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Journal of Hazardous Materials

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Biodegradation of 2,6-dibromo-4-nitrophenol by Cupriavidus sp. strain CNP-8: Kinetics, pathway, genetic and biochemical characterizationJun Mina,b, Weiwei Chena, Xiaoke Hua,b,⁎

a Key Laboratory of Coastal Biology and Bioresource Utilization, Yantai Institute of Coastal Zone Research, Chinese Academy of Sciences, Yantai 264003, Chinab Laboratory for Marine Biology and Biotechnology, Qingdao National Laboratory for Marine Science and Technology, Qingdao, China

G R A P H I C A L A B S T R A C T

A R T I C L E I N F O

Keywords:2,6-Dibromo-4-nitrophenolBiodegradationCatabolic mechanismCupriavidus sp.FADH2-dependent monooxygenase

A B S T R A C T

Compound 2,6-dibromo-4-nitrophenol (2,6-DBNP) with high cytotoxicity and genotoxicity has been recentlyidentified as an emerging brominated disinfection by-product during chloramination and chlorination of water,and its environmental fate is of great concern. To date, the biodegradation process of 2,6-DBNP is unknown.Herein, Cupriavidus sp. strain CNP-8 was reported to be able to utilize 2,6-DBNP as a sole source of carbon,nitrogen and energy. It degraded 2,6-DBNP in concentrations up to 0.7mM, and the degradation of 2,6-DBNPconformed to Haldane inhibition model with μmax of 0.096 h−1, Ks of 0.05mM and Ki of 0.31mM. Comparativetranscriptome and real-time quantitative PCR analyses suggested that the hnp gene cluster was likely responsiblefor 2,6-DBNP catabolism. Three Hnp proteins were purified and functionally verified. HnpA, a FADH2-dependentmonooxygenase, was found to catalyze the sequential denitration and debromination of 2,6-DBNP to 6-bro-mohydroxyquinol (6-BHQ) in the presence of the flavin reductase HnpB. Gene knockout and complementationrevealed that hnpA is essential for strain CNP-8 to utiluze 2,6-DBNP. HnpC, a 6-BHQ 1,2-dioxygenase wasproposed to catalyze the ring-cleavage of 6-BHQ during 2,6-DBNP catabolism. These results fill a gap in theunderstanding of the microbial degradation process and mechanism of 2,6-DBNP.

https://doi.org/10.1016/j.jhazmat.2018.08.063Received 24 June 2018; Received in revised form 18 August 2018; Accepted 20 August 2018

Abbreviations: 2,6-DBNP, 2,6-Dibromo-4-nitrophenol; 2,6-DBBQ, 2,6-dibromo-1, 4-benzoquinone; 2,6-DBHQ, 2,6-dibromohydroquinone; 6-BHQ, 6-bromohy-droxyquinol

⁎ Corresponding author at: Key Laboratory of Coastal Biology and Bioresource Utilization, Yantai Institute of Coastal Zone Research, Chinese Academy of Sciences,Yantai 264003, China.

E-mail address: xkhu@yic.ac.cn (X. Hu).

Journal of Hazardous Materials 361 (2019) 10–18

Available online 22 August 20180304-3894/ © 2018 Elsevier B.V. All rights reserved.

T

1. Introduction

Halogenated nitrophenols are widely used in the syntheses of agri-cultural and industrial chemicals and are common aromatic haloge-nated disinfection by-products (DBPs) during chloramination andchlorination of water [1,2]. Due to their mutagenic, teratogenic andcarcinogenic potential, halogenated nitrophenols in the environmenthave caused great problems for human health and ecosystems security.As an emerging aromatic halogenated DBP, 2,6-dibromo-4-nitrophenol(2,6-DBNP) has been identified in drinking water [3], swimming poolwater [2] and saline wastewater [4]. Previous toxicological studies hasdemonstrated that 2,6-DBNP was dozens to hundreds of times morecytotoxic than several regulated DBPs by the U.S. Environmental Pro-tection Agency (EPA) [5,6]. Recently, 2,6-DBNP has also been detectedin various mollusks from Bohai Sea, China [7], indicating its potentialrisk to human by bioaccumulation. Therefore, considering the hightoxicity of 2,6-DBNP and its wide distribution, removal of this haloge-nated nitrophenol from the environment is very meaningful.

Although several physico-chemical methods have been applied forhalogenated nitrophenols degradation [1,8], they are not cost-effectiveand are unable to mineralize the pollutants completely. Microorganismsplay key roles in the degradation of halogenated nitrophenols in theenvironment. Currently, bioremediation was considered to be the mostcost-effective and environmental-friendly means for pollution abate-ment, and it has been sucessfully used in treatment of numerous aro-matics-contaminated environments [9–15]. Therefore, in order toeliminate 2,6-DBNP from the environment by bioremediation, the es-sential prerequisite is to obtain a microorganism with the ability todegrade this toxicant.

Brominated nitrophenols (BNPs) are structurally analogues ofchlorinated nitrophenols (CNPs) which are the most common haloge-nated nitrophenols. To data, many microorganisms have evolved theircapacity to degrade various isomers of mono-chlorinated nitrophenols(MCNPs) including 2-chloro-4-nitrophenol (2C4NP) [16–19], 4-chloro-2-nitrophenol [20], 4-chloro-3-nitrophenol [21] and 2-chloro-5-ni-trophenol [22]. Noteworthily, the relative position of nitro and chlorosignificantly affected MCNPs degradation by microorganisms becauseof electron delocalization on the aromatic ring. Therefore, the enzyme(s) involved in 2C4NP degradation were not able to catalyze thetransformation of other MCNP isomers [16]. Moreover, even against thesame MCNP substrate, microorganisms have evolved different meta-bolic pathways [1]. The MCNPs-utilizers may also degrade the corre-sponding mono-brominated nitrophenols (MBNPs), but there is no ex-perimental evidence available. Further, in contrast to MBNPs,dibrominated nitrophenols (DBNPs) are more toxic and more resistantto microbial degradation because of an additional bromine on thebenzene ring. Thus, to date there is no data available on the microbialdegradation of DBNPs.

Herein, Cupriavidus sp. strain CNP-8 was proved to be able tocompletely mineralize 2,6-DBNP. The degradation kinetics of 2,6-DBNPby this strain was investigated. Moreover, the genes responsible for 2,6-DBNP catabolism was identified by comparative transcriptome assayand biological experimental validation. This is the first report of mi-crobiol degradation of 2,6-DBNP and enhance our understanding of theenvironmental fate of this pollutant.

2. Materials and methods

2.1. Strains, plasmids, primers and culture conditions

The bacterial strains and plasmids are described in Table 1, and theprimers are listed in Table S1. Cupriavidus sp. strain CNP-8 isolatedpreviously [22] has been deposited in the China Center for Type CultureCollection (accession number: M 2017546). Escherichia coli strains weregrown in lysogeny broth (LB) (tryptone: 10 g L−1, NaCl: 10 g L−1 andyeast extract: 5 g L−1) at 37 °C. Cupriavidus strains were grown in

minimal medium [23] (MM, without CaCl2) at 30 °C supplemented with2,6-DBNP. When necessary, 20% LB (v/v) was added into MM to en-hance the biomass. Kanamycin (50mg L−1) or chloramphenicol(34mg L−1) was added to the medium as necessary.

2.2. Biodegradation of 2,6-DBNP

Strain CNP-8 was initially cultivated in MM+LB (4:1, v/v) over-night. The harvested cells were washed thrice and suspended in freshMM to an initial OD600 of 0.05; followed by addition of 0.3 mM of 2,6-DBNP. The ability of strain CNP-8 to utilize 2,6-DBNP was determinedby monitoring the consumption of 2,6-DBNP, together with the growthof cells (OD600). To study the effect of simple carbon sources on 2,6-DBNP degradation, lactate, succinate or glucose (0.5 and 0.5 g L−1,respectively) was supplemented into MM containing 2,6-DBNP. Whole-cell biotransformation was carried out as described previously [24]. Allabove experiments were performed in triplicate.

2.3. 2,6-DBNP degradation kinetics

Degradation of 2,6-DBNP from 0.05 to 1.0mM by strain CNP-8 wasperformed to estimate the degradation kinetics parameters. Bacterialgrowth kinetics was modeled by the equation as follow:

X Xt t

µ ln ( / )0

0=

(1)

X: biomass concentration, μ: specific growth rate, t: time.Considering the toxicity of 2,6-DBNP against strain CNP-8, the

Haldane’s model with equation described as follow was used to in-vestigate the kinetics of 2,6-DBNP degradation [25].

SK S S Ki

µµ

( / )max

s2=

+ + (2)

S: 2,6-DBNP concentration, μmax: the maximal specific growth rate, Ks:the half saturation constant, Ki: the inhibition constant.

The biomass yield was calculated by the equation as follow [26]:

Y X XS S

0

0=

(3)

Xmax: the maximal biomass, X0: the initial biomass, S0: the initial 2,6-DBNP concentration, Sa: the 2,6-DBNP concentration when the biomassconcentration is maximal.

2.4. Comparative transcriptome and real-time quantitative PCR

The total RNA from 2,6-DBNP-induced and uninduced cells wasisolated by an Easy Pure RNA Kit (TransGen, Biotech, China). Then,cDNA was transcribed from total RNA followed by cDNA library se-quencing with Illumina HiSeq 2500 system (Novogene BioinformaticsTechnology Co., Ltd., Beijing, China). Reads generated by the sequen-cing machines were cleaned by discarding the adaptor and low-qualitysequences. Subsequently, the clean reads were then assembled andmapped to the genome sequence of strain CNP-8. Comparative analysisof the transcriptomes was performed as described previously [27]. Real-time quantitative PCR (RT-qPCR) was carried out using a TransStart TipGreen qPCR SuperMix kit (TransGen). The primers were listed in TableS1. The PCR reactions were performed in triplicates independently. Therelative expression levels of the target genes were estimated using the2 CT method [28] with the 16S rRNA gene as an internal control.

2.5. Protein expression and purification

The hnpA, hnpB, and hnpC genes were amplified from genomic DNAof strain CNP-8 using FastPfu DNA Polymerase (Transgene, China),with primers listed in Table S1. The PCR product was cloned into theNdeI and XhoI digested pET-28a(+), producing expression plasmids

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(Table 1). The plasmids with correct sequence of inserted genes weretransformed into E. coli BL21(DE3). The protein expression and pur-ification was carried out as described previously [29], with somemodification. BL21(DE3) harboring the expression plasmids was grownin LB at 37 °C to OD600 of about 0.5, and then induced with 0.2mMisopropyl-β-D-thiogalactopyranoside (IPTG) for 12 h at 16 °C. The har-vested cells were resuspended in binding buffer (50mM sodium phos-phate, 150mM NaCl, pH 8.0) and disrupted by ultrasonication. Thecellular debris was removed by centrifugation at 20,000×g for 30minat 4 °C, and the supernatants were applied to Ni-NTA agarose (GEHealthcare, USA) pre-equilibrated with binding buffer. Agarose columnwas washed sequentially with binding buffer containing 20, 40 and80mM imidazole to remove nonspecifically bound proteins. Finally,binding buffer containing 200mM imidazole was used to elute the his-tagged Hnp proteins. The purified proteins were dialyzed againstbinding buffer to remove imidazole, and their purities and molecularweights were determined by SDS-PAGE. Protein concentration wasmeasured by the Bradford assay [30].

2.6. Enzyme assays

All the enzyme assays were carried out at 30 °C, in 50mM Tris-HClbuffer (pH 7.5) in a total volume of 1ml using a Perkin Elmer Lambda365 UV/Vis spectrophotometer. The activity of H6-HnpB was de-termined spectrophotometrically by measuring the decrease of NAD(P)H at 340 nm (ε340= 6220 M−1 cm−1) [31]. The reaction system con-tained Tris-HCl buffer, NAD(P)H (0.2 mM), FAD (0.02mM) and H6-HnpB (5 μg). The Michaelis-Menten kinetics was investigated by in-creasing FAD from 0.2 to 20 μM when NADH was fixed at 200 μM orincreasing NADH from 10 to 200 μM when FAD was fixed at 20 μM.

For the activity assay of H6-HnpA against 2,6-DBNP, the 1ml re-action system contained Tris-HCl buffer, NADH (0.2mM), FAD(0.02mM), H6-HnpB (5 μg) and H6-HnpA (30 μg), and the enzyme ac-tivity was assayed by monitoring the decrease of 2,6-DBNP at 400 nm.The enzyme kinetics was investigated by increasing 2,6-DBNP from 2.5to 40 μM. The concentration of 2,6-DBNP was determined by HPLCanalysis. One unit of enzyme activity was defined as the quantity ofenzyme needed to consume 1 μmol substrate in 1min. To identify theproducts of 2,6-DBNP, the products in the reaction system (10ml) wereinitially acetylated as described previously [31] and then identified byGC–MS analysis.

Because 6-bromohydroxyquinol was not commercially available, itsstructure analogue hydroxyquinol was used to assay H6-HnpC activity.

The 1ml reaction system contained Tris-HCl buffer, ascorbic acid(1mM), HnpC (2 μg) and hydroxyquinol (0.1mM). The activity of H6-HnpC was determined by monitoring the consumption of hydro-xyquinol (λmax= 289 nm) and formation of maleylacetate(λmax= 243 nm).

2.7. Gene knockout and complementation

The upstream (800 bp) and downstream (849 bp) fragments of hnpAwere amplified from the genomic DNA and the chloramphenicol re-sistance gene (cm) was amplified from pXMJ19 (Table 1), with primerslisted in Table S1. These fragments were ligated into pK18mobsacB (pre-digested with EcoRI and HindIII) using In-Fusion HD Cloning Kit (Ta-kara) to yield the suicide plasmid pK18-hnpA. The plasmid was thentransformed into mobilizer strain E. coliWM3064 (Table 1). The suicideplasmid in WM3064 was transformed to strain CNP-8 by biparentalmating [32,33]. The double-crossover recombinant of was screened onLB plates containing sucrose (10%, w/v) and chloramphenicol (34 μg/ml), and the mutant strain CNP-8ΔhnpAwas confirmed by PCR analysis.hnpA amplified by PCR was digested by KpnI/EcoRI and then ligatedinto the KpnI/EcoRI-digested pRK415 (Table 1), yielding plasmid pRK-hnpA. It was transformed into WM3064, and then introduced into themutant strain by biparental mating, yielding complementary strainCNP-8ΔhnpA[pRK-hnpA]. The ability of mutant and complementarystrains to utilize 2,6-DBNP was determined by monitoring the substratedegradation and the cell growth.

2.8. Analytical methods

The concentrations of 2,6-DBNP was determined by HPLC analysiswith mobile consisting of buffer A (H2O with 0.1% acetic acid) andbuffer B (methanol) [34]. A linear gradient was set from 10% to 90% Bfrom 0 to 15min, then hold at 90% B for another 5min. The flow ratewas 1.0 ml min−1, and 2,6-DBNP were detected at 320 nm. Theacetylated products of 2,6-DBNP were identified by GC–MS analysis asdescribed [31]. The nitrite concentration was determined by colori-metric method as described previously [35]. Bromide ion was de-termined on ion chromatography (Dionex ICS-3000) equipped with anIon Pac As14 column (4×250, Dionex), with the eluent of NaHCO3

(8mM) and Na2CO3 (3.5mM) at a flow rate of 1.0ml min−1 [36].

Table 1Bacterial strains and plasmids used in this study.

Strain or plasmid Relevant genotype or characteristic(s) Reference or source

Cupriavidus sp.CNP-8 2,6-DBNP utilizer, wild type This studyCNP-8ΔhnpA CNP-8 mutant with hnpA gene deleted This studyCNP-8ΔhnpA[pRK-hnpA] mnpA gene was complemented by pRK-hnpA in CNP-8ΔhnpA This study

E. coli strainsDH5α supE44 lacU169 (φ80lacZΔM15) recA1 endA1 hsdR17 thi-1 gyrA96 relA1 TransGen BiotechBL21(DE3) F−ompT hsdS(rB−mB-) gal dcm, lacY1(DE3) TransGen BiotechWM3064 Donor strain for conjugation, 2,6-diaminopimelic acidauxotroph: thrB1004 pro thirpsLhsdS lacZΔM15 RP4-1360 Δ(araBAD)567

ΔdapA1341::[ermpir(wt)]Lab stock

PlasmidspET-28a(+) Expression vector, KanR NovagenpK18mobsacB Gene replacement vector derived from plasmid pK18; Mob+ sacB+ Kmr Lab stockpRK415 Broad host range vector, TcR Lab stockpXMJ19 Source of chloramphenicol resistance gene (cm) Lab stockpET-hnpA NdeI-XhoI fragment containing hnpA inserted into pET-28a(+) This studypET-hnpB NdeI-XhoI fragment containing hnpB inserted into pET-28a(+) This studypET-hnpC NdeI-XhoI fragment containing hnpC inserted into pET-28a(+) This studypK18-hnpA hnpA gene knockout vector containing DNA fragments homologous to the upstream and downstream regions of the hnpA and cm This studypRK-hnpA hnpA gene complementation vector by cloning hnpA into the KpnI/EcoRI restriction site of pRK415 This study

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2.9. Accession number

The transcriptome data have been deposited in the Sequence ReadArchive database of NCBI with accession numbers SRP150659,SRP150660, SRP150665 (three biological replicates of uninducedcells), SRP150666, SRP150667 and SRP150670 (three biological re-plicates of 2,6-DBNP-induced cells). The nucleotide sequence has beendeposited in NCBI under GenBank accession number MH271067.

3. Results and discussion

3.1. Degradation of 2,6-DBNP by strain CNP-8

As an emerging aromatic halogenated DBP, 2,6-DBNP has been re-cently identified in various water environment, especially in China[2–4,7]. However, very little is known about the biodegradation pro-cess of 2,6-DBNP to data. The time course assay of 2,6-DBNP de-gradation, together with the growth of CNP-8 and accumulation of ni-trite and bromide ions is shown in Fig. 1A. After a delay period ofapproximately 10 h, strain CNP-8 rapidly degraded 0.3mM 2,6-DBNPto undetectable levels within 25 h, and the OD600 increased from 0.051to 0.103. 0.57mM of bromide was released when 2,6-DBNP was de-graded completely, indicating that CNP-8 could remove both brominesfrom 2,6-DBNP. The non-stoichiometric release of nitrite during 2,6-DBNP degradation is owing to that CNP-8 utilized it as the source ofnitrogen. No degradation of 2,6-DBNP was observed in a negativecontrol without inoculation of CNP-8 cells. These results revealed thatstrain CNP-8 can degrade 2,6-DBNP, and utilize it as sole carbon, ni-trogen and energy sources for growth. In addition, this strain was also

able to degrade 2,6-dichloro-4-nitrophenol, the chlorinated analogue of2,6-DBNP, and utilize it for cell growth (data not shown). Bio-transformation showed that the 2,6-DBNP-induced CNP-8 (OD600 = 2)degrade 0.3 mM 2,6-DBNP within 30 min, while no apparent de-gradation of 2,6-DBNP was observed by the un-induced cells during thisperiod (Fig. 1B), indicating that the enzymes responsible for 2,6-DBNPdegradation were inducible. To our knowledge, CNP-8 is the first strainreported for 2,6-DBNP utilization. The studies on 2,6-DBNP degrada-tion by this strain will greatly improve our understanding of the fate of2,6-DBNP in the environment.

3.2. Effect of additional carbon on 2,6-DBNP biodegradation

Three simple carbon sources including lactate, succinate and glu-cose (0.5 and 5 g/L, respectively) was added into MM in the presence of0.3 mM 2,6-DBNP to determine their effect on 2,6-DBNP degradation bystrain CNP-8 (Table S2). All supplemented carbon sources at con-centrations of either 0.5 or 5 g/L supported the growth of CNP-8 todifferent extent. For substrate degradation, addition of lactate, succi-nate and glucose at concentration of 0.5 g/L was found to accelerate2,6-DBNP degradation, and succinate supplement resulted in higherdegradation rate than other to carbon sources. However, the degrada-tion rate of 2,6-DBNP decreased significantly with addition of 5 g/Lcarbon sources, although the biomass yield had a remarkable increase.This suggested that addition of easily degraded substrate at low con-centration was beneficial for the degradation of 2,6-DBNP by CNP-8,but the excessive addition would cause the delayed degradation of 2,6-DBNP. Similar phenomena have also been reported in the microbialdegradation of some other aromatic compunds [25,37]. Therefore, it isour view that addition of low concentration of simple carbon sources isan advisable strategy when CNP-8 was used for bioremediation of 2,6-DBNP-contaminated environments.

3.3. Kinetic studies of 2,6-DBNP degradation

Because strain CNP-8 is the first microorganism degrading 2,6-DBNP, the kinetic parameters for 2,6-DBNP biodegradation was un-available in the literature. However, this knowledge is very valuable inevaluating the capacity of microorganisms in biodegradation process[16]. In this study, degradation of different concentrations of 2,6-DBNPwas carried out to study the effect of initial substrate concentration onCNP-8 growth. This strain degraded 2,6-DBNP completely when theinitial concentration was no more than 0.7mM, and the delay periodsof both 2,6-DBNP degradation and CNP-8 growth extended with theincrease of 2,6-DBNP concentration (Fig. 2). Neither 2,6-DBNP de-gradation nor CNP-8 growth was observed when initial 2,6-DBNP was1mM (data not shown). These indicated that high concentration 2,6-DBNP could inhibite the growth of strain CNP-8. Indeed, aromaticcompounds exhibiting growth inhibition against their utilizers at highconcentration was inevitable [16,25,37–39]. Subsequently, Haldane’sgrowth kinetics model, widely used to depict growth kinetics of in-hibitory compounds [37,39,40], was used to determine the kineticparameters of 2,6-DBNP degradation. Experimental and predictedspecific growth rates of CNP-8 according to Haldane’s model wereshown in Fig. 3. The maximal specific growth rate (μmax) was 0.096 h−1

when 2,6-DBNP concentration was about 0.11mM. Particularly, thespecific growth rate increased with an increase of initial 2,6-DBNP to acertain concentration, and then decreased on further increasing 2,6-DBNP concentration, similar with the microbial degradation studies ofphenol [25], 4-chlorophenol [39] and 2,4,6-trinitrophenol [40]. Thehalf saturation constant (Ks) and inhibition constant (Ki) for 2,6-DBNPdegradation by CNP-8 was 0.05mM and 0.31mM, respectively. Bycomparing the kinetic parameters of 2,6-DBNP with those of a muchsimpler pollutant 2C4NP [26], CNP-8 had lower substrate affinity anddegradation rate for 2,6-DBNP, and higher inhibition by 2,6-DBNP ascompared to 2C4NP. This is likely due to that 2,6-DBNP is higher toxic

Fig. 1. Degradation of 2,6-DBNP in MM by strain CNP-8 (A), 2,6-DBNP-inducedand uninduced strain CNP-8 (B) under aerobic conditions. The initial OD600 ofthe inoculated cells for panels A and B were 0.05 and 2, respectively. The CNP-8growth and nitrite and bromide ions release were also measured for panel A.

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and more resistant to microorganism than 2C4NP.The biomass yield coefficient was estimated according to Eq. (3).

The yield coefficient changed from 0.05 to 0.19mgmg−1 when theinitial concentration of 2,6-DBNP varied from 0.05 to 0.7 mM, andseemed to be too low. It is probably attributed to that a larger portion ofthe molecular weight of 2,6-DBNP was provided by two bromine atoms.The value of yield coefficient changed slightly at low 2,6-DBNP con-centrations varying from 0.05 to 0.2mM. Beyond 0.2mM, the yieldcoefficient decreased with the increase of 2,6-DBNP up to 0.7 mM. This

is likely due to that more energy coming from 2,6-DBNP degradationwas needed to overcome the effect of 2,6-DBNP inhibition. Similarly,decrease of yield coefficient with increase of initial substrate con-centration in the inhibitory range have also been reported during themicrobial degradation of some other aromatics [16,25]. Overall, al-though substrate inhibition was inevitable, Ks was extremely low ascompared to Ki during 2,6-DBNP degradation, indicating that CNP-8was capable to degrade 2,6-DBNP from the kinetic viewpoint.

3.4. Prediction of 2,6-DBNP catabolic genes by comparative transcriptomeanalysis and RT-qPCR

Above biotransformation assay showed that the enzymes involvedin the catabolism of 2,6-DBNP were inducible; therefore, comparativetranscriptome analysis was initially carried out to find the potentialgenes responsible for 2,6-DBNP degradation. The results revealed thatthe transcription levels of 25 genes were increased over 30-fold in 2,6-DBNP-induced CNP-8 as compared to the uninduced cells (Table S3).Generally, the genes responsible for catabolism of nitroaromatic com-pounds are clustered in the genome. Therefore, based on the annota-tions of the 25 up-regulated genes, a gene cluster containing GM005195to GM005200 (59 to 648-fold up-regulated) was predicted to be re-sponsible for 2,6-DBNP catabolism. The cluster was named as hnpcluster, as annotated and outlined in Fig. 4A. Among the enzymes en-coded by these hnp genes, HnpA, HnpB and HnpC showed high identityto the FADH2-dependent 2,4,6-trichlorophenol (2,4,6-TCP) mono-oxygenase (TcpA), putative flavin reductase (TcpB) and 6-chlorohy-droxyquinol 1,2-dioxygenase (TcpC), respectively [41]. HnpD ishomologous to the maleylacetate reductases (PnpF), which was re-ported to convert 2-chloromaleylacetate to β-ketoadipate via mal-eylacetate [34]. HnpR, a LysR regulatory protein, was proposed tocontrols hnp gene expression as it is homologous to the transcriptionalregulator TcpR in 2,4,6-TCP degradation [42]. To verify the tran-scriptome results, we analyzed the mRNA levels of four hnp genes byRT-qPCR. The transcription levels of hnpA, hnpB, hnpC and hnpD in 2,6-DBNP-induced cells increased 254-, 160-, 143- and 37-fold, respectivelyas compared to the uninduced cells (Fig. 4B), similar to the tran-scriptome data.

3.5. Expression and purification of HnpA, HnpB and HnpC

To further confirm whether hnp genes are responsible for 2,6-DBNPcatabolism, the functions of the enzymes encoded by hnpA, hnpB andhnpC were validated experimentally. HnpA, HnpB and HnpC were ex-pressed as N-terminal His6-tagged fusion proteins in E. coli BL21(DE3).All three proteins were substantially soluble in E. coli grown at 16 °Cand easy for purification. Approximately 10.8mg H6-HnpA, 12.5mgH6-HnpB and 24.4mg H6-HnpC were, respectively, purified from 1 L ofculture medium. SDS-PAGE analysis showed that the molecular massesof the denatured H6-HnpA, H6-HnpB, and H6-HnpC were approximately59, 22, and 31 kDa (Fig. 5), respectively, which were consistent withthe deduced molecular mass from their amino acid sequence.

3.6. HnpB catalyzes the reduction of FAD

The flavin reductase activity of HnpB was assayed by monitoring thedisappearance of the NAD(P)H (λmax= 340 nm) in the presence ofFAD. NADH (λmax= 340 nm) was rapidly consumed by purified H6-HnpB with the production of NAD+ (λmax= 260 nm) (Fig. S1A). NoNADH consumption occurred when HnpB was omitted from the reac-tion system (Fig. S1B). HnpB exhibited the maximum FAD-reducingactivity (5.4 U mg−1) in the presence of NADH. Only 30% relativeactivity was observed when NADH was replaced by NADPH, indicatingthat NADH was preferred to NADPH as the co-substrate of HnpB. TheKm values of HnpB for FAD and NADH were 5.4 ± 1.1 and28.7 ± 4.7 μM, respectively.

Fig. 2. Biodegradation of 2,6-DBNP (A) and growth of CNP-8 (B) at differentinitial 2,6-DBNP concentrations. The initial OD600 values of the inoculated cellswas approximately 0.05.

Fig. 3. Experimental and predicted specific growth rates of strain CNP-8 due toHaldane's model.

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3.7. HnpA catalyzes the sequential denitration and debromination of 2,6-DBNP to 6-bromohydroxyquinol in the presence of HnpB

Purified H6-HnpA alone did not degrade 2,6-DBNP in the presenceof NADH and FAD. However, rapid degradation of 2,6-DBNP(λmax= 400 nm) occurred with the addition of H6-HnpB, together withconsumption of NADH (λmax= 340 nm) (Fig. 6A). In addition, theformation of an isobestic point at 280 nm indicated that 2,6-DBNP wastransformed to a new product (λmax≈250 nm). These results indicatedthat FADH2 provided by HnpB is necessary for the activity of HnpA,similar with the reports of the FADH2-dependent monooxygenase(TcpA) involved in catabolism of 2,4,6-TCP [43]. The maximal specificactivity of HnpA against 2,6-DBNP was 0.27 ± 0.05 U mg−1. Enzymekinetic assays showed that HnpA had a Km of 5.3 ± 1.2 μM and a Kcat/Km ratio of 0.09 ± 0.02 μM-1 min-1 for 2,6-DBNP.

E. coli BL21(DE3) carrying pET-hnpA alone was found to degrade2,6-DBNP, although there is no HnpB (Fig. S2). This is likely due to thatHnpA is able to use free FADH2 coming from reduction of FAD by theindigenous NADH-FAD oxidoreductase in E. coli [44], similar with thereports for several other halogenated aromatics monooxygenases[31,43]. Notably, there is no significant degradation of 2,6-DBNP byBL21(DE3)-pET-hnpA under anaerobic conditions, indicating that O2 is

essential for the catalytic activity of HnpA against 2,6-DBNP (Fig. S2).For product identification, ascorbic acid (1mM) was used to prevent

the autooxidation of products from 2,6-DBNP catalyzed by HnpA. Twocompounds with GC retention times of 17.7 and 22.1min, respectively,were detected during the GC–MS identification of the acetylated pro-ducts (Fig. 7A). The mass spectrum of compound I (17.7 min) exhibiteda molecular ion peak at m/z 352, together with fragments at m/z 310(loss of −COCH3) and m/z 268 (loss of two −COCH3) (Fig. 7B), in linewith the mass spectrum property of acetylated 2,6-dibromohy-droquinone (2,6-DBHQ) [45]. In addition, peaks at m/z 265.84 [M-2],267.92 [M], and 269.86 [M+2] as well as their relative intensities (1:2: 1) are characteristics of a compound containing two bromine atoms.The mass spectrum of compound II (22.1 min) was typical for a com-pound containing one bromine and proposed as acetylated 6-bromo-hydroxyquinol (6-BHQ) with molecular ion peak at m/z 330, togetherwith its fragments at m/z 288 (loss of −COCH3), m/z 246 (loss of two−COCH3) and m/z 204 (loss of three −COCH3) (Fig. 7C).

The identification of 2,6-DBHQ and 6-BHQ suggested that HnpAcatalyzed the sequential denitration and debromination of 2,6-DBNP inthe presence of HnpB. Generally, monooxygenases attacked the aro-matic ring substituted by an electron-withdrawing group (such as nitroor halogen group) producing a quinone compound [46]. Therefore, theidentification of 2,6-DBHQ proposed that the direct product of 2,6-DBNP catalyzed by HnpA was 2,6-dibromo-1, 4-benzoquinone (2,6-DBBQ). The production of 2,6-DBHQ during 2,6-DBNP conversion maybe attributed to the non-enzymatic reduction of 2,6-DBBQ by reductant(such as ascorbic acid or NADH in this study), and similar explanationhas been also proposed previously [43]. Moreover, the FADH2-depen-dent monooxygenases were reported to catalyze their substrates twicein tandem [31,43]. Therefore, the detection of 6-BHQ indicated that2,6-DBBQ from the first step could be further hydrolyzed to 6-BHQ via6-bromohydroxyquinone. These combined data suggested that HnpAtransforms 2,6-DBNP to 6-BHQ via two different reactions: (i) oxidizing2,6-DBNP to 2,6-DBBQ, and (ii) hydrolyzing 2,6-DBBQ to 6-BHQ(Fig. 4C), similar to the catalytic mechanism of some other FADH2-dependent monooxygenases [31,43].

3.8. HnpC and HnpD are likely involved in the lower pathway of 2,6-DBNPcatabolism

In this study, the direct catalysis of HnpC against 6-BHQ was notperformed since the commercial 6-BHQ is unavailable. Therefore, itsstructure analogue hydroxyquinol (HQ) was used to identify the ac-tivity of HnpC. The enzymatic reaction system contained 1mM ascorbicacid which could prevent the autooxidation of HQ. As shown in Fig. 6B,

Fig. 4. Organization of the hnp gene cluster (A), transcriptional analyses of the hnp genes under induced and uninduced conditions by RT-qPCR (B), and the proposedpathway of 2,6-DBNP degradation by strain CNP-8 (C).

Fig. 5. SDS-PAGE analysis of purified Hnp proteins. Lane M: molecular massstandards; lane 1: purified His6-HnpA; lane 2: His6-HnpB; lane 3: His6-HnpC.

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Fig. 6. Spectral changes during the transformation of 2,6-DBNP by purified HnpAB (A) and the transformation of hydroxyquinol by purified HnpC (B).

Fig. 7. Identification of the acetylated products of 2,6-DBNP by GC–MS. (A) The gas chromatogram is the extracted ion current chromatogram at m/z 268 ± 0.50and 204 ± 0.50 from the total ion current chromatogram. (B) The mass spectra of acetylated 2,6-DBHQ. (C) The mass spectra of acetylated 6-BHQ.

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H6-HnpC catalyzed the consumption of hydroxyquinol(λmax= 289 nm), together the formation of a product with a λmax of243 nm, in line with the spectral property of maleylacetate [47]. Noapparent degradation of HQ was observed for the negative control (noHnpC). Considering that HnpC has high identity with the 6-CHlorohy-droxyquinol (6-CHQ) 1,2-dioxygenase (TcpC), which was reported totransform 6-CHQ to 2-CHloromaleylacetate (2-CMA) [41]. Moreover,hnpC is clustered with hnpA and highly transcribed in 2,6-DBNP-in-duced CNP-8. These suggested that HnpC is responsible for ring-clea-vage of 6-BHQ to 2-bromomaleylacetate (2-BMA) during 2,6-DBNPdegradation (Fig. 4C). The activity of HnpD (maleylacetate reductase)was not characterized in this study. However, previous studies haveshown that maleylacetate reductases could catalyze the dechlorinationreaction of 2-CMA to generate β-ketoadipate via MA [34,48]. Moreover,hnpD is located downstream of hnpC and up-regulated in 2,6-DBNP-induced CNP-8. Therefore, HnpD was proposed to be responsible fordebromination of 2-BMA to β-ketoadipate via MA during 2,6-DBNP(Fig. 4C).

3.9. hnpA is essential for strain CNP-8 to utilize 2,6-DBNP

To verify the physiological role of HnpA in 2,6-DBNP catabolism invivo, an hnpA deletion mutant strain CNP-8ΔhnpA was constructed byhomologous recombination. Strain CNP-8ΔhnpA lost the ability to growon 2,6-DBNP, and hnpA gene complementation of the mutant CNP-8ΔhnpA (CNP-8ΔhnpA[pRK-hnpA], introduction of the plasmid pRK-hnpA) recovered the ability to utilize 2,6-DBNP (Fig. 8). Thus, hnpA1was necessary for CNP-8 to utilize 2,6-DBNP for growth.

4. Conclusions

Cupriavidus sp. strain CNP-8 was the first microorganism capable toutilize 2,6-DBNP. It could degrade 2,6-DBNP in concentrations as highas 0.7mM, with μmax= 0.096 h−1, Ks = 0.05mM and Ki = 0.31mM.The hnp gene cluster responsible for 2,6-DBNP degradation was iden-tified, and three enzymes were purified and functionally verified. HnpAcould catalyze the sequential denitration and debromination of 2,6-DBNP to 6-BHQ in the presence of HnpB. HnpC likely catalyzed thering-cleavage of 6-BHQ during 2,6-DBNP catabolism. hnpA was foundto be essential for CNP-8 to utiluze 2,6-DBNP by gene knockout andcomplementation. This report fills a gap in the understanding of themicrobial 2,6-DBNP degradation.

Conflict of interests

The authors declare no competing financial interest.

Acknowledgements

This work was supported by the National Natural ScienceFoundation of China (No. 31600085), the State’s Key Project ofResearch and Development Plan (2016YFC1402300), the Foreword KeyPriority Research Program of Chinese Academy of Sciences (QYZDB-SSW-DQC013), the State Key Laboratory of Microbial Metabolism,Shanghai JiaoTong University (MMLKF17-04), and the Yantai Scienceand Technology Project (2017ZH092).

Appendix A. Supplementary data

Supplementary material related to this article can be found, in theonline version, at doi: https://doi.org/10.1016/j.jhazmat.2018.08.063.

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