2.2 μmol min-1 mg-1, respectively. When the amount of NADH in the reaction mixture was in excess (200 μM), the Km values of H6-DcmB2 for FAD was 4.11  0.37 μM, the catalytic efficiency kcat/Km and Vmax values were 2.34  0.31 μM–1 S–1 and 22.2  2.7 μmol min-1 mg-1, respectively. As the function of DcmB2 was confirmed as an NADH/FAD oxidoreductase, conversion of 4-CP by the purified H6-DcmB1 was performed in the presence of NADH, FAD, and the purified H6-DcmB2. By the GC-MS analysis of the reaction mixture, 4-CC was identified as the product of the 4-CP conversion; the mass spectrum of the product was almost similar to that for authentic 4-CC (Supplementary Figure S2B). The oxidation activity assay of purified H6-DcmB1 and H6-DcmB2 demonstrated that the catalytic activity was closely related to the ratio of the two enzymes in the reaction mixture. A higher hydroxylation activity was clearly observed with an increase in H6-DcmB1 concentration in the reaction mixture (Supplementary Figure S4). Similar phenomena were observed in the cases of phenol hydroxylase PheA1A2 from Rhodococcus erythropolis UPV-135 and 4-nitrophenol monooxygenase from


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Rhodococcus imtechensis RKJ300.20 Maximal activity (0.12 U mg–1) was achieved when the molar ratio of H6-DcmB2 to H6-DcmB1 was approximately 1:35. The enzymatic kinetic assays led to the Km value of H6DcmB1 for 4-CP of 14.1  1.7 μM, the kcat/Km value of 0.21  0.01 μM–1 S–1 and the Vmax value of 2.73  0.29 μmol min-1 mg-1. 3.7 DcmC, DcmD, and DcmE are likely responsible for the sequential conversion of 4-CC The purified recombinant H6-DcmC was of approximately 35 kDa, consistent with its deduced molecular mass (Supplementary Figure S1). The 4-CC dioxygenase activity assay of the purified H6-DcmC revealed that the addition of the enzyme in the reaction mixture resulted in an increase of absorbance at 260 nm (Supplementary Figure S5), corresponding to the production of 3-CM.22 However, no specific absorbance change was observed upon H6-DcmC omission from the reaction mixture. Moreover, dcmC was evidently up-regulated when strain JT-3 was grown in MM using DCM or DCP as the sole carbon source. Due to the negligible increase in absorbance at 375 nm (corresponding to the formation of 5-chloro-2-hydroxymuconic semialdehyde), it was concluded that DcmC in strain JT-3, as a single subunit dioxygenase, might be responsible for the ortho-ring cleavage of 4-CC to form 3-CM, rather than its meta-cleavage, during the DCP degradation. The enzyme assays of DcmD and DcmE were not performed since their substrates were commercially unavailable. However, considering that DcmD and DcmE exhibited moderate identity with the chloromuconate cycloisomerase (AAC38251) and the maleylacetate reductase (AAC44727) of R. opacus 1CP, respectively, and their coding genes were significantly up-regulated in MM containing DCM or DCP, thus, it was speculated that DcmD and DcmE were likely involved in the further conversion of 3-CM. 3.8 DcmA was crucial for DCM metabolism of by the consortium L1 To evaluate the role of DcmA during DCM metabolism by the consortium L1, a mutant strain JT-



3∆dcmA, with a replacement of dcmA by a Cm-resistant gene, was constructed through double crossover recombination. Although the consortium comprising the mutant strain JT-3∆dcmA and strain JT-9 was able to grow with DCM as the sole carbon source, the two strains grew slowly and the maximum cell number was remarkably decreased compared to those of wild-type strains JT-3 and JT-9 (Figure 6A). Additionally, an accumulation of DCP was observed in MM inoculated with both the mutant strain JT-3∆dcmA and strain JT-9, and the consortium of these two strains was no longer able to rapidly degrade DCM, resulting in the poor growth on DCM (Figure 6A). Instead, the dcmA-complemented mutant strain JT-3∆dcmA(dcmA), together with strain JT-9, regained the ability to efficiently degrade DCM, though they exhibited slower growth and DCM degradation in comparison with the wild-type strains (Figure 6B). The above findings demonstrate that dcmA is necessary for the consortium L1 to efficiently degrade DCM.

AUTHOR INFORMATION Corresponding Author: *(G.-L.W.),

*(F.L.),

E-mail: [email protected]

E-mail: [email protected]

Funding This work was financed by grants from Provincial Natural Science Foundation of Anhui (1808085MC56), Major Program of Natural Science Research in Colleges and Universities in Anhui Province (KJ2018ZD039), Project of Outstanding Young Talents Supporting Plan in Universities of Anhui Province (gxyq2018020) and Construction Project from College Scientific Research Innovation Team of Anhui Province--Ecological


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Restoration and Utilization of Coal Mining Subsidence Area. Acknowledgments We are also grateful to Dr. Ma of Nanyang Technological University for assistance in the revision of manuscript.

Notes The authors declare no competing financial interest.



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Figure captions Figure 1. Degradation dynamics of DCM and its intermediates by strains JT-3 and JT-9. (A) Synergistic degradation of DCM by strains JT-3 and JT-9; (B) Synergistic degradation of DC by strains JT-3 and JT-9; (C) Individual degradation of DC by strain JT-3 and strain JT-9; (D) Individual degradation of DCP by strain JT-3 and strain JT-9. Figure 2. HPLC and LC-ITMS analyses of the DCM metabolites in the DCM degradation by the consortium L1. (A) HPLC chromatogram of the 5-d incubation sample; (B, D, F) Mass spectrum of the DCM metabolites I, II and III, respectively; (C, E, G) Second-order mass spectrum of the DCM metabolites I, II and III, respectively. Figure 3. (A) Organization of the dcm gene cluster of Rhodococcus sp. strain JT-3. The sizes and transcriptional directions of each gene and ORF were indicated by the black arrows. (B) Proposed degradation pathway of DCM by the consortium L1, together with the conversion reaction catalyzed by the dcm gene products. TCA, tricarboxylic acid. Figure 4. GC-MS analysis of the DCP metabolites in the DCP degradation by Rhodococcus sp. strain JT-3. (A) The gas chromatogram is the extracted ion current chromatogram at m/z 128.00  0.50 and 144.00  0.50 from the total ion current chromatogram. (B) Mass spectrum of authentic 4-chlorophenol (4-CP); (C) Mass spectrum of the GC peak at 11.28 min, identical to that of authentic 4-CP; (D) Mass spectrum of authentic 4chlorocatechol (4-CC); (E) Mass spectrum of the GC peak at 14.52 min, identical to that of authentic 4-CC. Figure 5. Transcriptional analyses of the gene cluster dcmAE and dcmB1B2CD of strain JT-3 with or without addition of DCM or DCP by RT-qPCR. DCM-containing MM was inoculated by both strains JT-3 and JT-9 and DCP-containing MM was inoculated by strain JT-3. All experiments were performed in triplicate, and error bars indicate standard deviations. Figure 6. Degradation dynamics of DCM by (A) the mutant strain JT-3∆dcmA and strain JT-9 or (B) by the dcmA-complemented mutant strain JT-3∆dcmA(dcmA) and strain JT-9.


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Figure 1 A

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Two dcm Gene Clusters Essential for the Degradation of Diclofop

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