Concurrent Haloalkanoate Degradation and Chlorate Reduction by Pseudomonas chloritidismutans AW-1^T

ABSTRACT

Haloalkanoates are environmental pollutants that can be degraded aerobically by microorganisms producing hydrolytic dehalogenases. However, there is a lack of information about the anaerobic degradation of haloalkanoates. Genome analysis of Pseudomonas chloritidismutans AW-1^T, a facultative anaerobic chlorate-reducing bacterium, showed the presence of two putative haloacid dehalogenase genes, the l-DEX gene and dehI, encoding an l-2-haloacid dehalogenase (l-DEX) and a halocarboxylic acid dehydrogenase (DehI), respectively. Hence, we studied the concurrent degradation of haloalkanoates and chlorate as a yet-unexplored trait of strain AW-1^T. The deduced amino acid sequences of l-DEX and DehI revealed 33 to 37% and 26 to 86% identities with biochemically/structurally characterized l-DEX and the d- and dl-2-haloacid dehalogenase enzymes, respectively. Physiological experiments confirmed that strain AW-1^T can grow on chloroacetate, bromoacetate, and both l- and d-α-halogenated propionates with chlorate as an electron acceptor. Interestingly, growth and haloalkanoate degradation were generally faster with chlorate as an electron acceptor than with oxygen as an electron acceptor. In line with this, analyses of l-DEX and DehI dehalogenase activities using cell-free extract (CFE) of strain AW-1^T grown on dl-2-chloropropionate under chlorate-reducing conditions showed up to 3.5-fold higher dehalogenase activity than the CFE obtained from AW-1^T cells grown on dl-2-chloropropionate under aerobic conditions. Reverse transcription-quantitative PCR showed that the l-DEX gene was expressed constitutively independently of the electron donor (haloalkanoates or acetate) or acceptor (chlorate or oxygen), whereas the expression of dehI was induced by haloalkanoates. Concurrent degradation of organic and inorganic halogenated compounds by strain AW-1^T represents a unique metabolic capacity in a single bacterium, providing a new piece of the puzzle of the microbial halogen cycle.

IMPORTANCE Halogenated organic and inorganic compounds are important environmental pollutants that have carcinogenic and genotoxic effects on both animals and humans. Previous research studied the degradation of organic and inorganic halogenated compounds separately but not concurrently. This study shows concurrent degradation of halogenated alkanoates and chlorate as an electron donor and acceptor, respectively, coupled to growth in a single bacterium, Pseudomonas chloritidismutans AW-1^T. Hence, besides biogenesis of molecular oxygen from chlorate reduction enabling a distinctive placement of strain AW-1^T between aerobic and anaerobic microorganisms, we can now add another unique metabolic potential of this bacterium to the roster. The degradation of different halogenated compounds under anoxic conditions by a single bacterium is also of interest for the natural halogen cycle in different aquatic and terrestrial ecosystems where ample natural production of halogenated compounds has been documented.

INTRODUCTION

Haloalkanoates are widely used as intermediates and raw materials for production of pesticides, pharmaceuticals, and other organic compounds (1). Each year, large amounts of these compounds are introduced into the environment, causing serious concerns due to their environmental toxicity and their carcinogenic and genotoxic effects on animals and humans (2). Microbial degradation plays an important role in detoxification and mineralization of haloalkanoates. Dehalogenation is often one of the first reactions during the degradation process, through which the halogen substituents, usually responsible for toxicity of these compounds, are removed (3). Bacterial strains capable of using haloalkanoates as the sole source of carbon and energy have been isolated and characterized from different genera, including Pseudomonas (4–8), Xanthobacter (9), and Methylobacterium (10).

Enzymes involved in dehalogenation of haloalkanoates are known as haloacid dehalogenases, which catalyze the hydrolytic dehalogenation of haloalkanoates and produce the corresponding hydroxyl alkanoates. Bacterial 2-haloacid dehalogenases that specifically act on α-substituted haloalkanoates are classified into three groups based on their substrate and stereochemical specificities. l-2-Haloacid dehalogenase (l-DEX) catalyzes the dehalogenation of l-2-haloalkanoates, d-2-haloacid dehalogenase (d-DEX) acts on d-2-haloalkanoates, and dl-2-haloacid dehalogenase (dl-DEX) acts on both enantiomers (11). For example, 2-haloacid dehalogenases catalyze the dehalogenation of d- or l-2-chloropropionate (d- or l-2CP, respectively) to l- or d-lactate, respectively, which is channeled to the tricarboxylic acid (TCA) cycle by further degradation to pyruvate and acetyl-coenzyme A (acetyl-CoA). The known haloalkanoate-dehalogenating bacteria degrade d- and l-2CP with molecular oxygen as a terminal electron acceptor. To our knowledge, no other terminal electron acceptors, such as chlorate, nitrate, Fe(III), or sulfate, have been reported to be used for bacterial growth on haloalkanoates; however, oxidation of l-2CP as a model compound coupled to reduction of these electron acceptors is thermodynamically feasible, with chlorate being the most favorable electron acceptor (Table 1). Oxidation of haloalkanoates coupled to chlorate reduction is of particular interest due to the concurrent removal of these two environmentally problematic compounds that potentially cooccur in environments as herbicides (12, 13) or as disinfection by-products (14). Chlorate-reducing bacteria generally reduce chlorate first to chlorite by chlorate reductase (encoded by the clr gene), and chlorite is then split into chloride and oxygen by chlorite dismutase (encoded by cld) (15–17). The molecular oxygen released from chlorite dismutation can be utilized as a terminal electron acceptor for the final mineralization of haloalkanoates.

TABLE 1.

Stoichiometric equations and standard Gibbs free energy changes for l-2-chloropropionate oxidation coupled to reduction of various electron acceptors

Electron acceptor (ox/red)a	Stoichiometric equation	ΔG°′ (kJ/mol)b
O2/H2O	C3H4O2Cl− + 3O2 + H2O → 3HCO3− + Cl− + 3H+	−1,284
ClO3−/Cl−	C3H4O2Cl− + 2ClO3− + H2O → 3HCO3− + 3Cl− + 3H+	−1,533
NO3−/N2	C3H4O2Cl− + 2.4NO3− → 3HCO3− + 1.2N2 + Cl− + 0.6H+ + 0.2H2O	−1,309
Fe3+/Fe2+	C3H4O2Cl− + 36Fe(OH)3(s) → 3HCO3− + 12Fe3O4(s) + Cl− + 53H2O + 3H+	−1,207
SO42−/H2S	C3H4O2Cl− + 1.5SO42− + H2O → 3HCO3− + 1.5HS− + Cl− + 1.5H+	−149
CO2/CH4	C3H4O2Cl− + 2.5H2O → 1.5HCO3− + 1.5CH4 + Cl− + 1.5H+	−124

^aox, oxidation; red, reduction.

^bStandard Gibbs free energy formation of the inorganic compounds were taken from Oelkers et al. (50) and http://www2.ucdsb.on.ca/tiss/stretton/database/inorganic_thermo.htm. Standard Gibbs free energy formation of 2-chloropropionate was taken from Dolfing and Janssen (51).

In this study, Pseudomonas chloritidismutans AW-1^T was selected as a potential degrader of haloalkanoates, coupled to chlorate reduction. This bacterium was previously isolated from an anoxic bioreactor (16) and is able to degrade a wide variety of electron donors, including n-alkanes, with chlorate as an electron acceptor (18, 19). Genome analysis of strain AW-1^T showed the presence of two putative haloacid dehalogenase genes, i.e., the l-DEX gene and dehI, predicted to encode l-DEX and halocarboxylic acid dehydrogenase (DehI), respectively. Hence, growth on haloalkanoates with chlorate as an alternative electron acceptor might represent a unique metabolic capacity in this bacterium. To test this hypothesis, different haloalkanoates were tested as electron donors and carbon sources with either chlorate or oxygen as an electron acceptor. The functionalities of the two putative 2-haloacid dehalogenases were determined by gene expression studies using reverse transcription-quantitative PCR (RT-qPCR) and in vitro dehalogenase activity measurements.

RESULTS AND DISCUSSION

Bioinformatic analysis.

The genome of strain AW-1^T (19) (GenBank accession no. AOFQ01000000) harbors two haloacid dehalogenase genes (the l-DEX gene and dehI) predicted to encode l-DEX and DehI, with 228 and 301 amino acid residues, respectively (Fig. 1). The amino acid sequence of l-DEX of strain AW-1^T shares 33%, 34%, 34%, and 37% identities with the l-DEX of Pseudomonas putida 109 (20), Pseudomonas putida AJ1 (7), Pseudomonas sp. strain YL (21), and Xanthobacter autotrophicus GJ10 (22), respectively. The amino acid sequence of DehI of strain AW-1^T shares 86%, 29%, 28%, and 26% identities with the d-DEX of Pseudomonas putida AJ1 (23), dl-DEX of Pseudomonas putida PP3 (24), dl-DEX of Pseudomonas sp. strain 113 (25), and dl-DEX of Methylobacterium sp. strain CPA1 (10), respectively.

FIG 1.

Multiple-sequence alignments of l-DEX (A) and d-DEX, dl-DEX, and DehI (B). White letters on a black background indicate amino acids that are identical in all sequences. Active-site residues are indicated with triangles. The d- and l-form halide-binding residues are indicated with squares and circles, respectively. The catalytic residues are indicated with stars. The source bacterial abbreviations are as follows: AW-1, Pseudomonas chloritidismutans AW-1^T; 109, Pseudomonas putida 109; AJ1, Pseudomonas putida AJ1; YL, Pseudomonas sp. YL; GJ10, Xanthobacter autotrophicus GJ10; PP3, Pseudomonas putida PP3; 113, Pseudomonas sp. 113; CPA1, Methylobacterium sp. CPA1. GenBank accession numbers are indicated at the C-terminal end. ClustalW multiple-sequence alignment was conducted using BioEdit version 7.2.5 (http://www.mbio.ncsu.edu/BioEdit/page2.html).

The proposed substrate binding and catalytic residues of the active site of the structurally characterized l-DEX of Pseudomonas sp. YL (26, 27) are identical in the l-DEX of strain AW-1^T (Fig. 1A), indicating dehalogenation of l-2-halopropionates and haloacetates by this enzyme. In contrast, only the catalytic residues of the active site of the structurally characterized dl-DEX of Pseudomonas putida PP3 (28) are identical in the DehI of strain AW-1^T and d-DEX of Pseudomonas putida AJ1 (23) (Fig. 1B). The halide-binding residues for l- and d-form halopropionates are identical only in the dl-DEX of Pseudomonas sp. 113 (25) and Methylobacterium sp. CPA1 (10) but not in the DehI of strain AW-1^T or d-DEX of Pseudomonas putida AJ1 (23). Moreover, the key residue for dictating stereoselectivity, Ala207, in dl-DEX (28) is replaced by Asn in d-DEX and DehI (Fig. 1B). These indicate that the DehI of strain AW-1^T is a d-DEX and mediates the dehalogenation of d-2-halopropionates and haloacetates. The DehI and l-DEX of strain AW-1^T share no sequence identity with each other. This is in agreement with previous studies showing that d-DEX (and dl-DEX) and l-DEX are evolutionarily unrelated and have different reaction mechanisms (29, 30).

Degradation of haloalkanoates by strain AW-1^T with either chlorate or oxygen as an electron acceptor.

Strain AW-1^T can utilize dl-2CP, l-2CP, d-2CP, l-2-bromopropionate (l-2BP), d-2-bromopropionate (d-2BP), chloroacetate, and bromoacetate as sole carbon and energy sources with chlorate or oxygen as an electron acceptor (Fig. 2, 3, and S1 in the supplemental material). Under chlorate-reducing conditions, the fastest degradation of haloalkanoates by strain AW-1^T was observed with l-2CP (Fig. 2A), dl-2CP (Fig. S1A), l-2BP (Fig. 3A), and d-2BP (Fig. 3B), with specific growth rates of 0.17, 0.12, 0.081, and 0.10 h⁻¹, respectively. Chloroacetate (Fig. 2E) and bromoacetate (Fig. 3C) were less favorable substrates, resulting in specific growth rates of 0.047 and 0.052 h⁻¹, respectively. d-2CP was the least favorable substrate, with the lowest specific growth rate (0.025 h⁻¹) among all substrates tested in this study (Fig. 2C). The chemical instability of d(l)-2BP in aqueous solution that could be spontaneously hydrolyzed to l(d)-lactate (11) might facilitate the dehalogenation of d-2BP to l-lactate and contribute to the higher specific growth rate of the strain AW-1^T with d-2BP (Fig. 3B) than with d-2CP (Fig. 2C). However, the uninoculated control experiment did not show any decrease in concentration for d- and l-2BP within 36 h, indicating a lack of abiotic d- and l-2BP dehalogenation (data not shown). Oxygen concentration in the cultures of strain AW-1^T grown on chlorate with either dl-2CP or chloroacetate did not surpass 0.009 mM dissolved oxygen (Fig. S2), indicating that oxygen produced from chlorate reduction was continuously consumed for mineralization of the haloalkanoates by strain AW-1^T. Interestingly, degradation of some haloalkanoates was faster with chlorate as an electron acceptor than with oxygen. For example, the specific growth rates of dl-2CP (Fig. S1B), l-2CP (Fig. S1D), and chloroacetate (Fig. S1H) by strain AW-1^T under aerobic conditions were 6.5-, 5.8-, and 3.9-fold lower, respectively, than the corresponding specific growth rates of these substrates under chlorate-reducing conditions. No growth was observed using β-substituted haloalkanoates, such as 3-chloropropionate, 3-bromopropionate, 3-iodopropionate, or 4-chlorobutyrate, nor with 2,3-dichloropropionate or 2-chlorobutyrate as the substrates with chlorate as an electron acceptor (data not shown). Therefore, the degradation of these substrates with oxygen as an electron acceptor was not tested in this study. Compared to the common degradation of α-substituted haloalkanoates, the degradation of β-substituted haloalkanoates was reported less frequently, and the responsible dehalogenase genes and enzymes have not been verified experimentally (1, 31, 32).

FIG 2.

Growth of P. chloritidismutans AW-1^T on l-2CP (A), d-2CP (C), chloroacetate (E), and acetate (G) with chlorate, and on acetate (I) with oxygen as electron acceptor, and relative expression of the l-2-haloacid dehalogenase gene (l-DEX gene), halocarboxylic acid dehydrogenase gene (dehI), and chlorate dismutase gene (cld) during growth on l-2CP (B), d-2CP (D), chloroacetate (F), and acetate (H) with chlorate, and on acetate with oxygen (J) as an electron acceptor. Error bars in the left panels indicate the standard deviation based on measurements from two random cultures out of 10 replicates for each growth condition sacrificed at each sampling point for growth, HPLC, and RT-qPCR analyses. Error bars in the right panels indicate standard deviation of triplicate qPCRs performed on samples withdrawn from two random replicate microcosms (n = 2 × 3).

FIG 3.

Growth of P. chloritidismutans AW-1^T on l-2BP (A), d-2BP (B), and bromoacetate (C) with chlorate as electron acceptor. Points and error bars represent the average and standard deviation of samples taken from duplicate cultures.

Dehalogenase activity assays.

The dehalogenase activity was determined in cell-free extracts (CFEs) of strain AW-1^T. The optimal pH for dehalogenase activity of the CFE from AW-1^T cells grown on dl-2CP and chlorate at 30°C for 24 h was 10.5 (Fig. 4). The optimal growth temperature of 30°C (16) was selected for further dehalogenase activity assays. Although higher dehalogenase activities were observed at higher temperatures, spontaneous release of bromide was detected in dehalogenase activity assays with d- and l-2BP as the substrates. This also confirmed the chemical instability of d(l)-2BP in aqueous solution, which might lead to the faster apparent degradation of d-2BP than d-2CP by strain AW-1^T. The CFEs prepared from both chlorate- and oxygen-grown cultures of strain AW-1^T showed dehalogenase activities with all the growth-supporting haloalkanoates tested in this study (Table 2). In addition, dehalogenase activity was also noted with 2-chlorobutyrate while it was not used as the growth substrate. No activity was observed with 4-chlorobutyrate, 3-chloropropionate, 3-bromopropionate, 3-iodopropionate, or 2,3-dichloropropionate (Table 2). The dehalogenase activity of the CFE obtained from AW-1^T cells grown in the presence of chlorate was up to 3.5-fold higher than the CFE obtained from AW-1^T cells grown in the presence of oxygen (Table 2). This is in line with the growth experiments that showed faster growth when chlorate was used as an electron acceptor than that in aerobic cultures (Fig. S1). Chlorite dismutase is a periplasmic enzyme (19, 33–35) and hence, utilization of the molecular oxygen derived from chlorite dismutation by oxygenases involved in the further oxidation of the dehalogenated haloalkanoates could be more efficient than using the oxygen from the extracellular environment. To this end, it should be noted that the solubility of chlorate in water (9.93 M at 25°C) is much higher than that of oxygen (0.000269 M at 25°C, under air), suggesting that exponentially growing cells of strain AW-1^T might be oxygen diffusion limited in the case of aerobic cultivation. Finally, thermodynamic analysis shows that chlorate is a more favorable electron acceptor than oxygen for complete oxidation of l-2CP (Table 1).

FIG 4.

Effect of pH (A) and temperature (B) on dehalogenase activity of the CFEs prepared from P. chloritidismutans AW-1^T cells grown on dl-2CP and chlorate at 30°C for 24 h. The pH (A) and temperature (B) yielding the highest dehalogenase activity were set as 100%, and activities are shown as percentages of the highest activity. The points are averages of two technical replicates, and the error bars represent the standard deviations.

TABLE 2.

Dehalogenase activity of the CFEs of P. chloritidismutans AW-1^T on various haloalkanoate substrates

Substrate	Dehalogenase activity (U/mg of protein)a
	dl-2CP + chlorateb	dl-2CP + oxygenc
l-2-Chloropropionate	1.58 ± 0.19	0.46 ± 0.05
d-2-Chloropropionate	0.09 ± 0.021	0.11 ± 0.35
dl-2-Chloropropionate	1.50 ± 0.04	0.59 ± 0.01
l-2-Bromopropionate	1.54 ± 0.02	0.89 ± 0.26
d-2-Bromopropionate	1.48 ± 0.26	0.42 ± 0.10
Chloroacetate	1.43 ± 0.09	1.33 ± 0.01
Bromoacetate	2.10 ± 0.03	1.71 ± 0.26
2-Chlorobutyrate	0.39 ± 0.13	0.09 ± 0.03
4-Chlorobutyrate	ND	ND
3-Chloropropionate	ND	ND
3-Bromopropionate	ND	ND
3-Iodopropionate	ND	ND
2,3-Dichloropropionate	ND	ND

^aValues of dehalogenase activity are the mean ± standard deviation of technical duplicate analysis. ND, not detected.

^bCFE was prepared from cells grown on dl-2CP and chlorate for 24 h.

^cCFE was prepared from cells grown on dl-2CP and oxygen for 90 h.

Transcription analysis.

Under all tested conditions, the time-zero expression of cld, the l-DEX gene, and dehI was comparable for all of the cultures, and the 16S rRNA gene was stably expressed throughout the growth phases of strain AW-1^T (Fig. S3). Among the three analyzed genes, dehI showed the highest induction under chlorate-reducing conditions with l-2CP, d-2CP, and chloroacetate as electron donors, which was significant in early and mid-exponential-growth phases (Fig. 2B, D, and F). Upregulation of dehI reached its highest level (∼14,000-fold) in l-2CP-fed cultures within 24 h and then decreased (Fig. 2B). In contrast, the expression of the l-DEX gene was relatively stable, and the highest upregulation (∼22-fold) was observed in the cultures amended with l-2CP after 18 h, which then decreased (Fig. 2B, D, and F). Similar to the l-DEX gene, cld also showed no significant upregulation in the cultures amended with the chloroalkanoates and chlorate (Fig. 2B, D, and F). In the cultures grown on nonchlorinated substrate (acetate) with either chlorate or oxygen, upregulations of dehI, the l-DEX gene, and cld did not surpass 18-, 26-, and 49-fold, respectively (Fig. 2H and J). These results collectively show the inductive expression of dehI by haloalkanoates and high constitutive expression of the l-DEX gene and cld independently of the electron donor and acceptor utilized (Fig. S2). In line with the expression pattern of cld, a previous proteomic study showed an abundance of chlorite dismutase in strain AW-1^T even when chlorate was replaced by oxygen (19).

Previous research on the degradation of organic and inorganic halogenated compounds has mainly focused on their degradation either as an electron donor or electron acceptor but not on concurrent degradation. This study showed concurrent degradation of halogenated compounds as an electron donor and acceptor in a single bacterium, representing a unique and untapped metabolic potential. A survey of available bacterial genomes showed a similar cooccurrence of genes involved in degradation of haloalkanoates and chlorate in other bacterial strains belonging to various genera, including, but not limited to, Bacillus, Exiguobacterium, Mycobacterium, Staphylococcus, and Roseiflexus (Table S1). However, none of these bacteria were experimentally tested for chlorate reduction and (or) haloalkanoate degradation, and thus further experimental verification is needed. This suggests that the potential catabolic machineries to degrade both halogenated organic and inorganic compounds by a single bacterium are widespread. Besides bioremediation prospects, such degradation of different halogenated compounds is of interest for the natural halogen cycle in different aquatic and terrestrial ecosystems where ample natural production of halogenated compounds has been documented (36–39).

MATERIALS AND METHODS

Chemicals.

Chloroacetate, bromoacetic acid, 2-chloropropionic acid, l-2-chloropropionic acid, l-2-bromopropionic acid, d-2-chloropropionic acid, d-2-bromopropionic acid, 3-chloropropionic acid, 3-bromopropionic acid, 3-iodopropionic acid, 2,3-dichloropropionic acid, 2-chlorobutyric acid, and 4-chlorobutyric acid were all purchased from Sigma-Aldrich. All inorganic salts used in this study were analytical grade.

Bacterial strain and growth conditions.

P. chloritidismutans AW-1^T was cultivated in 120-ml bottles containing 50 ml of anoxic medium, as previously described (16), with nitrogen or air (140 kPa) as the headspace and incubated statically in the dark at 30°C. Vitamins and trace elements were added as described by Holliger et al. (40), except that the trace elements were supplemented with the following (per liter of trace elements solution): 0.06 g of Na₂SeO₃ and 0.0184 g of NaWO₄·2H₂O. To obtain a preculture, 10 mM acetate and 10 mM chlorate were used as the electron donor and acceptor, respectively. When all acetate was consumed and the optical density at 600 nm (OD₆₀₀) reached ∼0.5, the preculture was transferred (5% [vol/vol]) into fresh medium with different haloalkanoates as the electron donor instead of acetate and either chlorate or oxygen as the electron acceptor. Haloalkanoic acids were neutralized with an equimolar amount of NaOH to produce the corresponding haloalkanoates and were filter-sterilized through a 0.2-μm-pore-size filter (Advanced Microdevices, Ambala, India) before being added to the medium at 3 to 10 mM final concentration. For transcription analysis, degradation of d-2CP, l-2CP, and chloroacetate with chlorate, degradation of acetate with chlorate, and degradation of acetate with oxygen were tested. To ensure sufficient biomass for transcription analysis, 10 replicate microcosms were prepared for each condition, and for each sampling occasion, two microcosms were randomly selected and sacrificed for RNA extraction after taking samples for high-performance liquid chromatography (HPLC) analysis of metabolites and OD₆₀₀ measurements. The specific growth rate was calculated according to the equation ln[OD_600(t2)/OD_600(t1)] = k(t₂ − t₁), where k is the specific growth rate; OD_600(t1) and OD_600(t2) are the optical densities of liquid cultures measured at 600 nm at the start and end of exponential-growth phases, respectively; and t₁ and t₂ are the starting point and endpoint (in hours) of exponential-growth phases, respectively.

RNA extraction and cDNA synthesis.

RNA was extracted from strain AW-1^T at different time points during growth on l-2CP (0, 12, 18, 24, and 36 h), d-2CP (0, 48, 96, 144, and 168 h), chloroacetate (0, 24, 30, 36, and 48 h), acetate with chlorate (0, 4.5, 9, 14, and 24 h), and acetate with oxygen (0, 9, 24, 39, and 48 h). RNA extraction was performed with a bead-beating procedure, as described earlier (41). RNA was purified using RNeasy columns (Qiagen, Venlo, The Netherlands) with DNase I (Roche, Almere, The Netherlands) treatment, according to the manufacturer's protocols. cDNA was synthesized from 500 ng of total RNA using the Maxima H Minus first-strand cDNA synthesis kit (Thermo Scientific, Vilnius, Lithuania), according to the manufacturer's protocols. The absence of genomic DNA was confirmed by 16S rRNA gene-targeted PCR with extracted RNA samples as the template.

qPCR assays.

Primers for amplification of cld, the l-DEX gene, and dehI were designed using the Primer3 online program (http://primer3.ut.ee/) or the NCBI online primer design tool (Primer-BLAST [http://www.ncbi.nlm.nih.gov/tools/primer-blast/]) (Table 3). Primers were tested in silico using OligoAnalyzer 3.1 (Integrated DNA Technologies). cld, the l-DEX gene, and dehI were PCR amplified using the following program: 95°C for 3 min, followed by 30 cycles of 95°C for 30 s, 55°C for 30 s, and 72°C for 30 s, followed by a final extension at 72°C for 10 min. The PCR products were purified using the GeneJET PCR purification kit (Thermo Scientific, Vilnius, Lithuania) and cloned into pGEM-T Easy vector (Promega, WI, USA). The plasmid was introduced into Escherichia coli JM109 competent cells (Promega). Primer specificity and efficiency of amplification were tested by temperature-gradient PCRs on the iQ5 iCycler (Bio-Rad, Veenendaal, The Netherlands) using plasmid or PCR product amplified with T7/SP6 primers from the plasmid containing target gene inserts. The same T7/SP6 PCR products were subsequently used to obtain qPCR calibration curves. qPCRs were performed using the iQ SYBR green supermix (Bio-Rad, CA, USA), as described earlier (42). The program for qPCR assays of cld, the l-DEX gene, and dehI was 95°C for 10 min, followed by 40 cycles of 95°C for 15 s, 60°C for 30 s, and 72°C for 30 s. Melting curves were measured from 65°C to 95°C with increments of 0.5°C and 10 s at each step. Transcript levels of cld, the l-DEX gene, and dehI were calculated by relative quantification using the 2^−ΔΔCq method (43). The 16S rRNA gene was used as the reference gene (44) and quantified as described previously (42). Gene expression over time was calibrated to the 0-h time point (44). A relative expression higher than 10 was arbitrarily set as representing significant induction (45).

TABLE 3.

Overview of qPCR primers used in this study

Gene name	Primer name	Primer sequence (5′ to 3′)
cld	CldF	ACACGACACCTACCTTAGCC
	CldR	CCCCAACGAACGTGGAATTT
l-DEX gene	l-DEXF	CTTTATCGGCGTGGTGAGTG
	l-DEXR	CCCACGGATCGAATAATGCC
dehI	DehIF	CTACCGGCCTTTCTTTGTCG
	DehIR	CTGATCAATCTCACGCACCG

Preparation of CFE and dehalogenase assay.

CFEs were prepared from 50-ml cultures of strain AW-1^T at early stationary phase grown with dl-2CP under either chlorate-reducing or aerobic conditions. Cells were harvested by centrifugation at 4,700 × g for 15 min at 4°C. The cell pellets were washed twice with 100 mM Tris-sulfate buffer (pH 7.5) and resuspended in 1 ml of the same buffer supplied with 10% glycerol. Cells were lysed by sonication using a Branson sonifier (Branson, CT, USA) equipped with a 3-mm tip by six pulses of 30 s with a 30-s rest in between pulses. Intact cells and cell debris were removed by centrifugation at 15,000 × g and 4°C for 15 min. The protein concentration of the supernatant was determined with the Qubit protein assay kit (Invitrogen, OR, USA), according to the manufacturer's instructions. Dehalogenase activity of the freshly prepared CFEs was measured by determining the release of halide ions under aerobic conditions without chlorate. The optimum pH and temperature for the dehalogenase activity were determined using two buffer types with distinct yet overlapping pH ranges (100 mM Tris-sulfate, pH 7.5, 8.0, 8.5, and 9.0; 100 mM glycine-NaOH, pH 9.0, 9.5, 10.0, 10.5, 11.0, 11.5, and 12.0) and different temperatures (20, 25, 30, 35, and 40°C). A control reaction mixture lacking CFE was included in each set of assays to detect any spontaneous release of halide ions. The dehalogenase assay system contained 400 μl of the buffer solutions, 20 mM haloalkanoates, and 50 μl of CFE. All reaction mixture components except the CFE were combined and allowed to equilibrate for 5 min at a given temperature, after which the reaction was initiated by adding 50 μl of CFE. The reaction was performed under aerobic conditions and terminated after 10 min by adding 75 μl of 2 N H₂SO₄. The release of halide ions was measured by ion chromatography. One unit of dehalogenase activity was defined as the amount of protein that catalyzes the dehalogenation of 1 μmol substrate per minute of reaction time.

Analytical methods.

Chlorate and halide ions were analyzed using the Thermo Fisher Scientific Dionex ICS-2100 ion chromatography system and a Dionex IonPac analytical column (AS19, 2 by 250 mm) equipped with a suppressed conductivity detector. The ions were analyzed under a three-step gradient profile consisting of 10 mM KOH for 4 min, 10 to 40 mM KOH for 16 min, and then 40 to 10 mM KOH for 1.5 min. Haloalkanoates were analyzed on a Thermo Fisher Scientific SpectraSYSTEM HPLC equipped with an Agilent column (MetaCarb 67H, 300 by 6.5 mm) and a refractive index (RI) detector. The mobile phase was with 0.01 N H₂SO₄. Oxygen was measured by taking 0.5-ml-headspace samples and was analyzed using a gas chromatograph equipped with thermal conductivity detector (GC-TCD; Shimadzu 2014) and a Restek column (MolSieve 13×, 200 by 3 mm). The column temperature was 60°C and held for 2.75 min. Cell growth was determined by measuring the OD₆₀₀ using a WPA CO8000 cell density meter (Biochrom, Cambridge, UK).

Genome annotation.

Bacterial genomes with a high-quality genome sequence available in the European Nucleotide Archive (ENA) version 121 were scanned for cooccurrence of l-DEX and chlorite dismutase (Cld) using protein domains IPR006439, IPR006328, IPR023214, and IPR010644. The DehI in the AW-1^T genome was found using the conserved regions of d- and dl-DEX from Pseudomonas putida PP3 (24), Pseudomonas sp. 113 (25), Methylobacterium sp. CPA1 (10), and Pseudomonas putida AJ1 (23). To avoid potential misannotations, the selected genomes were de novo reannotated using the SAPP framework (46, 47). Genes were identified using Prodigal (2.6.3) (48), and protein annotation was performed though protein domains using InterProScan (5.19-58.0) (49).