Characterization of 1,2-dibromoethane-degrading haloalkane dehalogenase from Bradyrhizobium japonicum USDA110

Enzyme and Microbial Technology 45 (2009) 397–404

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Characterization of 1,2-dibromoethane-degrading haloalkane dehalogenase from Bradyrhizobium japonicum USDA110 Christos C. Sfetsas a , Leonidas Milios a , Katholiki Skopelitou a , Anastasia Venieraki b , Rodanthi Todou a , Emmanouil Flemetakis b , Panagiotis Katinakis b , Nikolaos E. Labrou a,∗ a b

Laboratory of Enzyme Technology, Department of Agricultural Biotechnology, Agricultural University of Athens, Iera Odos 75, 11855 Athens, Greece Laboratory of Molecular Biology, Department of Agricultural Biotechnology, Agricultural University of Athens, Iera Odos 75, 11855 Athens, Greece

a r t i c l e

i n f o

Article history: Received 29 April 2009 Received in revised form 29 June 2009 Accepted 18 July 2009 Keywords: Bioremediation 1,2-Dibromoethane Haloalkane dehalogenase Hydrolase Rhizobial

a b s t r a c t Haloalkane dehalogenases (DHAs, E.C. 3.8.1.5) are very promising biocatalytic tools for the bioremediation of environmental pollutants which consists of haloalkanes. In the present work, we investigated the DHA from Bradyrhizobium japonicum USDA110 (BjDHA). The dehalogenase activity of B. japonicum USDA110 and RT-PCR analysis revealed that the BjDHA gene expression is induced by 1,2-dibromoethane (1,2-DBE) during the early exponential phase. The BjDHA gene was cloned, expressed in Escherichia coli BL21 (DE3) and characterized. The enzyme catalyzes the irreversible hydrolysis of a variety of haloalkanes to the corresponding alcohol, halide, and a hydrogen ion. The catalytic properties of the recombinant enzyme were investigated and the kinetic parameters (Km , kcat ) for a number of substrates were determined. The results showed that the BjDHA displays wide substrate specificity towards haloalkanes and particular high activity towards 1,2-DBE. The enzyme has a different catalytic triad topology compared to the Xanthobacter haloalkane dehalogenase and is more similar to the Rhodococcus enzyme. In addition, consistent with its broad specificity, the BjDHA has a substantially larger and more polar active site cavity compared to the Xanthobacter and Rhodococcus enzymes and as a consequence, BjDHA is able to dehalogenate longer and polar compounds. These properties make this enzyme very promising bioremediation tool for environmental applications. © 2009 Elsevier Inc. All rights reserved.

1. Introduction 1,2-Dibromoethane (1,2-DBE) is an extremely toxic chemical owing to its cancer-causing potential [1]. 1,2-DBE was used primarily as an antiknock additive to gasoline and is one of the most effective and widely used pesticidal soil fumigants [2]. Many years after its last known application as a soil fumigant, residual 1,2DBE is still found at remarkably high concentrations in soil because it strongly interacts with the soil matrix [3]. 1,2-DBE can slowly leach from such contaminated soils to groundwater over exceedingly long periods, and because of its slow chemical conversion in aqueous milieu, it is a continuous source of contamination of water supplies [3].

Abbreviations: BCH, bromocyclohexane; 1,10-BD, 1,10-bromodecanoate; BjDHA, Bradyrhizobium japonicum USDA110 dehalogenase; DHA, dehalogenase; 1,2-DBE, 1,2-dibromoethane; 1,2-DBP, 1,2-dibromopropane; IPTG, isopropyl-␤d-thiogalactopyranoside; PAGE, polyacryamide gel electrophoresis; SDS, sodium dodecyl sulfate; TOPO, topoisomerase. ∗ Corresponding author. Tel.: +30 210 5294308; fax: +30 210 5294308. E-mail address: [email protected] (N.E. Labrou). 0141-0229/$ – see front matter © 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.enzmictec.2009.07.013

Microbial dehalogenases (DHAs) consist a class of bacterial enzymes which catalyze the hydrolytic dehalogenation of haloalkanes to the corresponding alcohols, halide ions and protons, with water as the sole cosubstrate [4–11]. DHAs are monomeric enzymes of approximately 300 amino acids and a molecular mass of 30–35 kDa. Three experimental determined structures of haloalkane dehalogenases [12,13] are available and represented by the enzymes DhlA from Xanthobacter autotrophicus GJ10 [5,14], DhaA from Rhodococcus rhodochrous [15] and LinB from Sphingomonas paucimobilis UT26 [16]. They belong to the ␣/␤ hydrolase fold superfamily [17]. The shape of the molecule is spherical and is composed of two domains [18]: domain I which has an ␣/␤ type structure and includes the catalytic triad and domain II, which is also known as cap domain. The cap domain composed of a few helices inserted into the catalytic domain, usually at the C-terminal to ␤-strand 6. Cap domain has been found to contribute to substrate specificity. The major difference between different DHAs is the rate limiting step of the enzymatic reaction. The cloning, expression and partial characterization of DHAs of two rhizobial strains (Mesorhizobium loti and Bradyrhizobium japonicum) have recently reported [9]. From the biotechnology point of view DHA attracts attention because of many potential uses for the bioremediation of soil,

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water and air [19,20] from a wide range of environmental pollutants which consists fully or partially of haloalkanes. Recently, the genes of haloalkane dehalogenase and haloacid dehalogenase from the bacterium X. autotrophicus GJ10 were introduced into tobacco (Nicotiana tabacum ‘Xanthi’) plants. Transgenic tobacco plants were able to remediate a range of halogenated, aliphatic hydrocarbons [21]. In the present work we investigate the biochemical and catalytic properties of BjDHA using kinetic analysis, molecular modelling and expression studies.

cells were pelleted by centrifugation and genomic DNA was isolated according to a standard procedure [23]. The PCR reaction was carried out in a total volume of 50 ␮l contained: 6.8 pmole of each primer, 50 ng template genomic DNA, 10 mM dNTPs, 5 ␮l 10× Taq buffer and 4 U of Taq DNA polymerase (Stratagene, USA). The PCR procedure comprised 30 cycles of 2 min at 95 ◦ C, 1 min at 39 ◦ C and 2.5 min at 72 ◦ C. A final extension time at 72 ◦ C for 10 min was performed after the 30th cycle. The resulting PCR amplicon was TOPO (topoisomerase) ligated into a T7 expression vector (pCR® T7/CT-TOPO® ). The resulting expression construct pT7BjDHA was sequenced along both strands and were used to transform competent Escherichia coli BL21 (DE3) cells.

2. Materials

E. coli BL21 (DE3) ells, harboring plasmid pT7BjDHA-6His were grown at 37 ◦ C in 1 L LB (Bacto-tryptone, 1%, w/v; yeast extract, 0.5, w/v; NaCl, 1%, w/v) medium containing 100 ␮g/mL ampicillin. The synthesis of BjDHA was induced by the addition of 1 mM IPTG when the absorbance at 600 nm was approximately 0.6. Three to four hours after induction, cells were harvested (approx. 3 g) by centrifugation at 10,000 × g for 10 min (4 ◦ C), resuspended in potassium phosphate buffer (50 mM, pH 8.0, 9 mL) containing sodium chloride (0.3 M), sonicated, and centrifuged at 10,000 × g for 20 min. A sample of the supernatant (1.5 mL, 5.46 mg total protein) was loaded to a column of Ni-NTA-Sepharose (0.5 mL), which was previously equilibrated with potassium phosphate buffer (50 mM, pH 8.0) containing sodium chloride (0.3 M). Non-adsorbed protein was washed off with 10 mL equilibration buffer, followed by 10 mL of potassium phosphate buffer (50 mM, pH 6.2), containing sodium chloride (0.3 M). Bound BjDHA was eluted with equilibration buffer containing various concentrations of imidazole (2 mL, 5, 20, 100, 200, 500 mM). Collected fractions (2 mL) were assayed for BjDHA activity and protein (Bradford assay [22]). Protein purity was judged by SDS-PAGE. Purified enzyme fractions, before use, were dialyzed against HEPES/NaOH buffer (1 mM, pH 8.2, containing 20 mM Na2 SO4 , 1 mM EDTA) and stored at 4 ◦ C.

1,2-C2 H4 Br2 and 1,2-C3 H6 Br2 , perchloric acid, mercury thiocyanate, iron nitrate and phenol red and other haloalkanes were obtained from Sigma–Aldrich (USA). Isopropyl-␤-d-thiogalactopyranoside (IPTG), rifampicin, ampicillin and agarose were purchased from Genaxis. HEPES and other salts and buffers were from Merck (Germany). Yeast extract, peptone and agar were obtained from Scharlau (Spain). The expression kit (pCR® T7/CT-TOPO® ) was purchased from Invitrogen (USA). The DyNAzyme II DNA polymerase from Finnzymes (Finland) was used for the PCR reactions. 3. Methods 3.1. Bacterial growth conditions B. japonicum strain USDA110 was grown at 28 ◦ C on YMB or TY supplemented or not with the appropriate concentration of 1,2-DBE (0.2, 2.0, 5.0 mM). The YMB medium contained the following constituents: K2 HPO4 , 0.5 g/L; Mg2 SO4 ·7H2 O, 0.2 g/L; NaCl, 0.1 g/L; yeast extract, 1 g/L; mannitol, 10 g/L. The TY medium contained: 5 g/L tryptone; 3 g/L yeast extract; 10 mM CaCl2 . For the construction of the growth curves, 25 mL starter cultures were used to inoculate 200 mL of YMB to an initial OD600 of 0.05. 3.2. Determination of transcript levels using RT-PCR assay B. japonicum USDA110 cells were grown as described above in the presence (0.2, 2 mM) or in the absence of 1,2-DBE. At selected time intervals, cells were harvested by centrifugation at 10,000 × g for 10 min at 4 ◦ C and washed twice with 0.5 mM NaCl. Total RNA was isolated using the Trizol reagent (Invitrogen, Paisley, UK), according to the manufacture’s protocols, and quantified by spectrophotometry and agarose gel electrophoresis. Prior to RT-PCR, the total RNA samples were treated with DNase I (Promega, Madison, WI) at 37 ◦ C for 60 min in order to eliminate any traces of contaminating genomic DNA. First-strand cDNA was reverse transcribed using SuperScript II (Invitrogen) and random hexanucleotides (Invitrogen) from 2 ␮g of DNase I treated total RNA. BjDHA transcripts were specifically amplified using BjDHA-F 5 -TTT CGT ACG CGG ATT AGC CTT-3 and BjDHA-R 5 -CGA CCT CGG TAT GGT GGA AAT-3 primers. The different RNA preparations were normalized by the amplification of the constitutively expressed gene BjDNAK, coding for the heat shock protein 70 chaperone, using BjDNAK-F 5 -CAC GAT CGC CGT GTA CGA T-3 and BjDNAK-R 5 -GCC GAT TTC GAG AAT GGA GAT A-3 primers. Under our experimental conditions, an exponential increase of the amplification products was observed until after 25 amplification cycles (94 ◦ C/1 min, 54 ◦ C/1 min, 72 ◦ C/1 min), so all the amplification reactions were performed under these conditions in order to obtain semi-quantitative results. The RT-PCR products were analyzed by 1.5% (w/v) agarose gel electrophoresis, blotted and hybridized with the respective digoxigenin-11-rUTP labelled probe. 3.3. Time course of haloalkane dehalogenase activity in B. japonicum USDA110 B. japonicum USDA110 cells were grown as described above in the presence (2.0 mM) or in the absence of 1,2-DBE. At selected time intervals, cells were harvested by centrifugation at 10,000 × g for 10 min at 4 ◦ C, resuspended in HEPES/NaOH buffer (1 mM pH 8.2, containing 20 mM Na2 SO4 , 1 mM EDTA), sonicated, and centrifuged at 10,000 × g for 20 min. The supernatant was collected and was assayed for dehalogenase activity using 1,2-DBE as substrate and protein using the method of Bradford [22]. 3.4. Cloning and PCR amplification A BLASTp search of B. japonicum USDA110 sequences was carried out using the sequence of haloalkane dehalogenase DhaA from R. rhodochrous (NCBI accession number Q53042) as probe to identify DHA protein and gene sequence. PCR was used to amplify the full-length ORF from genomic DNA using the oligo-primers synthesized to the 5 region of the genes from the ATG start codon and to the 3 end of the gene. The primers sequences were as follows: 5 -ATG AGC AAG CCA ATC GAG ATC G-3 and 5 -CGC GGC GAG CTG CGG ACG CAC CG-3 . B. japonicum USDA110 was grown at 30 ◦ C in YMB medium containing 20 ␮g/mL rifampicin. After 3 days,

3.5. Heterologous expression and purification of BjDHA

3.6. Bioinformatics analysis and molecular modelling Sequences homologous to BjDHA were sought in the NCBI using BLAST [24]. The resulting sequence set was aligned with Clustal W [25]. ESPript [26] (carried out at http://prodes.toulouse.inra.fr/ESPript/cgi-bin/ESPript.cgi) was used for alignment visualization and manipulation. The molecular model of BjDHA was constructed using MODELLER 6 [27] (carried out at http://www.infobiosud.cnrs.fr/bioserver). The determined X-ray crystal structure of R. rhodochrous (PDB code 1cqwA [15]) determined at 1.5 Å resolution with which the BjDHA shares 51% sequence identity, was used as a template. An iterated protocol involving multiple model construction and rigorous protein structure quality assessment, using PROSA II [28] (carried out at http://www.lmcp.jussieu.fr/sincris-top/logiciel/prg-prosa.html), and Verify 3D [29] (carried out at http://nihserver.mbi.ucla.edu/Verify 3D) were used. The overall scores were used to choose the final model. For inspection of models and crystal structures we used the program PyMOL (DeLano Scientific). 3.7. Assay of enzyme activity and kinetic analysis Dehalogenase activities were measured at 22 ◦ C using two methods. In the first method, the reaction was monitored by determining the halide release as described by Iwasaki et al. [30]. In brief, 100 ␮L of crude enzyme extract or purified enzyme preparation (>0.01 U) was incubated for 20 min with 4.9 mL HEPES/NaOH buffer (1 mM, pH 8.2), containing 20 mM Na2 SO4 , 1 mM EDTA and 10 mM of 1,2-DBE. Samples (200 ␮L) were taken every 5–10 min, mixed with 1.8 mL distilled H2 O, 400 ␮L mercury thiocyanate (2 mM) and 160 ␮L (20 mM) ferric nitrate. The amount of halide release was determined by a standard curve obtained at 460 nm. Initial velocities were determined from the linear range of the curves obtained from a graph of [chloride released] versus time (min). One unit of dehalogenase activity is defined as the amount of enzyme that liberates 1 ␮mol of halide ion per minute at 22 ◦ C. Alternatively, qualitative enzyme assays were carried out using the dye indicator colorimetric method according to Holloway et al. [31], as following: crude extract or purified enzyme preparation (20–100 ␮L) was mixed with 900–980 ␮L of HEPES/NaOH buffer (1 mM pH 8.2, containing 20 mM Na2 SO4 , 1 mM EDTA, 10 mM haloalkane and 20 ␮g/mL of the dye indicator phenol red). The enzymatic reaction was measured at 22 ◦ C by determining the rate of proton formation which caused a pH decrease with concomitant change of dye indicator color at 540 nm. Steady-state kinetic measurements were performed in HEPES/NaOH buffer (1 mM, pH 8.2, containing 20 mM Na2 SO4 and 1 mM EDTA) by varying the concentration of the substrate (haloalkane, 0.1–10 mM). The kinetic parameters kcat and Km were calculated by non-linear regression analysis of experimental steady-state data. Turnover numbers were calculated on the basis of one active site per 34.1 kDa subunit. Kinetic data were analyzed using the computer program GraFit [32]. 3.8. Protein determination Protein concentration was determined at 25 ◦ C by the method of Bradford [22] using bovine serum albumin (fraction V) as standard.

C.C. Sfetsas et al. / Enzyme and Microbial Technology 45 (2009) 397–404 3.9. pH dependence on enzyme activity For the study of the pH dependence of BjDHA activity the following buffers were used in order to cover a pH range from 4.5 to 9.0: sodium acetate (100 mM) was used for pH 5.5, potassium phosphate (100 mM) was used to cover pH 6.5–7.5 and Tris–acetate (100 mM) to cover the pH range pH 8.2–9.0. The measurements were conducted at 22 ◦ C and 1,2-DBE was used as substrate. pKa values were estimated by fitting the experimental data to the equation reported by Blanchard and Cleland [33], using the computer program GraFit [32]. 3.10. Electrophoresis SDS polyacrylamide gel electrophoresis was performed according to the method of Laemmli [34] on a vertical slab gel containing 12.5% (w/v) polyacrylamide (running gel) and 2.5% (w/v) stacking gel. The protein bands were stained with Coomassie Brilliant Blue R-250.

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4. Results 4.1. A haloalkane dehalogenase of B. japonicum USDA110 In silico homology searches of B. japonicum USDA110 sequences using the amino acid sequence of the haloalkane dehalogenase from R. rhodochrous (Q53042) as a query sequence, revealed the presence of a single sequence corresponding to putative DHA homologues (designated Bjdha), in agreement with the work of Sato et al. [9]. BjDHA contained an open reading frame (ORF) of 930 bp, coding for a polypeptide of 310 amino acid residues with a predicted molecular mass of 34088.96 Da and a theoretical pI of 5.89. Fig. 1 shows the amino acid sequence alignments resulting from the BLAST search

Fig. 1. Multiple sequence alignment of haloalkane dehalogenases. The alignments were produced using Clustal W [25]. Numbering and the secondary structure of the model of BjDHA are shown above the alignment. Alpha helices and beta strands are represented as helices and arrows, respectively, and beta turns are marked with TT. Important active site amino acid residues are indicated by asterisk (*). Residues that are involved in the formation of the substrate binding site are indicated by (+). Conserved areas are shown shaded. A column is framed, if more than 70% of its residues are similar according to physico-chemical properties. NCBI accession number for dehalogenases are in parenthesis: MlDHA: Mesorhizobium loti dehalogenase (BAB51818); BjDHA: Bradyrhizobium japonicum dehalogenase (BAC46352); RrDHA: Rhodococcus rhodochrous dehalogenase (Q53042); SpDHA: Sphingomonas paucimobilis dehalogenase (1G42-A); XaDHA: Xanthobacter autotrophicus dehalogenase (P22643); MbDHA: Mycobacterium bovis dehalogenase (AF2122/97); MtDHA: Mycobacterium tuberculosis dehalogenase (Q50642); MaDHA: Mycobacterium avium dehalogenase (CAB65289); SaDHA: Streptomyces avermitilis dehalogenase (BAC72491).

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Fig. 2. Growth the B. japonicum USDA110 in the absence (−) or in the presence (+) of 0.2, 2, and 5 mM of 1,2-DBE, respectively.

of BjDHA. Fig. 1 also shows the catalytically important amino acid residues: Asp103, Glu127 and His280 [10]. Amino acids that are involved in the formation of the substrate binding site are also indicated: Asn37, Ala101, Trp104, Gly184, Ile219, Phe176, Phe213, Pro214, Ala253, Leu254, Tyr281 [14–16]. The structural and functional role of these residues will be discussed in the next section. 4.2. The BjDHA is induced by 1,2-dibromoethane at RNA and protein level The ability of B. japonicum USDA110 to grow in the presence of halogenated compounds was tested by growth of the bacteria in YMB supplemented with increasing concentrations of 1,2-dibromoethane (1,2-DBE) (Fig. 2). This analysis demonstrated that even low concentrations of 1,2-DBE (0.2 mM) resulted in a strong reduction of growth. A 10-fold increase in the concentration of the xenobiotic resulted in a slightly longer lag phase, while at 5 mM of 1,2-DBE the growth of the B. japonicum USDA110 was strongly inhibited. The dehalogenase activity towards 1,2-DBE was assayed in total protein extracts form B. japonicum USDA110 cells collected during the growth in the presence and absence of the 1,2-DBE (Fig. 3A). Interestingly, a strong induction of dehalogenase activity in the presence of 1,2-DBE was observed during the early exponential growth phase (at 5 days), while the activity was drastically reduced during the subsequent growth phases. In order to test whether the increased dehalogenase activity could be attributed to the up-regulation of the BjDHA gene, a semi-quantitative RT-PCR approach was used. This analysis demonstrated that during the early exponential growth phase in the presence of 2 mM 1,2-DBE, BjDHA transcript levels were significantly higher in comparison with the transcript levels when growth in 1,2-DBE-free media (Fig. 3B). In agreement with the observed enzyme activity, gene expression was much lower during the stationary growth phase. Interestingly, even a 10-fold lower 1,2-DBE concentration, was able to induce a similar accumulation of BjDHA transcripts during the early exponential growth phase, although transcript levels were barely detectable during the stationary phase. 4.3. Cloning, expression and purification of BjDHA from E. coli cells PCR was used to amplify the full-length sequence from genomic DNA of B. japonicum. The resulting PCR amplicon was cloned into the T7 expression vector pCR® T7/CT-TOPO to give the pT7BjDHA6His plasmid. This plasmid was used to transform the expression host E. coli BL21 (DE3). Cell-free extract of the E. coli transformants showed high dehalogenase activity (approximately 0.5–1 U/mg

Fig. 3. (A) Time course of DHA activity in B. japonicum USDA110. B. japonicum USDA110 cells were growth as described in Section 3 in the absence (−) or in the presence (+) of 2 mM 1,2-DBE. (B) Accumulation of BjDHA transcripts in the absence (−) or in the presence of 0.2 and 2 mM 1,2-DBE, during the early exponential growth phase (5 days) and the stationary growth phase (10 days), as determined by semiquantitative RT-PCR. Differences in the starting template were normalized by the parallel amplification of the BjDNAK transcripts.

protein) towards 1,2-dibromoethane. The extra six histidine (6His) residues tagged on the C-terminus of the recombinant enzyme enable BjDHA to be rapidly purified by immobilized metal ion affinity chromatography on Ni-NTA-Sepharose affinity column (Fig. 4 and Table 1). The purity of the final BjDHA preparation was evaluated by SDS-PAGE as illustrated in Fig. 4. 4.4. Kinetic analysis and molecular modelling The substrate specificity of the purified BjDHA was investigated in order to identify catalytic activities that may be related to its biological function. To this end, a broad range of substrates was examined and the results are listed in Table 2. BjDHA is characterized by a generally broad substrate specificity and exhibits significant differences in its individual activities for a wide variety of substrates. The kinetic parameters kcat , and Km for 1,2-DBE, 1,2dibromopropane (1,2-DBP), 1,10-bromodecanoate (1,10-BD) and bromocyclohexane (BCH) were determined by steady-state kinetic analysis (Table 3). The Km values for 1,2-DBE and 1,2-DBP were determined as 0.1 ± 0.02 and 0.7 ± 0.1 mM, respectively, indicating that the enzyme exhibits higher affinity for short haloalkanes. On the other hand, the enzyme displays higher Km and lower kcat values for longer (e.g. 1,10-dibromodecanoate) or cyclic haloalkanes (e.g. bromocyclohexane). The molecular model of BjDHA was constructed, using homology modelling, to put the kinetic data in a structural context (Fig. 5). Analysis of packing, solvent exposure and stereochemical properties suggests the final BjDHA model to be of good overall quality. This consideration, combined with the good resolution of the known crystal structure of R. rhodochrous (PDB code 1cqwA [15]), allowed the results of the kinetic analysis to be placed in a detailed

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Table 1 Purification protocol of recombinant BjDHA from E. coli BL21 (DE3) crude extract. Step

Volume (mL)

Activity (U)

Protein (mg)

Sp. Act. (U/mg)

Purification (fold)

Yield (%)

Crude extract Affinity chromatography on Ni-NTA-agarose

1.5 2

3.50 3.28

5.46 0.43

0.64 8.13

1 12.7

100 93.71

Procedures were performed at 4 ◦ C as described in Sections 2 and 3.

Fig. 6. pH dependence of log Vmax for 1,2-dibromoethane hydrolysis by BjDHA. Kinetic measurements were performed as described in Section 3.

Fig. 4. SDS-PAGE of recombinant BjDHA purification. Protein bands were stained with Coomassie Brilliant Blue R-250. Lane A: E. coli BL21 (DE3) crude extract after induction with 1 mM IPTG; lane B: flow through fraction from the Ni-NTA-Sepharose affinity column; lane C: BjDHA purified by affinity chromatography on Ni-NTAagarose.

structural context and to be compared with the results from other DHAs [18]. Analysis of the model reveals that the BjDHA has a different catalytic triad topology than the Xanthobacter enzyme [5] and is more similar to the Rhodococcus enzyme [15]. The catalytic triad is formed by Asp103, Glu127 and His280 (Figs. 1 and 5B). These amino acid residues are responsible for the catalytic reaction. Based on crystallographic and site-directed mutagenesis data it has been concluded that the reaction mechanism of DHAs consists of four main steps, which in the case of BjDHA may be described as following: in the first step the substrate binds in the binding site cavity and a Michaelis complex is formed. This is followed by a nucleophilic attack of Asp103 on the halogen-bound C1 atom of

the substrate, leading to the formation of a covalent alkyl-enzyme intermediate and a halide ion, which remains bound into to active site by the halide-stabilizing residue (Asn38). The intermediate is subsequently hydrolyzed by a water molecule activated by the general base His280 (Fig. 5B). Glu127 is in hydrogen bond with His280 and this residue may assist the histidine in its function as a general base by stabilizing the positive charge on the imidazole ring that emerges as the histidine extracts a proton from the water molecule. DHAs differ both in the type and location of one halide-stabilizing residue. For example, HLD-I members employ tryptophane (Trp175 of Xanthobacter enzyme) located in helix ␣4 [35], whereas HLD-II members (e.g. BjDHA, Rhodococcus enzyme) use asparagine (Asn38 of BjDHA) located in the loop following ␤-strand 2 (␤-strand 3 in the Rhodococcus enzyme). 4.5. Effect of pH on enzyme activity The effect of pH on enzyme activity was investigated in the range between pH 4.5 and 9.0 (Fig. 6). BjDHA exhibits an optimum pH 8.2. In the area pH 7.5–8.2 and pH 8.2–8.8 the enzyme preserves 65% and 50%, respectively, of its activity and for pH values below and above these areas the activity drops sharply almost to zero.

Fig. 5. Structural representations of BjDHA model. (a) The enzyme is represented as a cartoon with ␤-strands colored yellow and ␣-helices colored red. Bound iodide ion is shown as a sphere. (b) Active site residues of BjDHA. Residues are shown in a stick representation and labeled. (c) The substrate binding site of BjDHA. Important residues are shown in a stick representation and labeled. Bound iodide ion is shown as a dot sphere. The figures were produced using PyMol (DeLano Scientific). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of the article.)

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Table 2 Substrate specificity for purified recombinant BjDHA. Enzyme assays were carried out under standard conditions as described in Section 3. Results represent the means of triplicate determinations, with variation less than 5% in all cases. Substrate

Structure

Relative activitya (%)

1,2-Dibromoethane

100

1,2-Dibromopropane

18

Bromoethylamine

48

Iodoacetamide

2.9

Epichlorohydrin (1-chloro-2,3-epoxypropane)

1.4

1,4-Dioxobutyl-dibromobutane

54

1,6-Dibromohexane

2.8

1,10-Dibromodecane

1.1

Bromocyclohexane

4.2

1-Chloro-2,4-dinitrobenzene

NDb

p-Nitrobenzyl chloride

NDb

Permethrin

NDb

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Table 2 (Continued ) Substrate

Relative activitya (%)

Structure

NDb

Atrazine

a b

Relative activity was expressed as the rate for 1,2-DBE. Non-detectable activity.

Table 3 Kinetic parameters of BjDHA. Steady-state kinetic measurements were performed at 22 ◦ C in HEPES/NaOH buffer (1 mM, pH 8.2, containing 20 mM Na2 SO4 and 1 mM EDTA). The kinetic parameters kcat and Km were calculated by non-linear regression analysis of experimental steady-state data using the GraFit (Leatherbarrow [32]) program. Substrate

Km (mM)

1,2-Dibromoethane 1,2-Dibromopropane 1,10-Dibromodecanoate Bromocyclohexane

0.1 0.72 10.5 0.7

± ± ± ±

0.02 0.1 1.4 0.1

kcat (min−1 ) 394.4 29.7 4.1 17.8

± ± ± ±

25 5 0.3 0.5

kcat /Km (mM−1 min−1 ) 3944 41.3 0.39 25.4

5. Discussion Microbes that produce dehalogenases are widely distributed in nature, apparently having evolved to degrade naturally occurring halogenated compounds in order either to exploit them as a carbon source for growth or as a means of protection against the toxicity of these compounds [36]. Fig. 1 shows the amino acid sequence alignments resulting from the BLAST search of BjDHA. BjDHA shows 57% sequence identity with the M. loti enzyme, whereas significantly lower identity (e.g. 38–51%) was observed with other bacterial dehalogenases. The lowest identity (21–23%) was observed with the dehalogenases homolog from Mycobacerium avium and X. autotrophicus. The dehalogenase family can be divided into three subfamilies denoted HLD-I, HLD-II, and HLDIII, of which HLD-I and HLD-III are predicted to be sister-groups [35]. BjDHA belong to HLD-II group. This group also includes the following experimentally characterized haloalkane dehalogenases: DmbA from M. bovis and Mycobacterium tuberculosis and DhaA from Rhodococcus rhodochrous. HLD-I is represented by the experimentally confirmed haloalkane dehalogenase, DhlA, from Xanthobacter autotrophicus and DhmA from Mycobacterium avium [37,38]. The HLD-III subfamily it is still not as well-defined and established as HLD-I and HLD-II and members are not included in the alignments presented in Fig. 1. A strong induction of dehalogenase activity in the presence of 1,2-DBE was observed during the early exponential growth phase, while the activity was drastically reduced during the subsequent growth phases. The increased dehalogenase activity is attributed to the up-regulation of the BjDHA gene as determined by semiquantitative RT-PCR (Fig. 3). This analysis demonstrated that during the early exponential growth phase in the presence of 1,2-DBE, BjDHA transcript levels were significantly higher in comparison with the transcript levels when growth in 1,2-DBE-free media (Fig. 3B). High transcript accumulation during the exponential phase has been observed for several enzymes participating in the detoxification of xenobiotics [39]. Furthermore, our data suggests that both the expression of the BjDHA gene and dehalogenase activity are induced by low concentrations of 1,2-DBE in B. japonicum

USDA110, in contrast to the respective activity in M. tuberculosis where induction by haloalkanes did not result in a higher enzyme activity in cell extracts [37]. B. japonicum DHA in agreement with other characterized DHAs exhibits wide substrate specificity. DHAs catalyze a broad range of reactions, with different members of the family exhibiting quite varied substrate specificity. BjDHA shows high activity towards 1,2-DBE, whereas haloalkane dehalogenase from X. autotrophicus GJ10 exhibits lower specific activity [40]. Also unlike other dehalogenases, BjDHA showed activity towards long and cyclic haloalkanes (e.g. 1-bromodecanoate and bromocycloalkane) [4–8]. Also polar substrates such as bromoethylamine and 1,4dioxobutyl-dibromobutane, iodoacetamide and epichlorohydrin (1-chloro-2,3-epoxypropane) are substrates for BjDHA. Epichlorohydrin is carcinogenic, mutagenic, and genotoxic halohydrin and widely used as solvent and as starting materials for resins, polymers, agrochemicals, and pharmaceuticals [41]. A range of agrochemicals, such as atrazine (herbicide), permethryn (insecticide) and the synthetic xenobiotics (1-chloro-2,4dinitrobenzene and p-nitrobenzyl chloride) were also evaluated as possible substrates for BjDHA. All these compounds did not show appreciable activity. The high activity and specificity for 1,2-DBE (Tables 2 and 3) is probably the result of several highly favourable interactions between 1,2-DBE over other haloalkanes. Analysis of the modelled structure (Fig. 5) showed that the ligand may interact with the enzyme in a conserved manner, similar to that found in the Rhodococcus enzyme active site [15,42]. Although the active site residues Asp103, Glu127 and His280 are important requirement for catalysis, the combination of other factors may also contribute and seem to be important to substrate binding and catalysis. Among these is a hydrophobic cavity consisting of conserved residues (Asn38, Trp104, Ile219, Phe176, Phe213, Pro214, Ala253, Leu254, Tyr281, Fig. 5C). The only polar residues in this cluster are a tyrosine (Tyr281) and an asparagine residue (Asn38). As predicted from sequence alignments, Ile104, Cys176 and Val245, which are present in the Rhodococcus enzyme active site, is not conserved in BjDHA and have been replaced by Ala, Gly and Ala, respectively (Ala101, Gly184, Ala253, Fig. 5C). The greater volume of the active site cavity in the BjDHA enzyme, compared to that of Rhodococcus enzyme seems to be primarily due to the presence of smaller amino acid residues (e.g. Ala and Gly) at these positions. The large size and the mixed type (hydrophobic/polar) character of the substrate binding site, predict the catalytic activity for polar and large compounds (e.g. bromoethylamine, iodoacetamide, epichlorohydrin, 1,4-dioxobutyl-dibromobutane) consisted with the results listed in Table 2. Fig. 6 shows the log Vmax –pH profile for the 1,2-DBE hydrolysis. The log Vmax –pH profile yields the pKa , which reflects the ionization of the enzyme in complex with substrate. The pH transitions

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observed are controlled by two ionizable groups with pKa s of 6.3 and 9.3. Since there are no ionizable groups in the substrate (1,2DBE), these pKa s may correspond to the enzyme important groups. The pKa determined in the acidic pH region (pKa 6.3) is close to that expected for a histidine residue [43], probably the active site His280 which play important role in hydrolysis reaction, as it has been discussed in the previous section. The other pKa at the basic region appears to be close to that expected for a Lys or a Tyr residue. Since there is no Lys residue in the binding region which may be involved in substrate binding or catalysis (Fig. 5C) it is more conceivable to assume that this pKa may reflect the ionization of Tyr281. The functional role of Tyr281 is not clear. However, one may speculate that the formation of negative charge as a consequence of Tyr281 ionization at pH > 9 may destabilize the substrate binding site, leading to the reduced ability the enzyme to forms productive Michaelis complex with the hydrophobic substrate (1,2-DBE). In conclusion, in the present report we investigated the physiological role, catalytic and structural properties of BjDHA. The enzyme displays wide substrate specificity towards haloalcanes and high activity towards 1,2-DBE. These properties make this enzyme very promising bioremediation tool. References [1] Janssen DB, van der Ploeg JR, Pries F. Genetics and biochemistry of 1,2dichloroethane degradation. Biodegradation 1994;5:249–57. [2] Alexeeff GV, Kilgore WW, Li MY. Ethylene dibromide: toxicology and risk assessment. Rev Environ Contam Toxicol 1990;112:49–122. [3] Steinberg SM, Pignatello JJ, Sawhney BL. Persistence of 1,2-dibromoethane in soils: entrapment in intraparticle micropores. Environ Sci Technol 1987;21:1201–8. [4] Janssen DB, Jager D, Witholt B. Degradation of n-haloalkanes and alpha, omegadihaloalkanes by wild-type and mutants of Acinetobacter sp. strain GJ70. Appl Environ Microbiol 1987;53:561–6. [5] Janssen DB, Gerritse J, Brackman J, Kalk C, Jager D, Witholt B. Purification and characterization of a bacterial dehalogenase with activity toward halogenated alkanes, alcohols and ethers. Eur J Biochem 1988;171:67–72. [6] Negri A, Marco E, Damborsky J, Gago F. Stepwise dissection and visualization of the catalytic mechanism of haloalkane dehalogenase LinB using molecular dynamics simulations and computer graphics. J Mol Graph Model 2007;26:643–51. [7] Hardman DJ. Biotransformation of halogenated compounds. Crit Rev Biotechnol 1991;11:1–40. [8] Damborsky J, Nyandoroh MG, Nemec M, Holoubek I, Bull AT, Hardman DJ. Some biochemical properties and the classification of a range of bacterial haloalkane dehalogenases. Biotechnol Appl Biochem 1997;26:19–25. [9] Sato Y, Monincova M, Chaloupkova R, Prokop Z, Ohtsubo Y, Minamisawa K, et al. Two rhizobial strains, Mesorhizobium loti MAFF303099 and Bradyrhizobium japonicum USDA110, encode haloalkane dehalogenases with novel structures and substrate specificities. Appl Environ Microbiol 2005;71:4372–9. [10] Kurihara T, Esaki N. Bacterial hydrolytic dehalogenases and related enzymes: occurrences, reaction mechanisms, and applications. Chem Rec 2008;8:67–74. [11] Jesenská A, Sykora J, Olzynska A, Brezovsky´ J, Zdráhal Z, Damborsky´ J, et al. Nanosecond time-dependent Stokes shift at the tunnel mouth of haloalkane dehalogenases. J Am Chem Soc 2009;131:494–501. [12] Franken SM, Rozeboom HJ, Kalk KH, Dijkstra BW. Crystal structure of haloalkane dehalogenase: an enzyme to detoxify halogenated alkanes. EMBO J 1991;10:1297–302. [13] Holmquist M. Alpha/Beta-hydrolase fold enzymes: structures, functions and mechanisms. Curr Protein Pept Sci 2000;1:209–35. [14] Silberstein M, Damborsky J, Vajda S. Exploring the binding sites of the haloalkane dehalogenase DhlA from Xanthobacter autotrophicus GJ10. Biochemistry 2007;46:9239–49. [15] Newman J, Peat TS, Richard R, Kan L, Swanson PE, Affholter JA, et al. Haloalkane dehalogenases: structure of a Rhodococcus enzyme. Biochemistry 1999;38:16105–14.

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